The Enzymes. Volume XIII: Oxidation-Reduction. Part C: Dehydrogenases (II), Oxidases (II), Hydrogen Peroxide Cleavage. Third Edition

The Enzymes VOLUME XI11 OXIDATION-REDUCTION Part C DEHYDROGENASES (II) OXIDASES (II) HYDROGEN PEROXIDE CLEAVAGE Third ...

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The Enzymes VOLUME XI11

OXIDATION-REDUCTION Part C DEHYDROGENASES (II) OXIDASES (II) HYDROGEN PEROXIDE CLEAVAGE Third Edition

CONTRIBUTORS WINSLOW S. CAUGHEY

GREGORY R. SCHONBAUM

BRITTON CHANCE

DIANA L. STIGGALL

L. ERNSTER

JOHN A. VOLPE

J. IEUAN HARRIS

YOUSSEF HATEFI

WILLIAM J. WALLACE MICHAEL WATERS

J. B. HOEK

CHARLES H. WILLIAMS, JR.

J. RYDSTROM

TAKASHI YONETANI SHINYA YOSHIKAWA

ADVISORY BOARD BRITTON CHANCE BO MALMSTROM LARS ERNSTER VINCENT MASSEY

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

Volume XI11 OXIDATIOWREDUCTION Part C DEHYDROGENASES (II) OXIDASES (II) HYDROGEN PEROXIDE CLEAVAGE

THIRD EDITION

ACADEMIC PRESS New York San Francisco London 1976 A Subeidiary of Harcourt Brace Jovanovich, Publishers

COPYRIGHT 6 1976, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN A N Y FORM OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR A N Y INFORMATION STORAGE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING FROM THE PUBLISHER.

ACADEMIC PRESS, INC.

111 Fifth Avenue, New York. New' York 10003

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

Library of Congress Cataloging in Publication Data

Main entry under title: The Enzymes. Includes bibliographical references. CONTENTS: v. 1. Structure and control.-v. 2. netics and mechanism.-v. 3. Hydrolysis: peptide bonds. (etc.] 1. Enzymes. I. Boyer,PaulD.,ed. 1. Enzymes. QU135 B791eJ [DNLM: QP601.E523 574.1 '925 75-117107 ISBN 0-12-122713-8

PRINTED IN THE UNITED STATES OF AMERICA

Ki-

Contents . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . .. . . Contents of Other Volumes . . . . . . . . . . . . .

List of Contributors

1.

vii ix

xi

Glyceraldehyde-3-phosphate Dehydrogenase

J. IEUANHARRISAND MICHAEL WATERS

.

I. Introduction . 11. Molecular Properties 111. Catalytic Properties

2.

. . . . . . . . . . . . . 1 . . . . . . . . . . . . , 3 . . . . . . . . . . . . . as

Nicotinamide Nucleotide Transhydrogenases

J. RYDSTRBM, J. B. HOEK,AND L. ERNSTER

.

.

. . .

.

. . . . . . . .

I. Definitions . . . . , . . . . . 11. BBSpecific Transhydrogenases . . . . . . . 111. ABSpecific Transhydrogenases . . . . . , . IV. Physiological Roles of Nicotinamide Nucleotide Transhydrogenases

3.

. .

51 52 62

79

Flavin-Containing Dehydrogenases

CHARLES H. WILLIAMS, JR.

.

I. Introduction . . . . . . . 11. Pyridine Nucleotide-Disulfide Oxidoreductases 111. Lipoamide Dehydrogenase . . . . IV. Glutathione Reductaae . . . . . V. Thioredoxin Reductase . . . . . VI. Microsomal Electron Transport VII. NADH-Cytochrome b. Reductase . . . VIII. NADPH-Cytochrome P-450 Reductase . .

.

.

4.

. . . . . . . . . . .

. . . . . . . .

. . . . .

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

. . . . . . .

90 92 106 129 142 148

154 185

Metal-Containing Flavoprotein Dehydrogenases

YOUSSEF HATEFIAND DIANA L. STIGGALL I. Introduction . . . 11. NADH Dehydrogenases

. . . . . . . . . . . . . . . . . . . . . . . . V

175

177

vi

CONTENTS

. .

. . . . . . . . . . . . 222 . . . . . 256 . . 280 . . . . . . . . . . . . 263 . . 273 . . 279 . . 286 . . . . . . . . . . . . . . . . 295

I11 Succinate Dehydrogenases IV . ~-Glycerol-3-phosphate Dehydrogenase (EC 1.1.995) V Choline Dehydrogenase (EC 1.1.99.1) . . . . . . . VI . Lactate Dehydrogenases VII . Nitrite Reductases (EC 1.6.6.4) . . . . . . . . . VIII . Adenylyl Sulfate Reductases (EC 1.8.99.2) . . . . . . IX . Sulfite Reductases (H&:NADPH Oxidoreductases) (EC 1.8.12) X . Addendum

5

.

Cytochrome c Oxidare

WINSLOW S. CAUGHEY. WILLIAMJ . WALLACE. JOHNA . VOLPE.AND SHINYA YOSHIKAWA

I . Introduction . . . . . I1 Isolation and Characterization I11. Chemical and Physical Properties IV Mechanisms

. .

.

6

. . . . . . . . . .

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

299

305 313

337

Cytochrome c Peroxidare

TAKASHI YONETANI I. Introduction . . . . . . . I1. Preparation and Molecular Properties I11. Structural Aspects IV . Enzymic Activity V. Reaction Mechanism VI . Interaction with Cytochrome c VII . General Comments . . . . .

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

.

7

. . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

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. . . . . . .

. . . . . . .

. . . . . .

. . . . .

. . . . .

. 363 . 366 . 369 . 388 . 409

345 347 348 352 353 356 300

Catalase

GREGORY R . SCHONBAUM AND BRITTON CHANCE I . Introduction . . . . . . I1. General Enzyme Properties . . I11. The Nature of the Active Site . IV . Catalase-Mediated Redox Reactions Author Index

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

. . . .

. . . . . . . . . . . . . . . . 435 Topical Subject Index for Volumes I-XIII . . . . . . . . . . 459

Subject Index

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

WINSLOW S. CAUGHEY (299), Department of Biochemistry, Colorado State University, Fort Collins, Colorado BRITTON CHANCE (3631, Johnson Research Foundation, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania

L. ERNSTER (51),Department of Biochemistry, Arrhenius Laboratory, University of Stockholm, Stockholm, Sweden J. IEUAN HARRIS (1), Medical Research Council Laboratory of Molecular Biology, University Postgraduate Medical School, Cambridge, England YOUSSEF HATEFI (175),Department of Biochemistry, Scripps Clinic and Research Foundation, La Jolla, California

J. B. HOEK (51),Department of Biochemistry, University of Nairobi, Nairobi, Kenya J . RYDSTROM (51),Department of Biochemistry, Arrhenius Laboratory, University of Stockholm, Stockholm, Sweden GREGORY R. SCHONBAUM (363), Department of Biochemistry, St. Jude Children’s Research Hospital, and University of Tennessee Center for the Health Sciences, Memphis, Tennessee DIANA L. STIGGALL (175), Department of Biochemistry, Scripps Clinic and Research Foundation, La Jolla, California JOHN A. VOLPE (299), Department of Biochemistry, Colorado State University, Fort Collins, Colorado WILLIAM J. WALLACE (299), Department of Biochemistry, Colorado State University, Fort Collins, Colorado MICHAEL WATERS (1),Department of Biochemistry, Monash University, Clayton, Victoria, Australia vii

viii

LIST OF CONTRIBUTORS

CHARLES H. WILLIAMS, JR. (89), Veterans Administration Hospital, and Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan TAKASHI YONETANI (346), Department of Biochemistry and Biophysics, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania SHINYA YOSHIKAWA (299), Department of Biochemistry, Colorado State University, Fort Collins, Colorado

Preface This is the final volume of the Third Edition of “The Enzymes.” It completes the coverage of oxidation-reduction enzymes. As with previous volumes, the quality and quantity of information a t the molecular level in this volume are impressive. The first portion of the volume includes the remaining chapters on the nicotinamide nucleotidelinked dehydrogenases, namely, the transhydrogenases and the very important glyceraldehyde-3-phosphate dehydrogenase. The second portion completes the treatment of the great family of flavin-containing enzymes, with chapters on the flavoprotein dehydrogenases and the metalloflavoprotein dehydrogenases. The last portion includes chapters on catalase and peroxidase that use hydrogen peroxide, and on cytochrome oxidase, the enzyme responsible for most of the oxygen consumption by animals. The Third Edition has proved considerably longer and contains much more information than was thought likely when the edition was launched. The privilege of editing the treatise has given me a deep respect for the collective accomplishments of the many scientists whose continued efforts have made such a treatise possible. The quality and abundance of information found in this edition are a tribute to the individual research worker, often little recognized, and to the society that has made such a work possible. I know of no finer recognition of man’s potentiality and creativity than has been my fortune to experience in editing this multivolume treatise. Again, it is a pleasure to acknowledge the indebtedness of the users of these volumes to the Advisory Board that helped plan each volume, to the contributors for their unusually high level of excellence, to the staff of Academic Press for their high professional standards, and to Lyda Boyer, whose editorial and other assistance made many tasks lighter. PAULD. BOYER ix

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Contents of Other Volumes Volume I: Structure and Control

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

Index xi

xii

CONTENTS OF OTHER VOLUMES

Volume II: Kinetics and Mechanism

Steady State Kinetics W . W . Cleland Rapid Reactions and Transient States Gordon B. Hammes and Paul R . Schimmel Stereospecificity of Enzymic Reactions G. Popjhk Proximity Effects and Enzyme Catalysis Thomas C . Bruice Enzymology of Proton Abstraction and Transfer Reactions Irwin A. Rose Kinetic Isotope Effects in Enzymic Reactions J . H . Richards Schiff Base Intermediates in Enzyme Catalysis Esmond E. Snell and Samuel J. DiMari Some Physical Probes of Enzyme Structure in Solution Serge N . Timasheff Metals in Enzyme Catalysis Albert S. Mildvan Author Index-Subj ect Index

Volume 111: Hydrolysis: Peptide Bonds

Carboxypeptidase A Jean A. Hartsuck and William N . Lipscomb Carboxypeptidase B J . E . Folk Leucine Aminopeptidase and Other N-Terminal Exopeptidases Robert J . DeLange and Emil L. Smith Pepsin Joseph S. Fruton

CONTENTS OF OTHER VOLUMES

Chymotrypsinogen : X-Ray Structure J . Kraut The Structure of Chymotrypsin I?.M . Blow Chymotrypsin-Chemical George P . Hess

Properties and Catalysis

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

xiii

xiv

CONTENTS OF OTHER VOLUMES

Other Bacterial, Mold, and Yeast Proteases Hiroshi Matsubara and Joseph Feder Author Index-Subject

Index

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

Ureases F. J . Reithel Penicillinase and Other p-Lactamases Nathan Citri Purine, Purine Nucleoside, Purine Nucleotide Aminohydrolases C . L. Zielke and C. H . Suelter Glutaminase and 7-Glutamyltransferases Standish C . Hartman L-Asparaginase

John C. Wriston, Jr. Enzymology of Pyrrolidone Carboxylic Acid Marian Orlowski and Alton Meister Staphylococcal Nuclease X-Ray Structure F. Albert Cotton and Edward E . Hazen, Jr. Staphylococcal Nuclease, Chemical Properties and Catalysis Christian B. Anfinsen, Pedro Cuatrecasas, and Hiroshi Taniuchi Microbial Ribonucleases with Special Reference to RNases TI, T,,N1, and Uz Tsuneko Uchida and Fuji0 Egami Bacterial Deoxyribonucleases I. R. Lehman Spleen Acid Deoxyribonuclease Giorgio Bernardi Deoxyribonuclease I M . Laskowski, Sr.

CONTENTS OF OTHER VOLUMES

Venom Exonuclease M . Laskowski, Sr. Spleen Acid Exonuclease Albert0 Bernardi and Giorgio Bernardi Nucleotide Phosphomonoesterases George I . Drummond and Masanobu Yamamoto Nucleoside Cyclic Phosphate Diesterases George I . Drummond and Masanobu Yamamoto

E. coli Alkaline Phosphatase Ted W . Reid and Irwin B. Wilson Mammalian Alkaline Phosphatases H . N . Fernley Acid Phosphatases Vincent P. Hollander Inorganic Pyrophosphatase of Escherichia coli John Josse and Simon C. K. Wong Yeast and Other Inorganic Pyrophosphatases Larry G. Butler Glucose-6-Phosphatase, Hydrolytic and Synthetic Activities Robert C. Nordlie

Fructose-1,6-Diphosphatases 8.Pontremoli and B . L. Horecker Bovine Pancreatic Ribonuclease Frederic M . Richards and Harold W . Wyckoff Author Index-Subj ect Index

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

The Hydrolysis of Sulfate Esters A. B. Roy

xv

xvi

CONTENTS OF OTHER VOLUMES

Arylsulf atases R. G. Nicholls and A. B. Roy Carboxylic Ester Hydrolases Klaus Krkch Phospholipases Donald J . Hanahan Acetylcholinesterase Harry C. Froede and Irwin B . Wilson Plant and Animal Amylases John A. Thoma, Joseph E . Spradlin, and Stephen Dygert Glycogen and Starch Debranching Enzymes E. Y . C. Lee and W . J. Whelan Bacterial and Mold Amylases Toshio Takagi, Hirolco Toda, and Toshizo Isemura Cellulases D. R. Whitaker Yeast and Neurospora Invertases J . Oliver Lampen Hy aluronidases Karl Meyer Neuraminidases Alfred Gottschallc and A. S. Bhargava Phage Lysozyme and Other Lytic Enzymes Akira Tszlgita Aconitase Jenny Pickworth Glusker p-Hydroxydecanoyl Thioester Dehydrase Konrad Bloch Dehydration in Nucleotide-Linked Deoxysugar Synthesis L. Glaser and H.Zarkowslcy

CONTENTS OF OTHER VOLUMES

xvii

Dehydrations Requiring Vitamin B,, Coenzyme Robert H. Abeles Enolase Finn Wold Fumarase and Crotonase Robert L. Hill and John W . Teipel 6-Phosphogluconic and Related Dehydrases W . A. Wood Carbonic Anhydrase S. Lindslcog, L. E. Henderson, K . K . Kannan, A. Liljas, P. 0. Nyman, and B. Strandberg Author IndexSubject Index

Volume VI: Carboxylation and Decarboxylation ( Nonoxidative), lromerization

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

Ribulose-l,5-DiphosphateCarboxylase Marvin I. Siegel, Marcia WGhnick, and M . Daniel Lane Ferredoxin-Linked Carboxylation Reactions Bob B. Buchanan Amino Acid Decarboxylases Elizabeth A. Boeker and Esmond E. Snell Actoacetate Decarboxylase Irwin Fridovich

xviii

CONTENTS OF OTHER VOLUMES

Aldose-Ketose Isomerases Ernst A . Noltmann Epimerases Luis Glaser Cis-Trans Isomerization Stanley Seltzer Phosphomutases W. J. Ray, Jr., and E. J . Peck, Jr. Amino Acid Racemases and Epimerases E lija h Adams Coenzyme Bl,-Dependent Mutases Causing Carbon Chain Rearrangements H . A . Barker Blz Coenzyme-Dependent Amino Group Migrations Thressa C . Stadtman Isopentenylpyrophosphate Isomerase P . W . Holloway Isomerization in the Visual Cycle Joram Heller A6-3-KetosteroidIsomerase Paul Talalay and Ann M . Bemon Author Index-Subject Index

Volume VII: Elimination and Addition, Aldol Cleavage and condensation, Other C C Cleavage, Phorphorolysir, Hydrolysis (Fats, Glycoriderl

Tryptophan Synthetase Charles Yanojsky and Irving P . Crawjord Pyridoxal-Linked Elimination and Replacement Reactions Leodis Davis and David E. Metzler The Enzymatic Elimination of Ammonia Kenneth R . Hanson and Evelyn A . Havir

CONTENTS OF OTHW VOLUMES

Argininosuccinases and Adenylosuccinases Sarah Ratner Epoxidases William B. Jakoby and Thorsten A. Fjellstedt Aldolases B. L. Horecker, Orestes Tsolas, and C. Y.Lai Transaldolase Orestes Tsolas and B. L. Horecker

2-Keto-3-deoxy-6-phosphogluconicand Related Aldolaseo W. A. Wood Other Deoxy Sugar Aldolases David Sidney Feingold and Patricia Ann Hoflee 8-Aminolevulinic Acid Dehydratase David Shemin 8-Aminolevulinic Acid Synthetase Peter M . Jordan and David Shemin Citrate Cleavage and Related Enzymes Leonard B. Spector Thiolase Ulrich Gehring and Feodor Lynen Acyl-CoA Ligases Malcolm J. P. Higgim, Jack A. Kornblatt, and Harry Rudney a-Glucan Phosphorylases-Chemical and Physical Basis of Catalysis and Regulation Donald J . Graves and Jerry H . Wang Purine Nucleoside Phosphorylase R. E. Parks, Jr., and R. P. Agarwal Disaccharide Phosphorylases John J . Mieyal and Robert H. Abeles Polynucleotide Phosphorylase T . Godejroy-Colburn and M . Grunberg-Manago

xix

xx

CONTENTS OF OTHER VOLUMES

The Lipases P. Desnuelle p-Galactosidase Kurt Wallenfels and Rudolf Wed Vertebrate Lysozymes Taiji Imoto, L. N. Johnson, A . C. T. North, D. C. Phillips, and J . A . Rupley Author Index-Subject Index

Volume VIII: Group Transfer, Part A: Nucleotidyl Transfer, Nucleoridyl Transfer, Acyl Transfer, Phosphoryl Transfer

Adenylyl Transfer Reactions E. R. Stadtman Uridine Diphosphoryl Glucose Pyrophosphorylase Richard L. T u r n p h t and R. Gaurth Hansen Adenosine Diphosphoryl Glucose Pyrophosphorylase Jack Preiss The Adenosyltransferases S. Harvey Mudd Acyl Group Transfer (Acyl Carrier Protein) P. Roy Vagelos Chemical Basis of Biological Phosphoryl Transfer S. J. Benkovic and K . J . Schray Phosphofructokinase David P. Bloxham and Henry A . L a d y Adenylate Kinase L. Noda Nucleoside Diphosphokinases R. E. Parks, Jr., and R. P. Agarwal

CONTENTS OF OTHER VOLUMES

xxi

3-Phosphoglycerate Kinase R. K. Scope Pyruvate Kinase F. J. Kayne Creatine Kinase (Adenosine 5’-Triphosphate-Creatine Phosphotransferase) D.c. w a t t s Arginine Kinase and Other Invertebrate Guanidino Kinases J . F. Morrison Glycerol and Glycerate Kinases Jeremy W. Thorner and Henry Paulus Microbial Aspartokinases Paolo Truffa-Bachi Protein Kinases Donal A . Walsh and Edwin G. Krebs Author Index-Subject

Index

Volume IX: Group Transfer, Part B: Phosphoryl Transfer, One-Carbon Group Transfer, Glycosyl Transfer, Amino Group Transfer, Other Transferaser

The Hexokinases Sidney P. Colowick Nucleoside and Nucleotide Kinases Elizabeth P. Anderson Carbamate Kinase L. Raijman and M . E . Jones N5-Methyltetrahydrofolate-HomocysteineMethyltransferases Robert T . Taylor and Herbert Weissbach

Enzymic Methylation of Natural Polynucleotides Sylvia J. Kerr and Ernest Borelc Folate Coenzyme-Mediated Transfer of One-Carbon Groups Jeanne I. Rader and F. M . Huennekens

xxii

CONTENTS OF OTHER VOLUMES

Aspartate Transcarbamylases Gary R. Jacobson and George R. Stark Glycogen Synthesis from UDPG W . Stalmam and H , G. Hers Lactose Synthetase Kurt E. Ebner Amino Group Transfer Alexander E. Braumtein Coenzyme A Transferases W . P. Jencks Amidinotransferages James B. Walker

N-Acetylglutamate-5-Phosphotransferase Giza De'nes Author I n d e x a u b j e c t Index

Volume X: Protein Synthesis, DNA Synthesis and Repair, RNA Synthesis, Energy-Linked ATPases, Synthetases

Polypeptide Chain Initiation Severo Ochoa and Rajarshi Mazumder Protein Synthesis-Peptide Chain Elongation Jean Lucus-Lenard and Laszlo Beres Polypeptide Chain Termination W . P. Tate and C. T . Caskey Bacterial DNA Polymerases Thomas Kornberg and Arthur Kornberg Terminal Deoxynucleatidyl Transferase F . J . Bollum Eucaryotic DNA Polymerases Lawrence A. Loeb

RNA Tumor Virus DNA Polymerases Howard M . Temin and Satoshi Mizutani

CONTENTS OF OTHER VOLUMES

DNA Joining Enzymes (Ligases) I. R. Lehman Eucaryotic RNA Polymerases Pierre Chambon Bacterial DNA-Dependent RNA Polymerase Michael J . Chamberlin Mitochondria1 and Chloroplast ATPases Harvey S.Penefsky Bacterial Membrane ATPase Adolph Abrams and Jeffrey B. Smith Sarcoplasmic Membrane ATPases Wilhelm Hasselbach Fatty Acyl-CoA Synthetases John C. Londesborough and Leslie T . Webster, Jr. Aminoacyl-tRNA Synthetases Dieter Sol1 and Paul R. Schimmel C T P Synthetase and Related Enzymes D. E. Koshland, Jr., and A. Levitzki Asparagine Synthesis Alton Meister Succinyl-CoA Synthetase William A. Bridger PhosphoribosylpyrophosphateSynthetase and Related Pyrophosphokinases Robert L. Switzer Phosphoenolpyruvate Synthetase and Pyruvate, Phosphate Dikinase R. A. Cooper and H . L.Komberg Sulfation Linked to ATP Cleavage Harr y D . Peck, J r . Glutathione Synthesis A1ton Meis ter Glutamine Synthetase of Mammals Alton Meister

xxiii

xxiv

CONTENTS OF OTHER VOLUMES

The Glutamine Synthetase of Escherichia coli: Structure and Control E . R. Stadtman and A. Ginsburg Author Index-Subject

Index

Volume XI: Oxidation-Reduction, Transfer (1)

Part A: Dehydrogenases (II , Electron

Kinetics and Mechanism of Nicotinamide-Nucleotide-Linked Dehydrogenases Keith Dalziel Evolutionary and Structural Relationships among Dehydrogenases Michael G. Rossmann, Anders Liljas, Carl-Ivar Brandin, and Leonard J . Banaszak Alcohol Dehydrogenases Carl-Ivar Brand&, Hans Jornvall, Hans Eklund, and Bo Furugren Lactate Dehydrogenase J. John Holbrook, Anders Liljas, Steven J. Steindel, and Michael G. Rossmann Glutamate Dehydrogenases Emil L. Smith, Brian M . Awten, Kenneth M . Blumenthal, and Joseph F. Nyc Malate Dehydrogenases Leonard J . Banaszak and Ralph A . Bradshaw Cytochromes c Richard E . Dickerson and Russell Timkovich Type b Cytochromes Bunji Hagihara, Nobuhiro Sato, and Tateo Yamanaka Author Index-Subject Index Volume XII: Oxidatiorr-Reduction, Part B: Electron Transfer ( I l l , Oxygenases, Oxidares (I1

Iron-Sulfur Proteins Graham Palmer

CONTENTS OF OTHER VOLUMES

Flavodoxins and Electron-Transferring Flavoproteins Stephen G. Mayhew and Martha L. Ludwig Oxygenases : Dioxygenases Osamu Hayaishi, Mitsuhiro Nozaki, and Mitchel T. Abbott Flavin and Pteridine Monooxygenases Vincent Massey and Peter Hemmerich Iron- and Copper-Containing Monooxygenases V . Ullrich and W . Duppel Molybdenum Iron-Sulfur Hydroxylases and Related Enzymes R. C. Bray Flavoprotein Oxidases Harold J . Bright and David J . T. Porter Copper-Containing Oxidases and Superoxide Dismutase B. G. Malmstrom, L.-E. Andrdasson, and B. Reinhammar Author Index-Subj ect Index

xxv

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Dehydrogenase J. IEUAN HARRIS

MICHAEL WATERS

I. Introduction . . . . . . . . . . 11. Molecular Properties . . . . . . . A. Isolation . . . . . . . . B. Enryme Structure . . . . . . 111. Catalytic Properties . . . . . . . A. Studies of Pyridine Nucleotide Binding B. Mechanism of Action of GAPDH . . C. Metabolic Role of GAPDH . . . .

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1

3 3 5 28 28 38

45

1. Introduction ( 1 )

Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) catalyzes reversibly the oxidation and phosphorylation of D-glyceraldehyde 3-phosphate (G-3P) to 1,3-diphosphoglycerate (DPGA) according to the following reaction scheme:

ocoPo~*-

CHO HAOH AHaOPOa2-

+ HPO,*- + NAD+

~

HAOH

+ H+ + NADH

LHaOPO2-

It is thus a key enzyme in the glycolytic conversion of glucose to pyruvic 1. Abbreviations used are as follows : GAPDH, GlyceraldehydeSphosphate dehydrogenase ; G 3 P , glyceraldehgde 3-phosphate ; DPGA, 1,3-diphosphoglyceric acid; LDH, lactate dehydrogenase; MDH, mdic dehydrogenase ; and ADH, alcohol dehydrogenase . 1

2

J. IEUAN HARRIS AND MICHAEL WATERS

acid which represents an important pathway of carbohydrate metabolism in most organisms. That the oxidation of G-3P was associated with the coupled phosphorylation of adenine nucleotides was originally established by Meyerhof ( l a ) and by Needham and Pillai ( 2 ) . Meanwhile the eventual isolation of the participating enzyme was prompted by the earlier observations of Lundsgaard (3)and of Green et al. ( 4 ) on the inhibition of glycolysis and of alcoholic fermentation by halogenacetic acids, and by the subsequent work of Rapkine ( 6 ) ,associating this inhibition with sulfhydryl groups of GAPDH. The precise nature of the enzymic reaction was elucidated by Warburg and Christian (6) when they succeeded in preparing GAPDH in pure crystalline form from yeast. Subsequently, isolation of the crystalline enzyme from rabbit skeletal muscle was described by Dixon and Caputto (7) and by Cori et al. (8),and in retroTABLE I

SOURCESOF PUREGAPDH’s Source

Ref.

Rabbit muscle Yeast Cat, dog, pig muscle Rabbit, ox, human, chicken, turkey, pheasant, halibut, sturgeon, lobster muscle E . coli B . stearothermophilus T . aquaticus B . cereus Coelacanth muscle Cold-adapted Antarctic fish muscle Insects Rat muscle Kangaroo muscle Pea seed Photosynthetic plants

7, 8 10

14 16

16 17,18 19

20 dl 22 93

24 25

26 d7,28

la. 0. Meyerhof, Naturwissenschaften 25, 443 (1937). 2. D. M. Needham and P. Pillai, Nature (London) 140,65 (1937). 3. E. Lundsgaard, Bwchern. Z. 217, 162 (1930). 4. D. E. Green, D. M. Needham, and J. D. Dewan, BJ 31, 2327 (1937). 5. L. Rapkine, BJ 32, 1729 (1938). 6. 0. Warburg and W. Christian, Biochem. Z.303,40 (1939). 7. R. Caputto and M. Dixon, Nature (London) 156, 630 (1945). 8. G. T. Cori, M. W. Slein, and C. F. Cori, JBC 159,565 (1945).

1. GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE

3

spect there can be little doubt that K. Bailey’s “albumin” from rabbit muscle (9)“exhibiting a pronounced sheen upon agitation’’ was in fact GAPDH. Glyceraldehyde-3-phosphate dehydrogenase occurs widely and abundantly throughout nature. It comprises about 20% of the total soluble protein in yeast (10) and up to 10% of the soluble protein from muscle (8),and the relative ease of its preparation from a wide variety of different species has contributed to its popularity among enzymologists, protein chemists, and X-ray crystallographers (cf. 11). Moreover, study of the active enzyme-NAD complex has been facilitated by the fact that uniquely among NAD-linked enzymes crystalline muscle GAPDH contains firm bound NAD. Detailed reviews of these early studies have been given by Velick and Furfine ( l a ) and by Colowick et al. (IS). II. Molecular Properties

A. ISOLATION Pure crystalline GAPDH has been isolated from a number of different sources (cf. Table I) (7,8,10,14-28). Methods of purification have relied heavily upon its solubility as the enzyme-NAD complex in high concen9. K.Bailey, Nature (London) 145, 934 (1940). 10. E. G. Krebs, G. W. Rafter, and J. M. Junge, JBC 200, 479 (1953). 11. J. I. Harris, in “Structure and Function of Oxidation-Reduction Enzymes” (A. Akeson and A. Ehrenberg, eds.), p. 639.Pergam n, Oxford, 1972. 12. S. F.Velick and C. Furfine, “The Enzymes,” pol. 7, p. 243,1963. 13. S.P.Colowick, J. Van Eys, and J. H. Park, Compr. Biochem. 14,l (1966). 14. P. Elodi and E. SzorGnyi, Acta Phgsiol. 9, 339 (1956). 15. W.S.Allison and N. 0. Kaplan, JBC 239,2140 (1964). 16. G. D’Alessio and J. Josse, JBC 246, 4319 (1971). 17. R. E. Amelunxen, BBA 122, 175 (1966). 18. K. Suzuki and J. I. Harris, FEBS (Fed. Eur. Biochem. Soc.) Lett. 13, 217 (1971). 19. J. D. Hocking and J. I. Harris, FEBS (Fed. Eur. Biochem. Soc.) Lett. 34, 280 (1973). 20. K. Suzuki and K. Imahori, J . Biochem. (Tokyo) 73,97 (1973). 21. E. Kolb and J. I. Harris, BJ 130, 26P (1971). 22. F. C.Greene and R. E. Feeney, BBA 220,430 (1970). 23. C. W.Carlson and R .W. Rrosemer, Biochemistry 10, 2113 (1971). 24. N. K.Nagradova and M. K. Guseva, Biokhimiya 36, 496 (1971). 25. R. J. Simpson and B. E. Davidson, Aust. J. Biol. Sci. 24, 263 (1971). 26. R. G.Duggleby and D. T. Dennis, JBC 249, 162 (1974). 27. W.Hood and N. G. Carr, BBA 146, 309 (1967). 28. B. A. Melandri, P. Pupillo, and A. Baccarini-Melandri, BBA 220, 178 (1970).

4

J. IEUAN HARRIS AND MICHAEL WATERS

trations (up to 70% saturation) of ammonium sulfate so that the pure muscle enzyme can be obtained from a low salt extract of blended muscle by direct crystallization from 65 to 70% ammonium sulfate. Methods for preparing enzyme from bacterial sources such as Escherichia coli (16) and B. stearothermophilus (18) have been improved by the use of chromatography on ion exchangers, while more recently Hocking and Harris (19) have prepared pure enzyme from the thermophiles B. stearotherrnophilw and Thermus aquaticus by means of affinity chromatography on immobilized NAD'. This method of preparation utilizes the strong affinity of the enzyme for suitably immobilized NAD' (it remains bound to NAD-Sephsrose in 0.7 M NaCl and is then eluted from the resin with a pulse of 10 mM NAD') which allows it to be obtained pure and in high yield from relatively crude bacterial extracts as shown in Fig. 1.

FIG.1. Purification of (A) T.aquaticua and (B) B. stearothermophilua GAPDH; SDS-gel electrophoresis (a) before and (b) after NAD-Sepharose (cf. 37).

1. GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

5

B. ENZYME STRUCTURE 1. Primary Structure

A study of the enzyme by chemical methods involving the specific labeling of catalytically active cysteine residues ($9, 30) and the characterization of peptide fragments produced by enzymic cleavage (31) led Harris and Perham to conclude that GAPDH from a given source was composed of subunits comprising approximately 330 amino acid residues corresponding to a molecular weight of 36,000. These results, considered in conjunction with the physicochemical data of Harrington and Karr (32),showed that the active enzyme with a molecular weight of 146,000 was a tetramer and that it was in all probability composed of chemically identical subunits (31). Proof that the subunits are of identical primary structure was obtained by Harris and co-workers when complete amino acid sequences were established for enzyme from lobster muscle (33), pig muscle ( 3 4 ) , and yeast (36).Comparison of the three sequences (Table 11) shows that they are strictly homologous. Moreover, 60% of the residues occur in identical sequence in the three species showing that the sequence of GAPDH has been conserved to a much greater extent than the sequence of other comparable enzymes such as, for example, alcohol dehydrogenase ( 3 6 ) .Hocking and Harris (37) have subsequently determined the sequence of GAPDH from the thermophilic bacterium T . aquaticus, and comparison of this sequence with that of the lobster muscle enzyme shows a sequence identity of 50% which is again significantly higher than was found in comparison of bacterial and liver alcohol dehydrogenase (38) or bacterial and muscle triosephosphate isomerase

-

(399) 29. J. I. Harris, B. P. Meriwether, and J. H. Park, Nature (London) 198, 154 (1963). 30. R. N. Perham and J. I. Harris, JMB 7,316 (1963). 31. J. I. Harris and R. N. Perham, JMB 13,876 (1965). 32. W. F. Harrington and G . M. Karr, JMB 13, 885 (1965). 33. B. E. Davidson, M. Sajg6, H. F. Noller, and J. I. Harris, Nature (London) 218, 1181 (1967). 34. J. I. Harris and R. N. Perham, Nature (London) 219, 1025 (1988). 35. G. M. T. Jones and J. I. Harris, FEHS (Fed. Eur. Biochem. Soc.) Lett. 22, 185 (1972). 36. H. Jornvall, Proc. Nat. Acad. Sci. U . S. 70,2295 (1973). 37. J. D. Hocking and J. I. Harris, Ezperientia (1976) (in press); J. D. Hocking Ph.D. Dissertation, University of Cambridge, 1974. 38. J. Bridgen, E. Kolb, and J. I. Harris, FEBS (Fed. Eur. Biochem. Soc.) Lett. 33, 1 (1973). 39. S. Artavanis, Ph.D. Dissertation, University of Cambridge, 1974.

TABLE I1 COMPARISON OF THE AMINO ACIDSEQUENCE OF GAPDH FROM PIOMUSCLE, LOBSTEBMUSCLE,AND YEAST^.^ 10 Asn-Gly -Phe-Gly -Arg - Ile -Gly -Arg-Leu-Val Yeast

Val-Arg-Val-Ala- Ile

Leu-Ser -&g40

Asn-Asp-Pro-Phe Gly -Ala -Gln -Val Pro-Asx-Val -Glx -Val

(Ala

&:

50

- Ile

Asx,Asx,Pro,Phe, Ile

60

Tyr-Asp-Ser -Thr-His -Gly

t

Val-Val-Glu Ser -Thr-Gly -Val -Phe Ile -Val-Glu Ala- Ile - Asp

130

120 Ala-Pro-Met-Phe-Val

Q,

Y

150

160

*C

Ser - Lys-Asp-Met-Thr-Val

Val-Ser -Asn-Ala-Ser-CYS-Thr-Thr-Asn-Cys-Leu-Ala-Pro

Leu 170

180

Glu -Gly -Leu-Met-Thr-Thr-Val -His A l a - Ile Thr-Ala -Thr-Gln-LYSAla -Val (Met-Thr, Thr, Val, His) Ser -Le 200

Thr-Val -Asp-Gly -Pro-Ger

210

220

Ser -Thr-Gly -Ah-Ala-Lys-Ala-Val-Gly -Lys-Val

230

Gly -Lys-Leu-Thr-Gly - M e t - A h -

240

Phe-Arg-Val-Pro-Thr

250

Val -Ser -Val-Val -Asp-Leu-Thr Pro-Asp Val - A s x

Glu -Thr-Thr 260

270

Leu-Gly-Tyr-Thr-GluGLx -a

TABLE I1 (Continued)

Asx Ala

Ser

Leu-Gly -Asp-Ser -His

Ser

310

300

Ser-Trp-Tyr-Asp-Asn-Glu Aax-Asx-Glx Tyr

Thr

Val Asp-Leu Met-Val H i s Met-Ala-Ser-Lys-Glu

4 ~~

From (56). b Sequences not experimentally determined for the yeast chain are given within brackets and in a provisional order that maximizes sequence homology between the yeast and muscle enzymes. c C;s-149 forms part of the active site. a

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The amino acid sequence results clearly imply a unique sequence for each of the enzymes examined, and there is no decisive evidence for the existence among GAPDH’s of tissue-specific isozymes that differ in primary sequence despite reports of the occurrence of multiple electrophoretic forms in several different organisms (40, 41). I n no case was it demonstrated that these multiple forms are the products of different genes, and it is entirely possible that electrophoretically different tetramers may have arisen by amide loss [as in the case of muscle aldolases (42)] or through differential binding of NAD (41).

2. X - R a y Structure of Hobenzyme The elucidation of the subunit structure and of the amino acid sequences of the subunits of different GAPDH’s provided the necessary framework for the interpretation of chemical modification studies as well as of X-ray crystallographic studies of the tertiary and quaternary structure of the active enzymeINAD complex. The first X-ray diffraction data for GAPDH were obtained by Watson and Banaszak (43) with crystals of enzyme from lobster muscle. These crystals, which displayed the yellow color that is characteristic of the holoenzyme, were orthorhombic (P2,2,2, space group) with the tetramer as the asymmetric unit. Essentially similar results have also been obtained with enzyme crystals from human muscle (4,46) and from B. stearothermophilus (cf. 18). A more detailed study of the lobster muscle enzyme by Rossmann and co-workers (46-48) led to the computation of the first interpretable high resolution (3 A) structure for GAPDH. The first map (with the tetramer as asymmetric unit) was interpreted by averaging the four chemically equivalent but crystallographically different subunits and, with the aid of the amino acid sequence (33),it then became possible to trace the polypeptide chain within the individual subunits. A coordinate system of P , Q , and R axes 40. H.G. Lebherz and W. J. Rutter, Science 157, 1198 (1967). 41. S. F. Velick in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 57. Springer-Verlag, Berlin and New York, 1970. 42. C.F. Midelfort and A. H. Mehler, Proc. Nut. Acad. Sci. U.S . 69, 1816 (1972). 43. H.C.Watson and L. J. Banaszak, Nature (London) 204,918 (1974). 44. A. I. Gorjunov, N. S. Andreeva, T. Baranowski, and M. Wohy, J M B 69, 421 (1972). 45. H.C.Watson, E. Due& and W. D. Mercer, Nature (London) 240, l$O (1972). 46. M. Buehner, G.C. Ford, D. Moras, K. W. Olsen, and M. G. Roasmann, Proc. Nut. Acad. Sci. U.S . 70, 3052 (1973). 47. M. Buehner, G. C. Ford, D. Moras, K. W. Olsen, and M. G. Rossmann, J M B 82, 563 (1974). 48. M. Buehner, G. C. Ford, D. Moras, K. W. Olsen, and M. G. Rossmann, J M B 90, 25 (1974).

10

J. IEUAN HARRIS AND MICHAEL WATERS

Q

P

Q

P

FIO.2. Diagrammatic comparison of the association of subunits in GAPDH (left) and LDH (right) (48).

FIo. 3. Stereoviews of the Ca atom backbone in lobster muscle GAPDH: (a) one subunit viewed to illustrate the NAD+-binding and catalytic domains; (b) the NADtbinding domain viewed in the same orientation as in (a); (c) the catalytic

1.

GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

11

similar to that used previously for lactate dehydrogenase (LDH) (49) and malate dehydrogenase (MDH) (60) has been used (Fig. 2) to define the GAPDH structure in order to draw attention to the striking structural similarities that exist between the three dehydrogenases (46, 61). The major feature of the structure is that, although exhibiting apparent 222 symmetry, the tetramer consists functionally of a dimer of dimers related across the Q axis (cf. 46). The only true twofold axis is the Q axis whereas the other axes exhibit pseudosymmetry within the limits

domain viewed in the same orientation as in (a) ; and (d) complete tetramer viewed down the P axis demonstrating the dumbbell silhouette with the four active sites close to the center of the molecule (48, 6.5). 49. M. J. Adams, A. McPherson, Jr., M. G. Rossmann, R. W. Schevitz, and A. J. Wonacott, J M B 51, 31 (1970). 50. E. Hill, D. Tsernoglou, L. Webb, and L. J. Banaszak, J M B 72,577 (1972). 51. M. G. Rossmann, A. Liljas, C.-I. Branden, and J. J. Banaszak, Chapter 2, Volume XI.

12

J. IEUAN HARRIS AND MICHAEL WATERS

of resolution obtained. The region of major interaction between subunits is across the P axis; Q-axis-related contacts are relatively few and not highly conserved, while R-axis-related contacts are again more numerous and highly conserved. The conformation of C a backbone atoms in the GAPDH subunit is shown in Fig. 3a. The subunit is envisaged as consisting of two domains (Figs. 3b and 3c), each with a specific function. The first, comprising residues 1-149, is mainly involved in NAD+ binding while the second domain, comprising residues 149-334, provides residues for substrate binding, specificity, and catalysis. The “catalytic” domain also contains most of the residues that are involved in intersubunit contacts. a. The NAD-Binding Domain. The fold of the NAD+-binding domain in GAPDH is shown diagrammatically in Fig. 4. It consists of a sixstranded parallel /3 sheet flanked by helices and is similar to analogous nucleotide binding structures in LDH, MDH, and ADH (cf. 6 1 ) . The NAD+ in each of the four subunits is bound close to the molecular waist (cf. Figs. 2 and Fig. 3d) of the tetramer and close enough to interact via a section of antiparallel sheet (comprising residues 179-200) that extends across the R axis into the adjacent subunit. This intersubunit interaction was thought to link Lys-183 in one subunit to the pyrophosphate

Fra. 4. Diagrammatic representation of the NAD+-binding domain showing the six-stranded parallel p sheet flanked by helices (48,63).

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GLYCERALDEHYDE-3-PHOSPHATE

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moiety of NAD+ in the adjacent subunit (46, 48), compatible with earlier chemical evidence implicating Lys-183 in coenzyme binding ( 5 2 ) . A revised structure (@, 53) for this part of the molecule shows, however, that Lys-183 does not interact directly with either NAD+ or substrate. Nevertheless, it remains possible that interactions between other residues in the S-shaped loop (such as, for example, Pro-188 and Trp-193 with NAD+ in the adjacent subunit could be responsible for the cooperativity of NAD+ binding (cf. Section III,A,l) and the NAD+-promoted tetramerization of the dimeric moiety. I n this respect, and as shown in Fig. 2, GAPDH differs from LDH where each molecule of NAD’ is bound entirely within each subunit with little possibility for direct interaction between binding sites within the tetramer. The conformation of the NAD+ in GAPDH is nevertheless similar to that found in LDH. Thus it is bound in an open extended configuration in each of the four subunits but with the important difference of a 180° rotation about the C-1 to N-1 glycosidic bond linking the nicotinamide ring to the ribose. This change ensures that the “B” face of the ring is exposed to the substrate for hydride ion transfer giving GAPDH its B specificity. The B or syn configuration is stabilized by hydrogen bonds formed between the carboxyamide group and the invariant Asn-313 and with the nicotinamide phosphate. It should be noted that the alternative “A” configuration of the ring that occurs in MDH, LDH, and ADH is prohibited in GAPDH due to steric hindrance involving the main chain residues Ala-120 and Pro-121 and the carboxyamide group of the nicotinamide ring. The main chain hydrogen bonding scheme in the NAD+-binding domain is shown in Fig. 5 and the topography of the NAD-binding site itself is shown diagrammatically in Fig. 6. The adenine ring binds between Phe-34 and Phe-99; a t the side of the adenine binding pocket there are hydrophobic residues Pro-33, Met-77, and Pro-79, while the inside of the pocket is more hydrophilic in character owing to the presence of Asn-6 and Asn-31. Aspartate-32 forms a hydrogen bond to the 02’ atom on the adenosine ribose while Gly-7 approaches it closely from one side. The phosphates interact with the part of the chain comprising Gly-9, Arg-10, and Leu-11; Gly-97 and Ala-120 provide a hydrophobic environment for the nicotinamide ribose while, as mentioned previously, the carbonyl group of the nicotinamide forms a hydrogen bond to Asn-313. It should be noted that the residues found to be interacting with NAD’ are highly conserved in different 52. J. H. Park, D. C. Shaw, E. Mathew, and B. P. Meriwether, JBC 245, 2946 (1970). 53. D. Moras, I<. W. Olsen, M. N. Sabeson, G . C. Ford, and M. G . Rossmann, JBC 250, 9137 (1975).

14

J. IEUAN HARRIS AND MICHAEL WATERS

FIG.5. The main chain hydrogen bonding scheme in the NAD+-binding domain (residues 1-149)(48, 63).

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GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

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FIG.0. Stereoview of the NAD'-binding site showing amino acid side chains interacting with the coenzyme. Note Phe-34 and Phe-99 on either side of the adenine ring. Aspartate-32 and Gly-7, which are close to the adenine ribose, preserve their functions in other dehydrogenases (48,651.

FIG.7. Diagrammatic representation of the catalytic domain viewed to show the large pleated sheet forming the subunit interface across the P axis (68).

16

J. IEUAN HARRIS AND MICHAEL WATERS

GAPDH’s (Table 11).Moreover, residues corresponding to Asp-32 and Gly-7 are also found in structurally equivalent positions in LDH and ADH (61). The C-4 atom of the nicotinamide ring is close to the SH group of the active site cysteine (Cys-149), and this interaction (be it a covalent bond to a tetrahedral carbon atom or a charge-transfer complex) appears to be responsible for the “Racker band” in the holoenzyme (64,66) (cf. Section 111,A). Cysteine-149, to which the substrate G-3P is bound, occurs a t the junction between the two domains in the very center of the subunit. b. The CataEytic Domain. The second half of the subunit (residues 149-334) consists primarily of a nine-stranded antiparallel sheet (Fig. 3c and Fig. 7) which forms an intersubunit contact generated by the P axis. On the other side of the sheet there are three approximately parallel helices (comprising residues 147-166, 201-216, and 215-267, respectively) and the polypeptide chain ends in a long helix (residues 313-334) that is closely associated with the first domain so that the C-terminal residue comes close to the N-terminus (Fig. 3a). The hydrogen bonding scheme for the catalytic domain is shown in Fig. 8. The essential thiol (Cys-149) initiates a short helical region in which the nonreactive Cys-153 occurs after one turn so that it comes close to Cys-149 (cf. Section II,B,4,a). The polypeptide chain then returns to the vicinity of the active center as part of an antiparallel sheet that contains His-176. There follows an S-shaped antiparallel sheet (residues 179-200) that interacts with NAD+ in the R-axis-related subunit. This segment of the chain also contains Lys-191 which has been implicated as a possible binding site for phosphate. Histidine-176, conserved in all species (36, 37), occurs within hydrogen bonding distance of Cys-149, and these two residues are clearly implicated in the catalytic mechanism (see Section II,B,4,d). c. Conservation of Amino Acid Residues. The enzyme subunit comprises approximately 330 residues. Of these 36% are in helix and 40% in p sheet, in fair agreement with the prediction of ORD (66, 67) and infrared absorption (68) measurements. Table I11 lists diagrammatically the residues that have been conserved between the three completely 54. E. Racker and I. Krimsky, Nature (London) 169, 1043 (1952). 55. I. Krimsky and E. Racker, Science 122,319 (1955). 56. P. Zavodsky, L. B. Abaturov, and Y. M. Varshavsky, Acta Biochim.Biophys. 1, 389 (1966). 57. M. E. Magar, JBC 242, 2517 (1967). 58. D. W. Darnall and T. D. Barela, BBA 236, 593 (1971).

1. GLYCERALDEHYDE-3-PHOSPHATE

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FIG.8. The main chain hydrogen bonding scheme in the catalytic domain (residues 150-334) ( 5 5 ) .

18

J. IEUAN HARRIS AND MICHAEL WATERS

TABLE I11 AMINOACIDFUNCTIONS I N GAPDHO

Symbols and abbreviations aa follows: (-*-) consewed completely among the pig, yeast, and lobster sequences; (*) in active center region; P, in contact generated by P axis; Q, in contact generated by Q axis; R, in contact generated by R axis; and D, in contact region between first and second domains within a subunit (48).

analyzed GAPDH sequences (i.e., pig, lobster, and yeast, Table 11). Noted also are the particular functions that have been recognized for a given amino acid; that is, whether it is involved (a) in the active center, (b) in one of three types of subunit contacts, or (c) in domain boundary contacts. This information is summarized in Table IV. Correlation of conserved and variable regions with the three-dimensional structure (59) shows that residues involved in catalysis and in intersubunit contacts are conserved to a much greater extent than others. It follows that the sequence of the catalytic domain with its greater proportion of active site and subunit contact residues is more highly conserved than the sequence of the NAD-binding domain despite the fact that the latter represents a highly conserved structure. 59. I(. W. Olsen, D. Moras, (1975).

M. G. Rossmann, and J. I. Harris, JBC

250, 9313

1.

19

GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE

TABLE IV OF AMINOACID RESIDUES I N GAPDHa CONSERVATION

*

P

Q

R

D

AA

First Number conserved Total number contacts Percent conserved

17 19 90

0

0 -

3 3 100

5 5 100

9 13 69

71 148 48

Second Number contacts Total number contacts Percent conserved

11 12 92

27 33 82

2 5 40

10 12 83

2 5 40

124 186 67

Both Number contacts Total number contacts Percent conserved

28 31 90

27 33 82

5 8 63

15 17 88

11 18 61

195 334 58

Domain

symbols and abbreviations as follows: (*) involved in active center; P, in a P-axis-generated contact between subunits; Q, in a Q-axis-generated contact between subunits; R, in a R-axis-generated contact between subunits; D, in a domain-domain contact within a subunit; and AA, amino acids involved (48).

3. Structure of Apoenzyme Apoenzyme prepared from muscle holoenzyme by treatment with charcoal is unstable and difficult to crystallize (60, 61). Consequently, it has not so far been possible to solve the three-dimensional structure of apo-GAPDH by X-ray crystallographic methods. Suzuki and Harris (18) were able to prepare stable crystals suitable for X-ray diffraction analysis of both holo- and apoenzyme from the thermophile B. stearothermophilus. GAPDH from this source is considerably more stable than enzyme from mesophiles (17,18), and this stability is retained even in the absence of NAD+ (Fig. 9 ) . Wonacott and colleagues (62,cf. 18) have shown that these holoenzyme crystals are orthorhombic with space group P2,2,2; the unit cell, like that of the lobster muscle enzyme, consists of four tetramers. Apoenzyme crystals were found to be monoclinic (space group P 2 , ) , and the unit cell consists of two tetramers. It is known that the binding sites for NAD’ are not equivalent and that conformational changes occur when NAD+ interacts with apoenzyme in solution (for references see Section 111,A). These changes have been shown to involve a volume contraction of about 776, possibly because 60. C. S. Furfine and S. F. Velick, JRC 240,814 (1965). 61. P. M. Wassarman and H. C. Watson, in “Enzymes and Isoenzymes: Structure, Properties and Function” (D. Shugar, ed.), p. 51. Academic Press, New York, 1970.

20

J. IEUAN HARRIS A N D MICHAEL WATERS

D Temperature .C

FIG.9. Comparative thermal stabilities of GAPDH’s from

(A)rabbit

muscle,

( 0 )B. stearothermophilus, ( A ) T . aquaticus holoGAPDH; a n d y o ) T . aquaticus apoGAPDH (ST).

of the expulsion of solvent molecules as would be expected to occur if NAD+ binding gives rise to a more tightly packed tetramer. A separate determination of the three-dimensional structure of apoenzyme and identification of the structural changes that occur when NAD+ binds to the protein subunits will be necessary in order to understand fully the cooperative phenomena associated with the mechanism of the enzymic reaction (cf. Section 111,AJ). 4. Chemical Modification of Native Enzyme

a. Cysteine Residues. The cysteine content of GAPDH is variable, ranging between five SH groups per subunit for the lobster muscle enzyme (33) and one SH group per subunit for enzyme from T . aquaticus (19, 3 7 ) . The essential cysteine (Cys-149) is also the most reactive, and it is the selective reaction of this conserved residue in GAPDH that is responsible for the inhibition of glycolysis and of fermentation by iodoacetic acid (6, 9.9). Cysteine-149 also reacts selectively with a number of other reagents such as p-fluorodinitrobenzene (63),tetrathionate (64-66), iodosobenzoate 62. 63. 64. 65. 66.

A. J. Wonacott, unpublished results. S. Shaltiel and S. Soria, Biochemistry 8,4411 (1969). A. Pihl and R. Lange, JBC 237, 1356 (1962). W. S. Allison and N. 0.Kaplan, Biochemistry 3, 1792 (1964). D. J. Parker and W. 9. Allison, JBC 244,180 (1969).

1. GLYCERALDEHYDE-%PHOSPHATE

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21

(66, 671, and various organic mercurials (cf. 68, 69). It is also acylated during the enzyme-catalyzed hydrolysis of p-nitrophenylacetate (29, SO) and acetyl phosphate (70). Cysteine-153, conserved in all known GAPDH's [with the one exception of T.a q w t i w (S7')], is not reactive in the native structure although under certain conditions it is capable of forming an intrachain disulfide bond with Cys-149 (66-68).The initially reversible inactivation of GAPDH by iodosobenzoate or tetrathionate is thought to result from the formation of either a sulfenic acid or a sulfenyl thiosulfate derivative of Cys-149. Upon standing, and more especially upon heating or the addition of urea, these derivatives are able to react with Cys-153 (which occurs after one turn of a helix, and thus close to Cys-149 in the tertiary structure) to form the disulfide bond. Subsequent reduction with thiol does not restore enzymic activity, perhaps because formation of the ring introduces an element of strain into the helix which may then induce an irreversible conformational change, possibly associated with the displacement of Tyr-311 from the active site. It is, therefore, probably significant that Cys-153 has been replaced by serine in T. aquaticus GAPDH (37) since disulfide ring formation could lead to oxidative inactivation of the enzyme at the high growth temperature (7Oo-75OC) of this organism. With lobster GAPDH formation of the intrachain disulfide ring between Cys-149 and Cys-153 occurs following reaction of Cys-149 with DTNB ( 7 1 ) . This leads to a gradual unfolding of the structure so that the other three buried SH groups also become exposed and reactive toward DTNB. It should also be noted that the initial reaction of Cys-149 in the holoenzyme with DTNB causes NAD to be released (69) prior to disulfide bond formation. Reaction of the DTNB apoenzyme with Cys-153 and the irreversible changes accompanying this process may be a contributory factor in the instability of the apoenzyme derivative. b. Lysine Residues. A lysine residue in GAPDH, identified as Lys-183 in the primary sequence (34, 72),is acetylated irreversibly when apoenzyme is allowed to react with either p-nitrophenylacetate or acetyl phosphate (7'0, 73, 7'4) at alkaline pH. N-Acetylation occurs not by direct reaction with Lys-183 but as the result of an S to N migration of acetyl 67. J. I. Harris and R. N. Perham, Proc. Znt. Congr. Biochem., 6th, 1964 Vol. 32, Sect, IVS27, p. 293 (1964). 68. P. M. Wassarman, H. C. Watson, and J. P. Major, BBA 191, 1 (1969). 69. P. J. Harrigan and D. R. Trentham, BJ 124, 573 (1971). 70. E. Mathew, B. P. Meriwether, and J. H. Park, JBC 242, 5024 (1967). 71. P. M. Wassarman and J. P. Major, Biochemistry 8, 1076 (1969). 72. J. I. Harris and L. Polghr, JMB 14, 630 (1965). Q 25, 1 (1964). 73. L. Polg&r,A C ~Physiol. 74. L. Polgdr, BBA 118, 276 (1966).

22

J. IEUAN HARRIS AND MICHAEL WATERS

groups from Cys-149. This implies that Lys-183 is close to Cys-149 in the quaternary structure of the apoenzyme, an observation that was confirmed by the use of cross-linking reagents ( 7 5 ) .The acyl migration reaction does not occur in the presence of NAD', and it was not established if the reaction occurs within or between monomers. Acetylation of Lys183 was found to inhibit NAD binding as well as dehydrogenase activity, and for these reasons it was suggested ( 5 d ) that Lys-183 is itself involved in coenzyme binding. With yeast enzyme (35) the acetylation reaction appeared to be less specific; thus, at pH 8.5 some reaction was also noted with lysine residues at positions 212, 216, and 266, which may reflect differences in the mode of binding of NAD in the yeast enzyme. Reaction of the rabbit muscle holoenzyme with pyridoxal phosphate, which results in total inactivation, is specific for Lys-191 and Lys-212 ( 7 6 ) . With apoenzyme, on the other hand, pyridoxal phosphate reacts with Lys-212 only, suggesting that a conformational change involving Lys-191 takes place when NAD' is removed from the holoenzyme ( 7 7 ) . It is of interest that Lys-191 and Lys-212 are conserved in all the sequenced species of GAPDH (see Tables I1 and 111, also 35) ; Lys-183, on the other hand, is not conserved and its replacement by arginine in the B. stearotherrnophilus (78) and T . aquaticus (37) enzymes explains why the S to N acetyl transfer reaction does not occur with these enzymes. The position of Lys-183 in the three-dimensional structure of the lobster holoenzyme (53) indicates that it is about 10 A distant from the phosphate groups of the NAD'; moreover, its C-NH, group is 20 A distant from Cys-149 in the R-axis-related subunit. These distances are such as to suggest that the quaternary structure of the apoenzyme must be substantially altered with respect to the holoenzyme to permit a much closer approach of the two groups if direct S to N transfer (70, 73, 74) or crosslinking with 1,5-difluoro-2,4-dinitrobenzene (75) is to occur. It follows that there is no direct interaction of Lys-183 in the holoenzyme with either NAD' or substrate and that this particular lysine has no obvious role in the mechanism of the catalytic reaction normally performed by GAPDH. c. Tyrosine Residues. Iodination of lobster muscle GAPDH with K [ lZ5I] led to the identification of tyrosine residues of differing reactiv75. S. Shaltiel and M. Tauber-Finkelstein, BBRC 44, 484 (1971). 76. B. G. Forcina, G . Ferri, M. C. Zapponi, and S. Ronchi, Eur. J . Biochem. 20, 535 (1971). 77. M. C. Zapponi, G . Ferri, B. G . Forcina, and S. Ronchi, FEBS (Fed. Eur. Biochem. Soc.) Lett. 31,287 (1973.) 78. J. Bridgen rind J. I. Harris, Znt. Congr. Biochem, 9th, 1973 Abstract 2e.1, p. 59 (1973).

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ity (79). Tyrosine-46 was found to be the most reactive residue in the native holoensyme, but significant specificity was attained only in the presence of limiting amounts of iodine. Two other residues, Tyr-39 and Tyr-42, were moderately reactive and several of the other nine tyrosines in the subunit were also found to react, albeit a t appreciably slower rates. Iodination of Tyr-46 does not cause inactivation, and its special reactivity is entirely compatible with its exposed position on the outside of helix aC in the. three-dimensional structure of the tetramer. Tyrosine-39 and Tyr-42 are in the same helical segment but are partially shielded by interactions with neighboring subunits. Tyrosine-46, followed by Tyr-39 and Tyr-42, were also found to be the most susceptible to iodination in the pig holo- and yeast enzymes (79) again indicating that the threedimensional structure of GAPDH has been highly conserved. With pig spoensyme, on the other hand, Libor and Elodi (80) found Tyr-137 and Tyr-252 [which Thomas and Harris (79) found to be among the least reactive in the holoenzyme] to be more reactive than Tyr-46, Tyr-42, and Tyr-39, a result that is difficult to reconcile with the three-dimensional structure. The extent to which the reactivities of tyrosines can be correlated with their positions in the native three-dimensional structure is however difficult to assess with complete certainty owing to possible effects on the structure of side reactions such as oxidation of thiol groups (cf. Section II,B,4,a) and iodination of histidine residues. Thus, dependence of specificity on reaction time and reagent concentration could be the consequence of slow but irreversible conformational changes within the tetrameric structure, and in this connection it should be noted that apoensyme is less stable than holoeneyme in the presence of iodine a t alkaline pH (79). d. Histidine Residues. Although histidine has been implicated in the catalytic mechanism of GAPDH (see, for example, 81-83, and Section III,B,l), there is, perhaps surprisingly, no convincing chemical evidence for the direct involvement of a specific histidine in the active site. Thus, for example, Moore and Fenselau (84) could not link Cys-149 to any neighboring histidine residue in the rabbit enzyme with the bifunctional dibromoacetone; Allen and Harris (86) were also unable to achieve specific labeling of an essential histidine in the B. stearothermophilus en79. J. 0. Thomas and J. I. Harris, BJ 119, 307 (1970). 80. S. Libor and P. Elodi, Em-. J. Biochem. 12, 336 (1970). 81. E. J. Olson and J. H. Park, JBC 239, 2316 (1964). 82. P. Friedrich, L. PolgLr, and G . Szabolcsi, Acta Ph,gsiol. 25, 217 (1964) 83. P. J. Harrigan and D. R. Trentham, BJ 135, 695 (1973). 84. J. Moore. Jr. and A. Fenselau, Biochemistry 11, 3753 (1972). 85. G. A. Allen and J. I. Harris, unpublished results.

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J. IEUAN HARRIS AND MICHAEL WATERS

zyme with either 3-bromoacetypyridyine or bromopyruvate [reagents known to react selectively with His-195 in LDH (86, 87)’J.Iodination of His-50 had no effect on activity (86), and likewise Ovbdi and Keleti (88) showed that four of the eleven histidines in the pig enzyme were not essential for activity. The first positive evidence for histidine in the active site came from the photooxidation studies of Park and colleagues (89,90). These studies appeared to show that inactivation of the rabbit muscle enzyme was associated with the specific destruction of His-38. Significantly, however, this particular histidine does not occur in the lobster and yeast enzymes, and its involvement in the active site was therefore considered to be unlikely ( 3 6 ) .The three-dimensional structure (47) confirms that an invariant histidine (His-176, Table 11) is present in the active site within hydrogen bonding distance of Cys-149. Moreover, residue 38 (glutamic acid in lobster GAPDH) was shown to be on the surface of the tetramer and a considerable distance away from the active site. A histidine in this position would be expected to be particularly susceptible to photooxidation, and it seems likely therefore that an alternative explanation must be sought for the observed effects of photooxidation (89,90) on the activity of the rabbit enzyme. 5. Dissociation and Hybridization Early discrepancies in determinations of the molecular weight of rabbit muscle and yeast GAPDH’s (91-93) were resolved by the definitive sedimentation equilibrium studies of Harrington and Karr (32) in conjunction with the chemical investigations of Harris and Perham ( 3 1 ) . These established that the enzyme is a tetramer with a molecular weight of 146,000 and that the 7.5 S tetramer dissociates to an unfolded 1.85 monomer in 5 M guanidine. Loss of activity below pH 5 and above pH 11, and in 8 M urea, is also accompanied by dissociation and subsequent unfolding of the subunits as evidenced by sedimentation velocity, light 86. C. Woenckhaus, J. Berghauser, and G. Pfleiderer, Hoppe-Seyler’s Z. Physwl. Chem. 350,473 (1909). 87. M. G. Rossmann, M. J. Adams, M. Ruehner, G. C. Ford, M. L. Hackert, P. J. Lentz, A. McPherson, R. Schevitz, and I. E. Smiley, Cold Spring Harbor Symp. Quant.Biol. 36, 179 (1971). 88. J. Ov&diand T. Keleti, Acta Biochim. Biophys. 4, 365 (1909). 89. J. C. Bond, S. H. Francis, and J. H. Park, JBC 245, 1041 (1970). 90.S. H. Francis, B. P. Meriwether, and J. H. Park, Biochemistry 12, 346 (1973). 91. J. B. Fox, Jr. and W. B. Dandliker, JBC 218,53 (1950). 92. J. F. Taylor and C. Lowry, BBA 20, 109 (1950). 93. H-G. Elias, A. Garbe, and W. Lamprecht, Hoppe-Seyler‘s Z. Physbl. Chem. 319, 22 (1900).

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scattering, difference spectroscopy, and ORD measurements (94-98). High ionic strength also appears to promote dissociation (98-103), and on the basis of these observations it may be concluded that electrostatic interactions and hydrogen bonds are important in maintaining the tetrameric structure. There is evidence, too, that the rat muscle enzyme is dissociated a t low temperature with release of hydrophobic residues (24); that ATP promoted dissociation of yeast and rabbit muscle enzymes is enhanced a t Oo (104); and that detergents can cause dissociation without extensive unfolding of the subunits (67,106), showing that hydrophobic interactions also play a part in stabilizing the tetrameric structure. In all the above-mentioned studies, only the tetramer has catalytic activity. The only claims for an active dimer (106,107)are not supported by satisfactory experimental evidence. Deal and co-workers (104, 108) have, on the other hand, presented extensive studies of the ATP-induced dissociation of GPD to inactive dimers and monomers a t low temperatures. Furthermore, these subunits display very little unfolding, which has been taken to imply that dissociation is the major factor in the activity loss. These results were confirmed (24) with the rat skeletal muscle enzyme, which dissociates a t Oo to inactive dimers in the absence of ATP. In addition, the activity transport studies of Hoagland and Teller (109) have given strong evidence that only the tetrameric form is active and that the presence of all three substrates promotes tetramer formation. It was also shown that rabbit muscle enzyme exists in a dimer-tetramer equilibrium in dilute aqueous solution a t 5 O , with an association constant 94. P. Elodi, G. Jecsai, and A. Morolovsky, Acta Physiol. 17, 165 (1960). 95. P. Elodi and S. Libor, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 175. Springer-Verlag, Berlin and New York, 1970. 96. W. C. Deal and W. H. Holleman, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 23, 983 (abstr.) (1964). 97. Y. Shibata and M. J. Kronman, ABB 118, 410 (1967). 98. R. Jaenicke, D. Schmid, and S. Knof, Biochemistry 7,919 (1968). 99. E. A. Meighen and H. K. Schachman, Biochemistry 9, 1177 (1970). 100. R. Jaenicke, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 209. Springer-Verlag, Berlin and New York, 1970. 101. K. Kirschner and I. Schuster, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 217. Springer-Verlag, Berlin and New York, 1970. 102. G. M. Spotorno and M. R. Hollaway, Nature (London)226,756 (1970). 103. K. Suzuki and J. I. Harris, J. Bwchem. (Tokyo) 77,587 (1975). 104. S. M. Constantinides and W. C. Deal, Jr., JBC 244, 5695 (1969). 105. I. A. Bolotina, D. S. Markovich, M. V. Volkstein, and P. Zavodsky, BBA 132, 280 (1967). 106. A. I. Agatova, Biokhimivn 32,915 (1967). 107. J. Ovadi, M. Telegdi, J. Batke, and T. Keleti, Eur. J. Biochem. 22, 430 (1971). 108. S. T. Yang and W. C. Deal, Jr., Biochemistry 8,2806 (1969). 109. V. D. Hoagland and D. C. Teller, Biochemistry 8, 594 (1969).

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J. IEUAN HARRIS AND MICHAEL WATERS

of 2 X lo6 liters/mole. Removal of bound NAD’ shifts this equilibrium toward dissociation (KaS8apoenzyme = 0.4 X lo6 liters/mole) . Generally, similar conclusions were reached by others (1212, 99, 110-1112). Other evidence that a dimer-tetramer equilibrium exists is provided by hybridization studies with GAPDH’s from different sources. Thus, Kirschner and Schuster (101), Spotorno and Hollaway (log),and Stallcup and Koshland (113)were able to obtain Y,R, hybrids of yeast and rabbit muscle enzymes a t high enzyme concentrations and physiological pH suggesting that interchange of dimeric subunits can occur. Moreover, NAD’ was shown to prevent hybridizstion, which suggests that the coenzyme promotes tetramer formation. Lebherz et al (114) found that ATPinduced hybridization of trout GPD’s was likewise blocked by NAD’ or NADH. Using conditions of high ionic strength, Meighen and Schachman (99) were able to obtain small amounts of SRs and SsR hybrids of native and succinylated rabbit muscle enzyme, which implies that monomer formation had also occurred. Suzuki and Harris (103)used 3 M NaCl to hybridize a number of different GAPDH’s including the rabbit, pig, lobster, yeast, and E . coli enzymes. As shown in Fig. 10 different pairs of electrophoretically distinct enzymes gave rise to five-membered hybrid sets indicating that dissociation to monomers had occurred. The relative amounts of dimer and monomer formed by dissociation of tetramers comprising pairs of isologous associations will be determined by the tetramerdimer and dimer-monomer dissociation constants. In experiments involving yeast enzyme the dimeric form appears to predominate while with the rabbit, pig, lobster, and E. coli enzymes dimers readily dissociate to monomers. The relative ease with which enzymes from such phylogenetically different sources are able to form hybrid tetramers is indicative of a high degree of conservation of tertiary and quaternary structure among GAPDH’s. Moreover, the successful hybridization (103) of pig enzyme with lobster enzyme, which had been inactivated by carboxymethylation of Cys-149 in all four subunits, to form four enzymically active protein bands, including the tetramer that contained only one pig subunit, showed that individual subunits are able to express their activity independently even within hybrid tetramers formed with inactive chemically modified subunits of another species. 110. L. A. Fahien, JBC 241, 4115 (1966). 111. S. Lakatos, P. Zavodaky, and P. Elodi, FEBS (Fed. Eur. Biochem. Soc. Lett. 20, 324 (1972). 112. G. D. Smith and H. K. Schachman, Biochemistry 12, 3789 (1973). 113. W. B. Stallcup and D. E. Koshland, Jr., I M B 80,41 (1973). 114. H. G. Lebhera, B. Savage, and E. Abacherli, Nature (London) New Biol. 245, 269 (1973).

1. GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

FIG.10. Hybridization of GAPDH’s: (a) rabbit (R) and lobster (L) muscle, pig (P) and lobster (L)muscle; (b) rabbit (R) and yeast (Y), pig (P) and yeast (Y). The hybrid bands were separated by electrophoresis on cellulose acetate in 50 mM phosphate buffer, pH 7.0, and revealed in (a) by protein staining and in (b) by activity staining (cf 103). 27

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J . IEUAN HARRIS AND MICHAEL WATERS

111. Catalytic Properties

A. STUDIES OF PYRIDINE NUCLEDTIDE BINDING The binding of NAD+ and of coenzyme analogs to GAPDH has been studied extensively, and a detailed discussion of earlier work has been given by Colowick et al. (13). Racker and Krimsky (64) were the first to show that binding of NAD’ to the apoenzyme produces a unique yellow complex with ti broad absorption maximum around 365 nm. As normally isolated, GAPDH’s contain from 3 to 4 moles of tightly bound NAD’ per mole of tetramer, although there are exceptions such as the yeast and sturgeon muscle enzymes which are isolated with little or no bound coenzyme (cf. 16).The “Racker band” is either diminished or abolished by oxidation or alkylation of the active site thiol 149; and by acylation with the substrate (DPGA) or substrate analogs such as acetyl phosphate (cf. 116-1173,Modification of the thiol group results in a decrease in the binding constant for NAD’, although the cooperativity of binding is maintained (118).Moreover, no major change in conformation of the lobster muscle enzyme could be detected a t 6 A resolution (61) following carboxymethylation of Cys-149. Crystals of carboxymethylated enzyme were completely isomorphous with those of native enzyme and could be distinguished only by the conspicuous lack of yellow color implying that the loss of Racker band absorbance is an effect that is restricted to the active site. In this connection it is of interest that NAD’ protects the active site thiol against alkylation by most alkylating agents (119) with, however, the notable exception of iodoacetic acid (cf. 117). Kosower (120)suggested that the Racker band resulted from a charge-transfer interaction between the nicotinamide ring and an electron donor in the enzyme. The nucleophilic nature of Cys-149 pointed toward a thiolate anion as the likely electron donor (117,121, 122), although it was also suggested (123,124) on the basis of spectral studies with analogs that 115. E. Racker and I. Krimsky, JBC 198, 731 (1952). 116. D. R. Trentham, BJ 122,59 (1971a). 117. D. R. Trentham, BJ 109,603 (1968). 118. W. Boers and E. C. Slater, BBA 315, 272 (1973). 119. B. Eisele and K. Wallenfels, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 91. Springer-Verlag, Berlin and New York, 1970. 120. E. M. Kosower, JACS 78,3497 (1956). 121. L. Boross and E. Cseke, Acta Biochim. Biophys. 2, 47 (1967). 122. E. Cseke and L. Boross, Acta Biochim. Biophys. 5,385 (1970). 123. G. Cilento and P. Tedeschi, JBC 238, 907 (1961). 124. S.Shifrin, BBA 81,205 (1964).

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tryptophan might act as the donor in a charge-transfer complex with the nicotinamide. However, there is no tryptophan in the vicinity of the nicotinamide ring in the tertiary structure (47) whereas the thiol group of Cys-149 is ideally placed to form either a covalent bond or a chargetransfer complex with the NAD’. Strong interactions of this type between the essential thiol and the nicotinamide could adequately account both for the tight binding between NAD+ and the enzyme and for the loss of the Racker band upon modification of the thiol group. Moreover, it seems probable that this interaction would stabilize the thiolate anion. The fact that 260 nm excitation of the enzyme-NADH complex led to NADH emission a t 460 nm, as well as the apparent lack of spectral shift on NADH binding, led Velick (126)and others (IS, 126) to suggest that the nucleotide was bound to GAPDH in a folded conformation. The basis for this suggestion was the finding that in aqueous solutions, NADH emission a t 460 nm can be excited by light a t both 260 and 340 nm. This led to the idea that adenine and nicotinamide rings were coupled in parallel in a hairpin or folded conformation to allow for transfer of excitation energy between the rings. Loss of 260 nm excitation maximum and the accompanying spectral shift that occurs when NADH binds to LDH (127) were therefore taken to indicate an unfolding of the nucleotide into its open form. In addition, NMR studies have shown that the pyridine nucleotides exist predominantly in the folded conformation in solution (128) and that spectral shifts occur on binding to GAPDH (129) that are opposite to those seen with other dehydrogenases in which the coenzyme is known to bind in the open form. These interpretations are in conflict with the X-ray crystallographic model which shows beyond doubt that the coenzyme in GAPDH is bound, as in LDH, in the open form. It is possible, however, that the 180° rotation about the glycosidic bond linking the nicotamide ring to the ribose in NAD+ bound to GAPDH may account for the observed differences in the NMR spectra, and in this connection it is of interest that the spectral shifts in NADH absorption that occur on binding to B-specific dehydrogenases were also found to occur with GAPDH (130). It seems clear therefore that NAD’ is bound in the open form with the nicotinamide ring close to the reactive 125. S. F. Velick, in “Light and Life” (W. D. McElroy and B. Glass, eds.), p. 108. Johns Hopkins Press, Baltimore, Maryland, 1961. 1%. D. Eby and M. E . Kirtley, Biochemistry 10,2677 (1971). 127. S.F. Velick, JBC 233, 1455 (1958). 128. N. J. Oppenheimer, L. J. Arnold, and N. 0. Kaplan, Proc. Nut. Acud. Sn’. U.S. 68, 3200 (1971). 129. C. Y. Lee, R. D. Eichner, and N. 0. Kaplan, Proc. Nut. Acud. Sn’. U. S. 70, 1593 (1973). 130. H. F. Fisher, D. L. Adija, and D. G. Cross, Biochemistry 8, 4424 (1969).

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J . IEUAN HARRIS AND MICHAEL WATERS

thiol as demanded by the observation (131) that a hydride ion is transferred directly from the substrate to the nicotinamide ring. Substrate analog and kinetic studies gave some insight into the other groups involved in nucleotide binding. Thus, of the many nicotinamide ring analogs tested, only the acetyl pyridine, thionicotinamide, nicotinylhydroxamic acid, and nicotinic acid hydrazide analogs of NAD' can act as substrates, albeit with higher K , values (132). The activity of the acetylpyridine analog shows the the amide group of nicotinamide is not essential. The carbonyl group is important since, as is shown in the crystallographic model, it stabilizes the B conformation of the coenzyme by forming a hydrogen bond with Asn-313. It is of interest that the pyridine 3-aldehyde analog which is inhibitory .does not show Racker band absorption whereas the active acetyl pyridine analog does (133).Both deamino NAD' and acetyl pyridine NAD' retain the negative cooperativity of binding seen with NAD+ (126). Of the adenine analogs tested, only the 6-deamino analog exhibits activity, again with a high K , (126). From the model, it is likely that this amino group would interact with the highly conserved residue glutamate-76. Substitution of a phosphate on the 2 position of the ribose (NADP') or removal of the hydroxyl group (deoxyadenosine) abolishes activity and binding ability (108) in agreement with the binding of the 2-hydroxy group by Asp-32 seen in the model. Moreover, competitive inhibition by hydrophobic probes for the NAD+-binding site (24) may be the result of competition for the hydrophobic pocket bounded by Phe-34 and Phe-99, which contains the adenine moiety. The overall picture that emerges from these results is in general agreement with the X-ray crystallographic evidence and shows that the binding interactions between the protein and the NAD' are well distributed over the nicotinamide, pyrophosphate, ribose, and adenine parts. Moreover, strong binding requires all these interactions to be effective simultaneously. The fact that there is an adenine binding pocket that is distinct from the catalytic site is in agreement with the binding model proposed by Yang and Deal (108) and effectively disproves the closed NAD+ model (126,126).

1. Cooperativity of NAD' Binding The early studies of Velick et al. ( l S 4 ) first delineated the two aspects of NAD+ binding that have provided the basis for subsequent work. These 131. W. S. Allison, M. J. Connors, and D. J. Parker, BBRC 34, 503 (1969). 132. B. M. Anderson and N. 0. Kaplan, JBC 234, 1226 (1959). 133. N. 0. Kaplan, M. M. Ciotti, and F. E. Stolzenbach, ABB 69, 441 (1957). 134. S. F. Velick, J. E. Hayes, and J. Harting, JBC 203, 527 (1953).

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workers found that the dissociation constant for NAD’ changed as saturation of the enzyme was approached, and that binding of NAD‘ to the apoenzyme produced different solubility and stability characteristics, which led them to suggest that “NAD’ binding is associated with changes in the configurational state of the protein.’’ A number of subsequent studies showed that NAD’ binding produces changes in viscosity (136),and in optical rotatory dispersion (106,136, 137) and electron paramagnetic resonance spectra (138).NAD’ binding also alters the rate of exchange of the peptide bond amide groups with deuterium oxide and produces a small increase in sedimentation and diffusion parameters (66). The large difference between kinetically and spectrally determined dissociation constants (cf. la) and conflicting reports about the number (110,139) and independence of NAD+-binding sites (140,1.41) led to a period of confusion which was largely resolved by the finding of positive cooperativity of NAD’ binding in the yeast enzyme (149)and negative cooperativity of binding in the muscle enzyme (143,144). a. The Yeast Enzyme-A Concerted Binding Mechanism. Using a combination of equilibrium and rapid kinetic techniques, Kirschner and co-workers (101,142, 146-147) have shown that the binding of NAD’ to the yeast enzyme can be described by the concerted model of Monod et al. (148).According to this proposal, the enzyme exists in two symmetrical tetrameric forms, R and T, and a relatively slow T to R conversion occurs on addition of NAD’ to the apoenzyme which exists in about 98% T form. The T form is enzymically inactive and possesses highly reactive thiol groups. The R form, on the other hand, has a higher Racker band absorbance than the T form and a greater affinity for NAD’ (Kd lo-* M compared to 2 x M ) . The isomerization can therefore be clearly distinguished both in terms of a time-dependent interconversion process and by the separate characteristics of the two forms. The coopera135. P. Elodi and G. Szabolczi, Nature (London) 184,56 (1959). 136. B. H. Havsteen, Acta Chem. Scand. 23, 2193 (1969). 137. I. Listowsky, C. S. Furfine, J. J. Betheil, and S. Englard, JBC 240, 4253 (1965). 138. W. Balthasar, Eur. J . Biochem. 22, 158 (1971). 139. A. L. Murdock and 0. J. Koeppe, JBC 239, 1983 (1964). 140. A. Stockell, JBC 234, 1286 (1959). 141. B. Chance and J. H. Park, JBC 242, 5093 (1967). 142. K. Kirschner, M. Eigen, R. Bittmann, and B. Voigt, Proc. Nat. Acad. Sci. U . S. 56, 1661 (1966). 143. A. Conway and D. E. Koshland, Jr., Biochemistry 7 , 4011 (1968). 144. J. J. M. De Vijlder and E. C. Slater, BBA 167, 23 (1968). 145. K. Kirschner, in “The Regulation of Enzyme Activity and Allosteric Interactions” (E. Kvamme and A. Pihl, eds.), p. 39. Academic Press, New York, 1968. 146. K. Kirschner, J M B 58, 51 (1971). 147. K. Kirschner, E. Gallego, I. Schuster, and D. Goodall, J M B 58, 29 (1971). 148. J. Monod, J. Wyman, and J.-P. Changeaux, J M B 12, 88 (1965).

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tivity is not readily detected a t 20° and pH 7.4, but increases as the temperature or pH is raised to 40° and pH 8.5.Even a t the higher temperature and pH, each form of the tetramer displays equivalent and independent binding sites, as predicted by the concerted model; and there is no evidence for significant quantities of the hybrid forms which would be predicted by the sequential model of Koshland et al. (149). Later studies (160) showed that the binding of NADH is hyperbolic in contrast to that of NAD’. Thus, NADH binds equally well to both R and T forms, with the lower affinity characteristic of NAD’ binding to M).These observations have been the T form (Kd NADH 2 x rationalized by postulating that strong interaction with the adenine and pyridinium carboxamide sites is possible only in the enzymically active R form, and since reduced NAD+ does not possess the quaternary pyridinium ring, it cannot stabilize the R form on binding. Thus, the enzyme would be active in the reverse reaction only in the presence of NAD’, which would convert the enzyme to the R form, as has been observed (161; but see Section III,B for an alternative hypothesis). In this connection it should be noted that bound NADH fluorescence is enhanced in the yeast enzyme (160)and quenched in the muscle enzyme (197),which would imply a difference in the nucleotide binding site, in keeping with the difference in cooperativity characteristics. Support for a concerted model for the yeast enzyme has come from X-ray small angle scattering experiments (169) as well as from hydrodynamic and optical rotation studies (165, 164). A volume contraction of about 5% occurs on binding of NAD’ to the apoenzyme, presumably related to tightening of the tetramer and expulsion of water molecules. The relation between NAD’ bound (R) and change of volume (Y) was hyperbolic, in accord with the concerted model. It was later shown (166) from buoyant density and preferential hydration studies that water is indeed excluded from the yeast enzyme on binding to NAD’, such that a volume contraction of about 6% occurs. Furthermore, fluorimetric and calorimetric titrations over the range 5O-4Oo showed independence of 149. D. E. Koshland, G. Nkmkthy, and D. Filmer, Biochemistry 5, 365 (1966). 150. G. von Ellenrieder, K. Kirschner, and I. Schuster, Eur. J . Biochem. 26, 220 (1972). 151. J. J. M. De Vijlder and B . J . M. Harmsen, BBA 178, 434 (1969). 152. H. Durchschlag, G. Puchwein, 0. Kratky, I. Schuster, and K. Kirschner, Eur. J . Biochem. 19,9 (1971). 153. R. Jaenicke and W. B. Gratzer, Eur. J . Biochem. 10,158 (1969). 154. R. Jaenicke, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.) p. 209. Springer-Verlag, Berlin and New York, 1970. 155. D. L. Sloan and S. F. Velick, JBC 248, 6419 (1973).

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33

NAD' binding a t pH 7.4 (166) and positive cooperativity a t pH 8.5 (166), in agreement with earlier studies (101). Eisele and Wallenfels (119) have also provided support for the concerted model with studies on the rate of stereoselective inhibition of the yeast enzyme by the antipodes of u-iodopropionic acid and a-iodopropionamide a t varying NAD' concentrations. I n similar experiments the muscle enzyme showed negative cooperativity. b. The Muscle Enzyme-A Complex Sequential Binding Mechanism. Conway and Koshland (143) showed, on the basis of equilibrium binding studies, that each molecule of NAD+ binds with decreasing affinity (Kl M) to the tetrameric rabbit lo-", K 2 K s 3 X lo-', and K , 2.6 x muscle enzyme. These binding constants cannot be described by the statistical relationship predicted from the concerted model, where each molecule binds to an equivalent site on the R state tetramer, and can only be accounted for by introducing terms for negative interaction between the subunits (149) implying the existence of hybrid states between the R and T forms. Here, in contrast to the yeast enzyme, symmetry forces are not as strong as ligand-induced conformational distortions; thus, symmetry is not maintained. Slater and colleagues (14.4, 161, 167-159) obtained independent evidence for negative cooperativity of NAD+ binding to both the rabbit and lobster muscle enzymes by means of equilibrium dialysis and ultracentrifugation as well as by stopped-flow, fluorimetric, optical rotatory dispersion, and circular dichroism methods. Quantitative estimation of the Racker band indicated that binding of the fourth NAD' did not appear to contribute to the absorbance a t 365 nm. It has since been shown (160-162) that this effect is produced either with enzyme in which the active site thiol is partly oxidized or by an insufficient range of titration with NAD'. Heterogeneity in NAD+-binding affinity, that was in addition indicative of two pairs of equivalent and independent sites, was found for the sturgeon muscle enzyme by Seydoux et al. (160). Subsequently, Allen 156. S. F. Velick, J. P. Baggott, and J. M. Sturtevant, Biochemistry 10,779 (1971). 157. J. J. M. De Vijlder, W. Boers, A. G. Hilvers, B . J . Harmsen, and E. C. Slater, in "Pyridine Nucleotide Dependent Dehydrogenases" (H. Sund, ed.), p. 233. Springer-Verlag, Berlin and New York, 1970. 158. W. Boers, C. Oosthuizen, and E. C. Slater, BBA 250, 35 (1971). 159. J. J. M. De Vijlder, A. G. Hilvers, J. M. J. Van Lis, and E. C I Slater, BBA 191, 221 (1969). 160. F. Seydoux, S. Bernhard, 0. Pfenninger, M. Payne, and 0. P. Malhotra, Biochemistry 12,4290 (1973). 161. B. D. Peczon and H. 0. Spivey, Biochemistry 11,2209 (1972). 162. N. C. Price and G. K. Radda, BBA 235,27 (1971).

34

J. IEUAN HARRIS AND MICHAEL WATERS

and Harris (163) found negative cooperativity of NAD' binding to apply in the case of enzyme from a prokaryote B . stearothermophilus, showing that the phenomenon is not confined to eukaryotic cells. Moreover, as in the case of the sturgeon enzyme, the binding isotherm appeared to reflect the presence of two distinct pairs of equivalent binding sites. Yeast GAPDH is thus so far unique in possessing a concerted mechanism of coenzyme binding. I n accord with the complex sequential model, Koshland and colleagues (143,164) have reported that binding of only the first molecule of NAD' influences the reactivity of the active site Cys-149 in all four subunits. The major structural changes in the tetramer, as evidenced by thiol reactivity, appear therefore to have occurred in response to the binding of the first molecule of NAD'. This phenomenon has been observed with ORD (137) and other (165) optical techniques, as well as by difference sedimentation ( I l a ) , calorimetric (156), stopped-flow (144), and small angle X-ray scattering (166) studies. The intrinsic catalytic activity of each of the four sites remains constant with addition of each mole of NAD' when measured with substrate analogs (164), indicating that the catalytic rate is directly proportional to the amount of bound NAD' and that it is not affected by conformational changes accompanying NAD' binding to neighboring subunits. This observation suggested a possible physiological role for negative cooperativity in modulating the effects of variations in cellular NAD+ levels on the activity of the enzyme through changes in its affinity for NAD', although it has since (166) been pointed out that the high and relatively invariant levels of NAD' in the cell renders this possibility unlikely. Schlessinger and Levitzki (167)have probed further into the mechanism of negative cooperativity by using a variety of spectroscopic methods to study the interactions of rabbit muscle GAPDH with both NAD' and a fluorescent derivative of the coenzyme. Their results give added support to the concept that the structural transitions that occur when NAD' binds to the enzyme are sequential and that the largest structural change occurs in binding the first molecule of NAD'. I n addition, they show that whereas each NAD' molecule induces the same conformational change a t the nicotinamide subsite there are distinct and progressive structural changes a t the adenine subsite which are transmitted to adj acent vacant subunits thereby accounting for their decreased affinity for 163. G. A. Allen and J. I. Harris, Biochem. J. 151, 747 (1975). 164. J. Teipel and D. E. Koshland, Jr., BBA 198, 183 (1970). 165. W. Bloch, R. A. MacQuarrie, and S. A. Bernhard, JBC 246, 780 (1971). 166. I. Simon, Eur. J. Biochem. 30, 184 (1972). 167. J. Schleasinger and A. Levitzki, J M B 82,547 (1974).

1.

GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE

35

NAD’. These results offer a molecular explanation for the negative cooperativity in coenzyme binding and a t the same time for the finding that the intrinsic catalytic activity of each site is independent of NAD’ saturation (164). It was further suggested that ligands interacting a t the adenine subsite of the NAD’ binding site induce the “half of the sites reactivity” effect that has been observed with a number of alkylating reagents (see Section 1111A,2) and that both negative cooperativity in coenzyme binding and half-site reactivity result from ligand-induced conformational changes in an a priori symmetrical tetramer. 2. The Preexisting Asymmetry Model Many subunit enzymes show the phenomenon of “half-site” reactivity ; that is, the reaction with a substrate or substrate analog shows a stoichiometry equal to half the number of chemically identical subunits. A possible explanation which has been proposed for the half-site reactivity of the enzyme cytidine triosephosphate synthetase with a substrate analog (168) is that reaction with one substrate molecule induces a change in an adjacent subunit such that a second substrate molecule is prevented from reacting. Analogous studies with GAPDH showed that whereas the four active site sulfhydryl groups per tetrameric molecule react equivalently with alkylating reagents such as iodoacetic acid or iodoacetamide only two of the sites react with the pseudosubstrate p- (2-furyl)acrylolyl phosphate (FAP) unless forcing conditions are used to achieve higher stoichiometry. These observations considered in conjunction with the original crystallographic data of Watson and Banaszak (43) led Malhotra and Bernhard (169) to postulate that the chemically identical subunits in GAPDH are arranged asymmetrically and that the half-site reactivity also applied to the normal physiological substrate, diphosphoglyceric acid. Later work (69,160) has however shown that all four sites are reactive toward DPGA. One must therefore conclude that half-site reactivity is induced by certain ligands but not by others, and several possible explanations have been put forward to account for the phenomenon (cf. 168). In the case of GAPDH these may be summarized as follows: 1. 2. acyl 3.

The tetramer contains subunits of different primary structure. The active sites within the tetramer overlap so that binding of an group to one subunit prevents acylation of the adjacent subunit. The four subunits in the tetramer are identical, but acylation of

168. A. Levitrki, W. B. Stallcup, and D. E. Koshland, Jr., Biochemistry 10, 3371 (1971). 169. 0. P.Malhotra and S. A. Bernhard, JBC 243, 1243 (1968).

36

J. IEUAN HARRIS AND MICHAEL WATERS

one subunit induces conformational changes in the adjacent subunit which lowers the reactivity of that subunit toward further acylation. 4. Identical polypeptide chains form an asymmetric dimer (sad in which the conformation of the two subunits is not identical and the tetramer is a dimer of asymmetric dimers (ma1) . 5. Half-site stoichiometry of acylation is induced by bound coenzyme blocking acylation a t the R-axis-related subunit. I n considering these possible explanations, model 1 is excluded by the amino acid sequence results (cf. 36, 170) and model 5 is excluded by the finding of half-site reactivity for the muscle apoenzyme (171)and for the yeast apoensyme (113,172).Bernhard and co-workers (160, 173, 174) maintained that models 2 and 3 are excluded for several reasons. First, a number of alkylating agents, including DTNB which is considerably larger than FAP, give full-site reactivity. This tends to exclude the overlapping active site model 2. Second, iodoacetate and iodoacetamide, in addition to giving full-site reactivity, react independently a t each site, which is not in accord with the prediction of model 3. Moreover, the two unreacted sites in the diacyl enzyme can be alkylated with iodoacetate without affecting the NAD+-induced acceleration of deacylation and NAD+-induced change in the spectrum of the di (2-furylacryloyl) enzyme, implying that there is little interaction between adjacent sites of the type expected in a cooperativity model. There remains model 4, and MacQuarrie and Bernhard (176)have utilized the full-site reactivity by iodoacetamide and half-site reactivity by FAP to provide support for this model. Thus, di (2-furylacryloyl) ensyme was prepared, and the two remaining sites were blocked with iodoacetate. Acyl groups were then removed from this derivative by arsenolysis, and the resulting dialkyl enzyme was tested for stoichiometry with FAP. Only one acyl group could be incorporated into the dialkyl enzyme. This result cannot be explained in terms of an induced asymmetry model, and indeed, can only be explained by a preexisting asymmetry model if there is a subunit rearrangement. I n addition, alkylation of the enzyme with varying quantities of iodoacetate, followed by acylation of these derivatives with FAP, showed a 2: 1 ratio of alkylation to acylation, independent 170. J. I. Harris, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H.Sund, ed.), p. 57. Springer-Verlag, Berlin and New York, 1970. 171. A. Levitzki, BBRC 54, 889 (1973). 172. W. B. Stallcup and D. E. Koshland, Jr., J M B 80,77 (1973). 173. R. A. MacQuarrie and S. A. Bernhard, Biochemistry 10, 2456 (1971). 174. 0. P. Malhotra and 5. A. Bernhard, Proc. N a t . Acad. Sci. U. S. 70, 2077 ( 1973). 175. R. A. MacQuarrie and S. A. Bernhard, J M B 55,181 (1971).

1. GLYCERALDEHYDE-%PHOSPHATE

DEHYDROGENASE

37

of the degree of prior alkylation. This linear relationship is not predicted by an induced asymmetry model or an overlapping active site niodel but is consistent with the preexisting asymmetry model (174). Similar studies with alkylating and acylating reagents carried out with yeast GADPH by Stallcup and Koshland (113,172, 176) have shown that, as in the case of NAD+ binding, the fungal enzyme again differs from the muscle enzyme. Thus, for example, yeast enzyme showed only half-site reactivity toward iodoacetic acid and iodoacetamide (confirming earlier observations with the bifunctional reagent p-fluoro-rn,m'-dinitrophenylsulfone (177)) . In addition, and in marked contrast to the behavior of muscle enzyme (117, 176), a nonlinear relationship was noted between extent of alkylation and loss of enzymic activity. Similar results were obtained with a number of other alkylating and acylating agents indicating that the structure of the modifying reagent has only a secondary influence. The half of the sites' effect was therefore considered to result from ligand-induced negatively cooperative changes in the enzyme tetramer. This view was strengthened by the finding that the slow rate of alkylation of the third site in a dialkyl enzyme is the same for a reordered enzyme prepared according to MacQuarrie and Bernhard (176) as it is for a normal randomly alkylated enzyme. This showed, in accord with the negative cooperativity hypothesis, that the reactivity of the third site is a function of the number of alkyl groups bound rather than the number of available reactive sites. Moreover, relative values of two rapidly reacting groups, a third more slowly reacting, and a fourth very slowly reacting group, would necessitate three different subunit conformations in a preexisting asymmetry model which is therefore considered to be unlikely to apply in the case of the yeast enzyme. In an extension of these studies to the natural substrate, Stallcup and Koshland (172) showed further that acylation of the first two sites with DPGA decreases the rate of acylation of the third site, and that DPGA (or other acylating agents) attached to the first two sites increases the rate of deacylation of the third site through subunit interactions. These additional observations provide further support for the concept of negative cooperativity as a cause for half-site reactivity. They could also explain many of the results that have been obtained with muscle enzyme where apparent half-site stoichiometry might have resulted from rapid hydrolysis of the acyl enzyme (83). In contrast to the reactive thiol (Cys-149) groups the Lys-183 residues of yeast GAPDH were found to react independently with acylating and alkylating reagents (17 2 ) . Significantly, however, modification of the 176. W. B. Stallcup and D. E. Koshland, Jr., J M B 80,63 (1973). 177. D. Givol, FEBS ( F e d . Eur. Biochem. Soc.) Lett. 5,163 (1969).

38

J. IEUAN HARRIS AND MICHAEL WATERS

Lys-183 residues abolishes the half-site reactivity of Cys-149, so that all four thiol groups now react independently with iodoacetate or iodoacetamide. The equivalent reaction rates of the four thiol groups in the N-acetylated enzyme argues against preexisting asymmetry unless it were to be envisaged that a ligand effect could override a preexisting asymmetry. In any event it seems clear that yeast GAPDH exhibits positive cooperativity in NAD+ binding and negative cooperativity in acylation, whereas the muscle enzyme exhibits negative cooperativity in NAD+ binding and apparent preexisting asymmetry in acylation. These differences presumably reflect differences in binding sites and in subunit interactions which cannot so far be identified from the X-ray crystallographic model of the lobster muscle enzyme.

B. MECHANISM OF ACTIONOF GAPDH 1. Physiological Activity GAPDH catalyzes the reversible oxidative phosphorylation of G-3P in a reaction that couples the oxidation of an aldehyde to the synthesis of a high energy phosphate anhydride, l13-diphosph~oglycerate(cf. 178) , according to the equation G-3P

+ NAD+ + Pi

1,3-DPGA

+ NADH

The use of arsenate instead of phosphate results in the formation of l-arseno-3-phosphoglyceratewhich is rapidly and nonenzymically hydrolyzed to 3-phosphoglycerate rendering the overall reaction irreversible and thus amenable to steady-state kinetic analysis. Other aldehydes can also be oxidized giving rise to the corresponding acyl phosphates (179). GAPDH also exhibits a number of other activities which, although unphysiological, have greatly aided efforts to elucidate the mechanism of the normal physiological reaction. The nature of these other activities, such as acyl transfer and esterase activities, which have been studied extensively [principally by Park and colleagues (cf. IS)] can be explained in terms of the formation of acyl enzyme intermediates. The normal catalytic mechanism involves the formation of a covalent phosphoglyceroyl thioester, and proof for the existence of such a high energy intermediate was first obtained by Krimsky and Racker (55) who succeeded in preparing an acetyl enzyme with acetyl phosphate. This was shown to be an enzymically active intermediate since the acetyl group could 178. F. Lipmann, Advan. Eneymol. 6,231 (1946). 179. A. P. Nygaard and J. B. Sumner, PBB 39, 119 (1952).

1.

GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

39

be reduced by NADH to give acetaldehyde and NAD'. Formation of the acetyl enzyme was inhibited by iodoacetate, consistent with the proposal of a thioester linkage between the acetyl group and the protein. The reactive group involved in thioester formation was subsequently identified as Cys-149 by Harris et al. (29). The importance of thiol groups in the enzymic reaction had been revealed by the early studies of Segal and Boyer (180), who postulated that the primary step in the reaction was the formation of a thiohemiacetal intermediate. This proposal is consistent with the observation that the rate of reaction of a number of aliphatic and aromatic aldehydes with the enzyme is facilitated by electron withdrawing substituents on the aldehyde group, which implies the attack of a nucleophilic group (such as a thiol) on the carbonyl carbon (181). Moreover, the redox reaction has been shown (182) to proceed by way of a direct hydride ion transfer from the substrate to the p-4 position of the nicotinamide ring. It does not involve a reduction of the enzyme itself (183), and the B specificity of GAPDH is now known (see Section II,B,2,a) to be the result of a 180° rotation of the nicotinamide ring about the glycosidic linkage which exposes the other face of the ring for attack at C-4 (47, @). A second function of NAD+ in the reaction is unrelated to its redox role since it is required as a cofactor for most of the nonoxidative reactions involving the reactive thiol group, with the exception of the esterase activity (13, 54, 184-188). The mechanism originally postulated by Segal and Boyer (180) (and shown schematically in Fig. 11) is in accord with the above-mentioned observations, and has since received extensive support from the stoppedflow kinetic studies of Trentham and colleagues (69, 83, 116, 117, 189, 190).

This type of ping-pong mechanism (cf. 191) is contrary to the random or ordered sequential addition of reactants, with deacylation prior to dis180. H. L. Segal and P. D . Boyer, JBC 204, 265 (1953). 181. T. H. Fife, T. Rikihisa, and B. M. Benjamin, Biochemistry 10, 3875 (1971). 182. F. A . Loewus, H. R. Levy, and B. Vennesland, JBC 223, 589 (1956). 183. W. S. Allison, H. B. White, 111, and M. J. Connors, Biochemistry 10, 2290 (1971). 184. J. Harting and S. F. Velick, JBC 207, 867 (1954). 185. J. H. Park, B. P . Meriwether, P. Clodfelder, and L. W. Cunningham, JBC 236, 136 (1961). 186. E. L. Taylor, B. P. Meriwether, and J. H. Park, JBC 238, 734 (1963). 187. A . G. Hilvers and J. H. M.Weenen, BBA 58,380 (1962). 188. R. G. Duggleby and D. T . Dennis, JBC 249, 175 (1974). 189. D. R. Trentham, BJ 122, 71 (1971). 190. P. J. Harrigan and D. R. Trentham, BJ 135, 701 (1974). 191. M. Lazdunski, Progr. Bioorg. Chem. 3, 82 (1974).

40

J . IEUAN HARRIS AND MICHAEL WATERS ,S-CHOHR

E

NA1l' I-CHOHR

R

Y E'ADt

/SH

,SCOR

NAD+

,

R +H+ C O O e k -7 O R ENAD+

WAD+

SCOR

E'

where RCHO refers to GAP and RCOOPOs to DPGA

FIQ.11. Reaction mechanism of GAPDH adapted from Segal and Boyer (180).

sociation of NADH, that was postulated on the basis of steady-state studies (60, 192-194) with muscle enzyme. A recent steady-state analysis (196) carried out with the pea seed enzyme (26)) under conditions that were designed to eliminate product inhibition by DPGA is, however, entirely consistent with the ping-pong mechanism, and it seems likely that the earlier steady-state kinetic studies would have been influenced by product inhibition with NADH or DPGA as well as by nonspecific salt effects (85). Dissociation of NADH prior to phosphorolysis is a feature common to many mechanisms for GAPDH (cf. 13) since this is necessary to explain the NAD' requirement for phosphorolysis of the acyl enzyme (196). Indeed, apart from the earlier steady-state kinetic studies, the only report a t variance with this concept is that of Smith (19'7), who examined the ternary acyl enzyme-NADH complex fluorimetrically and was unable to displace the NADH even a t high concentrations of NAD'. It has, however, been suggested (117)that phosphate is required for this displacement to occur. 192. T. Keleti and J. Batke, Acta Physiol. 28, 195 (1965). 193. B. A. Orsi and W. W. Cleland, Biochemistry 11, 102 (1972). 194. C. M. Smith and S. F. Velick, JBC 247, 273 (1972). 195. R. G. Duggleby and D. T. Dennis, JBC 249,167 (1974). 196. E. Racker and I. Krimsky, Fed. Proc., Fed. Amer. SOC.Eap. B i d . 17, 1135 ( 1958). 197. T. E. Smith, Biochemistry 5,2919 (1966).

1. GLYCERALDEHYDE-%PHOSPHATE

DEHYDROGENASE

41

Trentham (116) has shown that NAD+ strongly accelerates the formation and breakdown of acyl enzyme from DPGA as well as from G-3P. Moreover, the relationship is reciprocal, since acylation of the enzyme weakens NAD+ binding (69) and thus accounts, a t least in part, for earlier discrepancies between the kinetically and fluorimetrically determined NAD' dissociation constants (60), as well as for the strong product inhibition by NADH (198). In fact, only if the holoenzyme is acylated is the dissociation of NAD+ in the reverse reaction sufficiently rapid to account for the catalytic rate. The rate-limiting step of the oxidative phosphorylation is NADH release a t high pH (>7.5) and phosphorolysis of the acyl enzyme a t low pH. This is because the rate of phosphorolysis is highly p H dependent, possibly increasing more than 2 X 104-fold from pH 5.4 to 8.6, while the rate of NADH release is independent of pH over this range, with the result that the two converge around pH 7.5. The pH dependence of phosphorolysis (190) may reflect a requirement for the phosphate trianion (PO,3-). At high enzyme concentrations (> 0.1 mg/ml) , the conversion of the predominant gem-diol form of G-3p to its reactive aldehyde form becomes rate limiting (146, 199). This in vitro interconversion does not apply in vivo since the reactive aldehyde form of G-3P is the product of both the aldolase and triosephosphate isomerase reactions, and the hydration of the aldehyde is a slow process. I n the reverse reaction, the rate determining step is a process associated with NADH binding, probably a conformational change, at high pH, and release of G-3P a t low pH (189, 1 9 9 ~ )At . high ionic strengths, acylation becomes rate limiting (83)* With the natural substrate it is now generally agreed that all four sites of the muscle enzyme tetramer are simultaneously active both in the forward and reverse reactions (160, 161, 189) despite earlier claims (169) that only the fourth site turns over. Smith and Velick (194) have undertaken an extensive steady-state kinetic analysis of forward and reverse reactions catalyzed by the liver and muscle enzymes, under pseudophysiological conditions, in a n effort 198. I. Krimsky and E. Racker, Biochemistry 2, 512 (1963). 199. D. R. Trentham, C. H. McMurray, and C. I. Pogson, BJ 114, 19 (1969). 199a. An alternative proposal now favored by Trentham which removes the necessity to postulate an NADH-induced conformational change is that aldehyde release is rate limiting under all conditions of low salt. The precursor for aldehyde release is the NAD+-aldehyde enzyme. A t high pH this complex is in rapid equilibrium with the NADH-acyl enzyme, which is the major species and therefore the predominant steady-atate intermediate. At low pH, however, the rapid equilibrium can favor the aldehyde-apoenzyme complex suggesting that NAD' dissociation from the NAD+-aldehyde enzyme is favored at low pH.

42

J. IEUAN HARRIS AND'MICHAEL WATERS

to understand the factors allowing gluconeogenesis through GAPDH in liver. Although the conclusions with regard to the forward reaction are complicated by product inhibition caused by DPGA, the results obtained for the reverse reductive dephosphorylation reaction provide an explanation for the possible metabolic significance of negative cooperativity in muscle and in liver. At the low DPGA concentrations encountered in vivo, NAD' acts cooperatively to convert the DPGA saturation curve from a sigmoidal to a hyperbolic form, thus sensitizing the enzyme to the lower concentrations of the acyl phosphate. At the same time, NAD+ acts to abolish substrate inhibition by NADH toward nonacylated enzyme sites. At higher concentrations of acyl phosphate, NAD' acts as a weak competitive inhibitor toward NADH, in contrast to the strong competitive inhibition by NADH toward NAD' seen in the forward reaction. The latter observations can be explained by the decrease in binding affinity for NAD' that occurs on acylation (GO),which Smith and Velick (194) suggested is largely the result of an isomerization process (cf. 143).Presumably, because the enzyme is almost totally acylated and because of the high NAD' concentrations, cooperative effectcl are not seen in the forward reaction in vitro; i.e., the enzyme exists in a single conformation. While there is no doubt that bound NAD+ enhances acylation (117) as well as deacylation (7O),it is difficult to reconcile the above-mentioned cooperative effects with the observation (165)that NAD' occupation of one site in the rabbit tetramer did not affect the catalytic activity of the other three sites toward aldehyde substrates. Evidence against a slow isomerization of the T + R variety, which might be expected to affect thiol reactivity in other subunits, is the fact that the sturgeon apoenzyme is almost instantaneously active in acylation, following the addition of NAD' (116). The nature of the NAD+ enhancement of acylation and deacylation is of considerable interest since it has a bearing on the mechanism of enzyme catalysis in general, and also because it explains the NAD' requirement of a number of the minor activities of GPD (see Section III,B,2). Studies with the simple alkylating agents iodoacetate and iodoacetamide have shown that NAD' promotes alkylation of Cys-149 by negatively charged iodoacetate, but inhibits alkylation by the uncharged iodoacetamide molecule (117,196).Since NADH and the adenine nucleotides do not facilitate alkylation by iodoacetate or arsenolysis of acetyl phosphate (200-202), it has been suggested that the positively charged pyridinium ring facilitates attack by negatively charged alkylating or 200. S. H. Francis, B. P. Meriwether, and J. H. Park, JBC 246, 5427 (1971). J. H. Park, JBC 248, 5433 (1971).

201. S. H. Francis, B. P. Meriwether, and 202. A. Feneslau, JBC 245, 1239 (1970).

1. GLYCERALDEHYDE-3-PHOSPHATE

DEHYDROGENASE

43

deacylating agents through ion pair formation. This "ion pair" concept is supported by the inhibitory effect of high ionic strength on both alkylation by iodoacetate and acylation by acetyl phosphate, which presumably occurs because the positive charge is masked by interaction with anions (83, 122). Cseke and Boross (122,203) have shown that the PKa of Racker band absorbance and of thiol anion carboxymethylation is lowered from about 8 in the apoenzyme to around 5.5 (204) in the presence of NAD'. This has been taken as evidence that NAD' has lowered the pK, of Cys-149 (117,203), especially since the pK, of carboxymethylation and of Racker band absorbance vary together, depending on the nature of the solvent anion. If the nucleophilicity of the thiol anion is insensitive to its basicity, as the evidence suggests, then this lowered thiol group dissociation induced by NAD' would explain the thiol group reactivity a t lower pH (< 7.0). It cannot explain the fact that the reactivity is only exhibited toward iodoacetate and not iodoacetamide, since reactivity of the thiol group should be identical toward the two alkylating agents. Consequently, it is still necessary to postulate ion pair formation in the presence of NAD'. Whatever the nature of this effect it is manifest a t low ratios of NAD+:enzyme (< 1 mole/mole) which suggests migration of NAD' from alkylated sites to nonalkylated sites induced by the lowered affinity of NAD' for alkylated subunits (117, 202). While the scheme outlined above accounts for the NAD+-dependent activation of thiol-149, it does not account for the fact that Cys-149 is highly activated compared to simple aliphatic thiols even in the absence of NAD'. It therefore becomes necessary to postulate the presence of a basic group to activate the thiol group by hydrogen bonding to its proton in a manner similar to that found with papain and thiol-subtilisin, where histidine is the basic group (83, 205, 206'). The effect of NAD' on the pK, of the thiol could then be explained by postulating that NAD' binding alters the conformational alignment of the thiol and basic residues in order to draw a proton further away from the thiol group. The inhibitory effect of anions and the anion-induced variation in the pK, of the thiol base proton would then be the result of interference in both base thiol and pyridinium thiol interactions (122).It has been suggested that a strongly basic group adjacent to the thiol base pair is responsible for anion binding and that the bound anion creates an electron-rich region which in turn increases the pK, of the thiol base pair (122, 20od). The 203. E. Cseke and L. Boross, Acta Biochim. Biophys. 2, 39 (1967). 204. M.T.A. Behme and E. H. Cordes, JBC 242,5500 (1967). 205. G. Loae, Phil. Trans. Roy. Snc. London, Ser. B 257, 237 (1970). 206. L. Polghr, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 38, 187 (1974).

44

J. IEUAN HARRIS AND MICHAEL WATERS

H<

,SCORENAD+

+

- NAD+ E

HB,

/WOR

FIO.12. Reaction mechanism of GAPDH according to Harrigan and Trentham ( 8 3 ) .

crystallographic evidence shows that Lys-191 and Arg-231 contribute toward the formation of this site which appears to be suitably located to participate in binding inorganic phosphate during the phosphorolysis reaction. These two residues are highly conserved (Table 11), and modification of Lys-191 is known to cause inactivation ( 7 6 ) . Presumably the powerfully inhibitory analog, threose 2,4-diphosphate, which forms a stable acyl enzyme even in the presence of phosphate (207),also binds a t this site. A second anion binding site created primarily by the nicotinamide ribose and located where the phosphate group of the substrate (G-3P) would be expected to bind (46, 47) might also account for substrate inhibition by phosphate (60) and for the greater affinity of the enzyme for G-3P as compared to glyceraldehyde (179). I n the crystallographic model the highly conserved His-176 occurs within hydrogen bonding distance of Cys-149 (cf. Section II,B,4,d). It is thus ideally placed to be the histidine residue that interacts with the active site thiol group during catalysis. It may also be envisaged to be the residue that extracts a proton from Cys-149 during the initial acylation step according to the reaction scheme postulated by Harrigan and Trentham (83) and illustrated schematically in Fig. 12. The role of histidine in deacylation but not in the redox reaction has also been discussed by Francis et a2. (90). 2. Other Activities of GAPDH I n addition to its dehydrogenase activity GAPDH possesses several other activities. These have been reviewed extensively by Colowick et a2. (IS) and will be referred to here only insofar as they help to clarify the mechanism of the dehydrogenase reaction. a. Acyltransferase Activity. GAPDH catalyzes the exchange of into acyl phosphates, such as acetyl phosphate and DPGA, and arsenate 207. E. Racker, V. Klybas, and M. Schramm, JBC 234, 2610 (1969).

1. GLYCERALDEHYDE-%PHOSPHATE

DEHYDROGENASE

45

will induce an irreversible arsenolysis of acyl phosphate in the presence of the enzyme (184,208). CHaCOOPOsHz CH&OOPOaHz

+ Ha*'POc % CHSCOO''PO~HZ+ HaPo,

+ HaAsOi % CH~COOASOIHZ+ Hap04 1Hz0 CHaCOOH + HshO4

It has been suggested (70)that NAD+ acts by facilitating deacylation of the thiol ester, and the later kinetic studies (189) are in agreement with this view. Only NAD+ analogs which are active in the oxidative phosphorylation reaction will substitute for NAD' in the enzyme-catalyzed arsenolysis of DPGA (188) or acetyl phosphate (90). At higher pH (8.5) an irreversible S to N transfer of acetyl groups occurs. b. Esterase Activity. NAD+-free GAPDH catalyzes the hydrolysis of aryl esters (e.g., p-nitrophenyl acetate) a t rates up to five times that of chymotrypsin (186). This reaction also proceeds via a thiolester intermediate with Cys-149 (29) and is, therefore, inhibited by iodoacetate. RCOOAr

+ ESH

ES

-+

COR

Ha0

RCOOH

ArOH

The rate of the reaction is related to the electron withdrawing ability of the phenolic moiety (81) and may be compared to the reaction of aromatic aldehydes with the enzyme (181). These reactions are in accord with the concept of a nucleophilic attack of the thiol of the carbonyl carbon. Although NAD' inhibits the esterase reaction, adenine nucleotides stimulate it, which suggests that NAD+ is blocking access of the aromatic ester to the reactive thiol in a manner similar to its effect on iodoacetamide alkylation (7'0). Acyl shifts involving Cys-149 and Lys-183 also occur readily with p-nitrophenyl acetate a t higher pH and in the absence of NAD' (70,73, 74).

C. METABOLIC ROLEOF GAPDH GAPDH is common to both the glycolytic and gluconeogenic pathways. The muscle and liver enzymes are similar in structure and properties (194, 209), and the different behavior of the enzyme in muscle and liver must therefore be ascribed to differences in the cellular environment. 208. P. Oesper, JBC 201,421 (1954). 209. J. M. Lamhert and R. N. Perham, FEBS ( F e d . Eur. Biochem. Soc.) Lett 40, 305 (1974).

46

J . IEUAN HARRIS AND MICHAEL WATERS

Evidence has been obtained (210, 211) that the rate of glycolysis in rat liver is proportional to the cytosolic redox state (i.e., the ratio of free XAD': NADH) . It was also inferred that the rate of gluconeogenesis is inversely proportional to (but not necessarily controlled by) this ratio. These conclusions were drawn from studies utilizing the changes in cytosolic NAD': NADH ratios induced by different dietary and hormonal states and presumably reflect the ability of GAPDH to influence the rate of glycolysis by responding to changes in the redox state. Moreover, it has been suggested (211) that the primary effect derives from an alteration in the phosphorylation state of the adenine nucleotide pool since the NAD+:NADH ratio is coupled to the ATP/ADP X Pi ratio through the phosphoglycerate kinase reaction "AD+] [NADH]

1

=-x K

[3-phosphoglycerate] [ATP] X[GAP1 [ADPI[Pil

Thus, a high NAD+:NADH ratio will be associated with a high ATP:ADP X P, ratio, and with an increased glycolytic flux. Conversely, a decrease in the NAD+:NADH ratio will be accompanied by a fall of the ADP:ADP X Pi ratio and, paradoxically, a rise in the flux in the direction of gluconeogenesis. That is, while the overall NAD': NADH ratio is set by the redox potentials of the cytosolic dehydrogenases at that pH, it is possible to vary this ratio and hence the direction of carbon flow through GAPDH by changes in metabolite concentrations. The major change in the ATP:ADP X Pi ratio results from raised ADP and Pi levels, although an increase in the latter would to some extent counter the increased gluconeogenic flow by a substrate effect on GAPDH (212). The percentage fall in ATP level is only slight in starvation and alloxan diabetes although the percentage rise in ADP is considerable (210,213). The NAD+:NADH ratio in the cytoplasm can, under some conditions (e.g., in starvation), be considerably lower in liver than in muscle (212, 214), giving rise to conditions more favorable for gluconeogenesis. I n addition, oleate infusion was found (216)to promote gluconeogenesis in rat liver, possibly as the result of a lowered NAD+:NADH ratio; and sharply lowered ratios were also found in the livers of alloxan diabetic 210. R. L. Veech, L. Raijman, and H. A. Krebs, BJ 117,499 (1970). 211. K. A. Gumaa,. P. McLean, and A. L. Greenbaum, Essays Biochem. 7, 39 (1971). 212. D. H. Williamson, P. Lund, and H. A. Krebs, BJ 103, 514 (1967). 213. J. Elliott, E. Dade, D. M. W. Salmon, and D. A. Hems, BBA 343,307 (1974). 214. L. A. Jedeikin and S. Weinhouse, JBC 213,271 (1955). 215. J. R. Williamson, JBC 242,4476 (1967).

1. GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE

47

rats (211,216), in accord with the increased gluconeogenesis characteristic of diabetes. These considerations rest heavily on the assumption that GAPDH is catalyzing an equilibrium reaction, and that it does not fulfill a regulatory function dependent upon inhibitory effectors. While this assumption may hold true in liver under normal circumstances ( H I ) , there is evidence to suggest that this is not always the case. Thus, for example, of eight glycolytic enzymes studied in adipose tissue taken from alloxan diabetic rats (217), the GAPDH-phosphoglycerate kinase system alone did not move in parallel with the other enzymes studied, and was the slowest to be restored with insulin. GAPDH was also found (218, 219) to be considerably displaced from its equilibium value in mouse brain and rat heart. Cross-over studies (216, 219) have demonstrated that GAPDH becomes the rate-limiting (and possibly regulatory) gluconeogenic step during pyruvate and oleate infusion of rat livers, and there is good evidence (220) that the enzyme can be rate-limiting in ascites cells. It has also been suggested (221) that GAPDH in cerebral cortex slices may form a regulatory system with pyruvate kinase, similar to the hexokinase-phosphofructokinase system in that ATP resulting from the action of GAPDH and phosphoglycerate kinase might be expected to inhibit pyruvate kinase leading to an accumulation of DPGA and the consequent product inhibition of GAPDH. The activities of glycolytic and gluconeogenic enzymes have been determined for most rat and human tissues (222). The values for GAPDH (Table V) show that in liver it is 50 times higher than the activity of the least active enzyme, glucokinase; while in muscle it is over 300 times greater than the enzyme of lowest activity, hexokinase. For adipose tissue, on the other hand, total GAPDH activity is only marginally higher than the activity of the least active enzyme, phosphofructokinase, suggesting that in this tissue (although presumably not in heart or brain where GAPDH activity is about 20 times greater than the activity of the rate-limiting enzyme) GAPDH might be expected to be a rate-limiting or regulatory enzyme. In assessing these possibilities it should nevertheless be recalled that the effective GAPDH activity in the cell is controlled by a variety of factors. Thus, in the first place, a t the normal 216. 217. 218. 219. 220.

H. A . Krebs, Recent Ad7lan. Enzyme Regul. 5,409 (1967). E. D. Saggerson and A. L. Greenbaum, BJ 115, 405 (1969). 0. H. Lowry and J. V. Passoneau, JRC 239, 31 (1964). J. R. Williamson, JBC 240, 2308 (1965). D. P. Kosow and I. A . Rose, Fed. Proc., F e d . Amer. SOC.Exp. Biol. 31, 1219

(abstr.) (1972). 221. F. S. Rolleston and E. A. Newsholme, BJ 104,524 (1967). 222. C. E. Shonk and G. E. Boxer, Cancer Res. 24,709 (1964).

48

J. IEUAN HARRIS AND MICHAEL WATERS

TABLE V MAXIMAL ACTIVITIES OF GAPDH I N RAT AND HUMAN TISSUES' Tissue

Liver

Kidney

Brain

Heart muscle

Skeletal muscle

Spleen

Adipose

Rat 63 f 2.36 69 f 6.5 81 f 10.5 90 f 7.6 294 f 36 25 f 5.7 0.06 f 0.16 Human 65 63 9 61 284 2.1 a

Micromoles per minute per gram wet tissue It S.E.M.

physiological pH the enzyme is operating at only about 20% of its maximum activity (92s). Moreover, this falls to as low as 5% a t p H 6.5, a pH that might well be attained during peak production of lactate in muscle (224, 225). GAPDH is also inhibited by various adenine nucleotides (108, 900,224-926) although, in vivo, ATP is the only nucleotide normally present at sufficiently high concentrations to cause significant inactivation. Thus, a t normal physiological concentrations of ATP GAPDH could be inhibited to the extent of 67% a t pH 7.4 and of 87% a t pH 6.8 (225). I n muscle phosphocreatine could also be an effective inhibitor of GAPDH (224). I n the erythrocyte GAPDH is operating a t less than 1% of its maximum capacity (227) because of product inhibition by NADH and a high K , for phosphate. The enzyme is also susceptible to appreciable product inhibition of the forward reaction by DPGA (96,60),which could account for its inhibition following pyruvate infusion of rat hearts (819). It has also been suggested (228, 229) that binding of GAPDH to erythrocyte membranes and to F-actin could be a means of regulating the activity of the enzyme. Consideration of these many factors illustrates the GAPDH could well play an important regulatory role in glycolysis despite its apparent high in vitro activity. Studies with the purified enzyme have, in addition, led to some interesting suggestions concerning its possible in vivo behavior. Thus, isolation of acyl enzyme from muscle led to the proposal (165) that scyl enzyme itself is a glycolytic intermediate of considerable importance since the active site concentration of GAPDH in muscle is in excess of 70 pill (60). Moreover, under conditions of sudden glycolytic flow (as 223. G. T. Cori, M. W. Slein, and C. F. Cori, JBC 173, 605 (1948). 224. M. Oguchi, E. Gerth, B. Fitzgerald, and J. H. Park, JBC 248,5562 (1973). 225. M. Oguchi, B. P. Meriwether, and J. H. Park, JBC 248, 5562 (1973). 226. N. K. Nagradova, M. K. Vorona, and R. A. Asriyants, Biokhimiya 34, 503 (1969). 227. G. C. Mills and F. L. Hill, ABB 146, 306 (1971). 228. H. Arnold and D. Pette, Eur. J . Biochem. 6,163 (1968). 229. B. C. Shin and K. L. Carraway, JBC 248, 1436 (1973).

1.

GLYCERALDEHYDE-%PHOSPHATE DEHYDROGENASE

49

in muscle contraction), acyl enzyme might act to “buffer” glycolytic flow through the triose phosphates below GAPDH until phosphofructokinase responded to the altered adenine nucleotide ratio. It has also been suggested (166, 194) that deacylation of GPD under such conditions would enhance the bind of NAD’ (since NAD’ does not bind as strongly to the acyl enzyme) and thereby favor the forward reaction. Concomitantly, product inhibition by DPGA would be relieved by ATP synthesis a t the phosphoglycerate kinase step, in response to the lowered cytosolic ATP:ADP X Pi ratio. In addition, the lowered intracellular p H resulting from lactate production would increase the NAD+:NADHratio, and this would also favor glycolysis by its effect on the GAPDH reaction. In the liver, no such glycolytic transients occur; thus, the enzyme would be permanently acylated, thereby favoring NADH binding and the reverse reaction, especially in the more reduced hepatocyte (214). The presence of NAD’ would serve to activate the reverse reaction a t low DPGA concentrations, while the higher level of NADH would induce positive cooperativity of DPGA binding (194). These considerations do not take into account the major glycolytic controls exerted by the hexokinase-phosphofructokinase system (230) but they do show that G3PDH could act as a regulatory enzyme in response to the NAD+:NADHand ATP:ADP x Pi ratios in the cell. These ratios in conjunction with appropriate substrate (i.e., G-3P, DPGA, and Pi) concentrations prime the enzyme for glycolysis or gluconeogenesis in accord with the particular environment and needs of the cell.

ACKNOWLEDGMENTS The authors are grateful to Dr. M. G . Rossmann for providing original copies of Figs. 2-8 and to Dn. D. R. Trentham and J. Armstrong for reading parts of the manuscript and for helpful comments.

230. H. A. Krebs, Essays Biochem. 8, 1 (1972).

This Page Intentionally Left Blank

Nicotinamide Nucleotide Transhydroenases J . RYDSTROM

J . B. HOEK

L . ERNSTER

I . Definitions . . . . . . . . . . . . . . . . . I1. BB-Specific Transhydrogenases . . . . . . . . . . . A . Historical . . . . . . . . . . . . . . . B . Occurrence . . . . . . . . . . . . . . . C . Purification and Assay . . . . . . . . . . . D . Molecular Properties . . . . . . . . . . . . E . Reaction Mechanism and Regulation . . . . . . . I11. AB-Specific Transhydrogenaaes . . . . . . . . . . . A . Historical . . . . . . . . . . . . . . . B . Occurrence . . . . . . . . . . . . . . . C . Preparations and Assay . . . . . . . . . . . D . Molecular Properties . . . . . . . . . . . . E . Relationship to the Energy-Coupling System . . . . . F. Kinetics and Reaction Mechanism . . . . . . . . G . Reconstitution . . . . . . . . . . . . . I V . Physiological Roles of Nicotinamide Nucleotide Transhydrogenaaes . A . Redox State of Mitochondrial Nicotinamide Nucleotides . B. Role of Transhydrogenme in Mitochondrial Monooxygenation Reactions . . . . . . . . . . . . . C. Role of Transhydrogenme in Mitochondrial Glutamate and Isocitrate Metabolism . . . . . . . . . . . D . Role of Transhydrogenase in Fatty Acid Synthesis . . .

51 52 52 53 54 57 59 62 62 64 66 69 71 75 78 79 81 83 85 88

.

1 Definitions

The term “nicotinamide nucleotide transhydrogenase” (EC 1.6.1.1) is used in this chapter to denote those enzymes that catalyze the reversible 51

52

J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

transfer of hydrogen between the two naturally occurring nicotinamide nucleotides, NAD ( H ) and NADP(H) ,i.e., the reaction NADH

+ NADP+

NAD+

+ NADPH

without the mediation of any further substrates. Thus, the term does not include those enzymes that catalyze transhydrogenation between only one naturally occurring nicotinamide nucleotide and an artificial nicotinamide nucleotide analog [most NAD (H)- or NADP (H)-specific flavoproteins belong in this category] ; nor does it include so-called dual-specific dehydrogenases, i.e., enzymes that react with both NAD(H) and NADP(H) , but where a transhydrogenation between the two requires the mediation of a third substrate (e.g., glutamate dehydrogenase) . Finally, the present chapter does not deal with the enzyme DT-diaphorase (EC 1.6.99.2), a flavoprotein that oxidizes both NADH and NADPH but does not seem to catalyze a transhydrogenation between the two. Nicotinamide nucleotide transhydrogenases may be divided into two classes. One class is present in certain bacteria, and possibly in some plants, is an easily extractable, water-soluble enzyme ; is not functionally linked to the energy-transfer system of the bacterial membrane; is a flavoprotein; and is specific for the 4B-hydrogen atom of both NADH and NADPH. The other class is present in both certain bacteria and in mitochondria ; is a firmly membrane-bound water-insoluble enzyme; is functionally linked to the energy-transfer system of the bacterial or mitochondria1 membrane; is not known to be a flavoprotein; and is specific for the 4A-hydrogen atom of NADH and the 4B-hydrogen atom of NADPH. For the sake of convenience, the two classes of enzyme will be referred to below as BB-specific and AB-specific transhydrogenases, respectively. II. BB-Specific Transhydrogenases

A. HISTORICAL Investigations by Colowick et al. (1) on isocitrate dehydrogenase in Pseudomoms fluorescens led to the discovery that in the presence of extracts of these bacteria NAD' could be reduced by isocitrate provided a catalytic amount of NADP' was added. It was proposed that a specific enzyme, called pyridine nucleotide transhydrogenase, catalyzed the for1. S. P. Colowick, N. 0. Kaplan, E. F. Neufeld, and M. M. Ciotti, JBC 195, 95 (1952).

2.

NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

53

mation of NADP+ plus NADH from NAD+ plus NADPH. Determination of the stoichiometry of oxidized and reduced nicotinamide nucleotides ( g ) ,respectively, indicated the net reaction NAD+

+ NADPH + NADH + NADP+

Indirect transfer of hydrogen ( I ) , transfer of phosphate (2, S) or nicotinamide moieties ( 4 ) could be eliminated, suggesting that reduction of NAD’ by NADPH was direct and stereospecific (6). Regulation of the reduction of NADP’ by NADH by various nucleotides (3)indicated that the transhydrogenase was an allosteric enzyme. However, despite extensive kinetic studies of the transhydrogenase carried out mainly by Kaplan and co-workers ( 6 ) with a purified enzyme, its detailed reaction mechanism and regulation are still unclear. The stereospecificity of the Pseudomonas enzyme for the 4B hydrogen of both NADH and NADPH (7), in contrast to those of the mammalian and certain bacterial transhydrogenases which are AB specific, led Hoek et al. (8) to propose that the so-called BB- and AB-specific transhydrogenases represent two different classes of enzymes (see Section I). Several other laboratories have been involved in elucidating the properties of BB-specific transhydrogenases from other sources, e.g., the laboratories of Veeger (9) and Chung ( I O ) , who found that the transhydrogenase from Azotobacter vinelandii is closely related to the Pseudomonaa enzyme. B. OCCURRENCE Nicotinamide nucleotide transhydrogenase was originally discovered in Pseudomonas fluorescens. Part of the work done with these bacteria by Kaplan and co-workers (see 7) appeared later to have involved Pseudomonas aeruginosa. There is little doubt, however, that these two strains contain transhydrogenases that are closely related. Kaplan and co-workers (3) also demonstrated the presence of transhydrogenase in 2. N. 0. Kaplan, S. P. Colowick, and E. F. Neufeld, JBC 199, 107 (1952). 3. N . 0. Kaplan, S. P. Colowick, E. F. Neufeld, and M. M. Ciotti, JBC 205, 17 (1953). 4. N. 0. Kaplan, S. P. Colowick, L. J. Zatman, and M. M. Ciotti, JBC 205, 31 (1953). 5. A. San Pietro, N. 0. Kaplan, and S. P. Colowick, JBC 212, 941 (1955). 6. P. T. Cohen and N . 0. Kaplan, JBC 245, 4666 (1970). 7. D. D. Louie and N . 0. Kaplan, JBC 245,5691 (1970). 8. J. B. Hoek, J. Rydstrom, and B. Hojeberg, BBA 333,237 (1974). 9. H. W. J. van den Broek and C. Veeger, “Pyridine Nucleotide Dependent Dehydrogenases,” p. 335. Springer-Verlag, Berlin and New York, 1969. 10. A. E. Chung, J. Bucteriol. 102, 438 (1970).

54

J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

Azotobacter vinelandii, Azotobacter chroococcurn, and Azotobacter agile. The occurrence of transhydrogenases in plants, i.e., spinach, pea leaves, turnip greens, parsley, watercress, and Euglena gracilis, was reported by Keister et al. (11) ; among these the spinach enzyme appears to be identical with ferredoxin-NADP reductase (19, IS). The same holds for the transhydrogenase obtained from the alga Burnilleriopsis filifomzis (14) 1 6 ) . Whether these plant and alga transhydrogenases may be classified as BB- or AB-specific eneymes remains to be established, although their kinetic properties partly resemble those of the Pseudomonas enzyme. Keister and Hemmes (16) isolated a transhydrogenase from Chromatiurn, with properties similar to those of the Azotobacter enzyme.

C. PURIFICATION AND ASSAY The first demonstration of transhydrogenase activity in Pseudornonas jluorescem by Colowick et al. (1) was carried out with a crude extract obtained from cells grown on citrate as the sole carbon source. This extract could be fractionated further by acetone precipitation followed by calcium phosphate adsorption and subsequent elution with potassium phosphate. A second acetone fractionation and calcium phosphate adsorption gave a total purification of about 200-fold. This preparation was devoid of dehydrogenase activity using glutamate, isocitrate, lactate, 6-phosphogluconate, glucose 6-phosphate, and ethanol as substrates in combination with either NAD+ or NADP+.A considerably more elaborate and extensive purification was reported by Cohen and Kaplan ( l 7 ) ,using a Pseudomonas aeruginosa strain grown on glucose medium. After sonic disruption of the cells and centrifugation, the supernatant was fractionated with ammonium sulfate. The sediment was redissolved and chromatographed in two steps on DE-11 and hydroxylapatite columns. Finally, a two-step differential centrifugation gave a total purification of about 800-fold, yielding a homogeneous preparation that could be crystallized and that exhibited no dehydrogenase activity with a variety of substrates tested. Spinach transhydrogenase was isolated by Keister et al. (11) by acetone extraction and precipitation with protamine sulfate followed by adsorp11. D. L. Keister, A. San Pietro, and F. E. Stolzenbach, JBC 235, 2989 (1960). 12. M. Shin, K. Tagawa, and D. Arnon, Biochem. Z.338,84(1963). 13. W. W. Fredericks rind J. M. Gehl, JBC 246, 1201 (1971). 14. P. Boger, Plantn 92, 105 (1970). 15. P. Boger, Proc. Znt. Congr. Photosyn. Res., 2nd, 1971 p. 449 (1972). 16. D. L. Keister and R. B. Hemmes, JBC 241,2820 (1966). 17. P. T. Cohen and N. 0. Kaplan, JBC 245,2825 (1970).

2.

NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

55

tion on bentonite. This preparation could be purified further on Dowex 50W,giving a total purification of 330-fold. The preparation was reported to exhibit a number of other activities, e.g., NADPH-menadione reductase and NADPH-cytochrome c reductase. Shin et al. (12) extended this purification by means of repeated DEAE and Sephadex chromatography steps. A final ammonium sulfate precipitation yielded a crystalline preparation whose activity was not much different from that of the enzyme prepared by Keister et al. (11). The Chromatium transhydrogenase was isolated by Keister and Hemmes (16) from a sonicated extract of Chromatium strain D. After high-speed centrifugation and ammonium sulfate precipitation, two subsequent steps of DEAE-cellulose chromatography were employed. The purity of the preparation was tested for various reductase.and dehydrogenase activities. The Azotobacter vinelandii transhydrogenase has been purified by two different laboratories. Chung (10) described a method that employed heat treatment and alternating DEAE-cellulose and calcium phosphate gel chromatography. The purity of the preparation was not stated in terms of contaminating activities. This procedure was later modified by Middleditch et al. (18) to include gel filtration and sucrose gradient fractionation. A purification of about 900-fold was achieved. Information regarding contaminating activities in this preparation was not given, but the transhydrogenase sedimented as a single band in the ultracentrifuge. Similarly to Chung (lo), van den Broek et al. (19) employed heat treatment that precipitated a considerable amount of contaminating protein. Fractionation with ammonium sulfate, calcium phosphate gel, and ammonium sulfate, in that order, prior to two sequential differential centrifugation steps, gave about 650-800-fold purification. It appears that this preparation was not completely devoid of contaminating activities such as pyruvate dehydrogenase and transacetylase ; these activities could not be separated from the transhydrogenase activity by ultracentrifugation. The properties of the various preparations obtained from Pseudomoms fluorescem, spinach, Pseudomonas aeruginosa, Azotobacter vinelandii, and Chromatium, respectively, are summarized in Table I. Reduction of NAD' by NADPH or reduction of NADP' by NADH cannot be assayed directly since the spectral differences between NADH and NADPH or between NAD' and NADP' are negligible. Either reaction may be measured by removing aliquots from the reaction mixture 18. L. E. Middleditch, R. W. Atchison, and A. E. Chung, JBC 247, 6802 (1972). 19. H. W. J. van den Broek, J. S. Santema, J. H. Wassink, and C. Veeger, Eur. J. Biochem. 24, 31 (1971).

56

J. RYDSTR~M, J . B. HOEK, AND L. ERNSTER

TABLE I PREPARATIONS OF BB-SPECIFIC TRANSHY DROQENASE

Purification (-fold)

Specific activity (pmoles/ min/mg protein)

Pseudomanas JEuoreacens

200

11.5"

Spinach

300

Chrmatium

275

24"

7

Pseudomonas aeruginosa Azotobacter vinelandii Azolobaeter vinelandii Azotobacter vinelandii

800

217O

900

3000

Source

1.35"

650-800 220-260"

900

362"

Re covery (%)

FROM

VARIOUSSOURCES

Contaminations

Ref. 1

15

Devoid of various reductase and dehydrogenase activities tested Several, e.g., NADPH-menadione reductase and NADPH-cytochrome c reductaseb Devoid of various reductase and dehydrogenase activities tested None (crystalline)

14

Not stated

lo

15

Pyruvate dehydrogenase and transacetylase Not stated

19

16

35

4.5

11,18

16

17

18

Substrates used were NADPH plus NAD+.

* These activities may be catalyzed by the purified enzyme since the ,preparation was crystalline. Substrates used were NADPH plus thio-NAD+.

and terminating the reaction by the addition of either acid or alkali (20). I n the former case only oxidized nicotinamide nucleotides remain intact, whereas in the latter case only reduced nicotinamide nucleotides remain intact. After neutralization, the nicotinamide nucleotides can be determined enzymically. An alternative method is to keep the concentration of one of the substrates constant by means of a suitable enzymic regenerating system and to measure the change in concentration of the other substrate a t 340 nm. This type of assay was used originally by Colowick et al. (1) and later by Chung (10) and van den Broek et al. (19). The advantage with the regenerating system assay is that the reaction may be followed directly in a spectrophotometer, provided that the specificity as well as the capacity of the regenerating system is sufficiently high; 20. M. Klingenberg and W . Slenczka, Biochem. 2.331,486 (1959).

2.

NICOTINAMIDE NUCLEQTIDE TRANSHYDROGENASES

57

i.e., suitably, the maximal rate of the regenerating system should exceed that of the transhydrogenase reaction by a t least 10 times. I n order to measure transhydrogenation spectrophotometrically without a regenerating system, coenzyme analogs, e.g., thio-NAD (P) (absorption maximum for the reduced form a t 400 nm), have often been used (6, 90). However, coenzyme analogs are not natural substrates and therefore care should be taken in interpreting kinetic data obtained with these artificial substrates. On the other hand, the use of analogs appears to be the method of choice in cases where rapid reactions are measured or where low concentrations of products interfere (see 21).

D. MOLECULAR PROPERTIES Transhydrogenase from Pseudomonas aeruginma purified by Cohen and Kaplan (17) was reported to have a molecular weight of several million (21,92). Its structure, as revealed by electron microscopy, appeared like unbranched filamentous aggregates of nonuniform length in the range of 500-5000 A, and the widths of the majority of the filamentous helixes had an apparent dimension of 80-100 A ( 1 7 ) . In the presence of 2’-AMP (or NADP’) these structures were found to dissociate into smaller fragments of about 900,000 daltons, composed of some 20 subunits of 40,000-45,OOo daltons (81). The subunits were arranged in a cylindrical fashion with six or eight larger units peripheral to the fragment (91). Similarly, van den Broek et al. (19) found that, in the absence of NADP’, the purified Azotobacter transhydrogenase forms long helicallike structures approximately 120 A in width and 15,000-18,000 A in length. Upon addition of NADP+ the aggregates were transformed into shorter fragments with a minimum molecular weight of about 58,000. These data were later confirmed by Middleditch et al. (18) who also determined the amino acid composition of the Azotobacter transhydrogenase. It is still not clear, however, whether nucleotide-dependent transformations similar to those described by van den Broek et al. (19) occur in vivo. The kinetic properties (see Section II,E) and spectral characteristics (7, 9, 10, 17, 19) of the BB-specific transhydrogenases strongly suggest that these enzymes are flavoproteins. Consistent with this suggestion, Louie and Kaplan ( 7 ) showed that, in the presence of 1 M urea, 3H was taken up from the medium and was incorporated into the reduced nicotinamide nucleotide. Direct proof for the occurrence of FAD in both the 21. D. D. Louie, N. 0. Kaplan, and J. D. Lean, J M B 70,651 (1972). 22. D. D. Louie and N . 0. Kaplan, “Pyridine Nucleotide Dependent Dehydroge-

rimes," p. 351. Springer-Verlag, Berlin and New York, 1969.

58

J.

RYDSTRBM, J .

B. HOEK, AND L. ERNSTER

Pseudomonas and Azotobacter transhydrogenases was provided by Cohen and Kaplan (17) and by van den Broek et al. (19), respectively, who showed that inactivation by heat treatment could be reversed by addition of FAD. FAD could not be replaced by FMN. Reduction of the enzyme with either NADH or NADPH largely increased the heat sensitivity, whereas oxidized nicotinamide nucleotides or FAD had the opposite effect (17, 19). The number of flavins per 50,000-dalton molecular weight was calculated to be 0.58 to 1.1 ( 1 7 ) . Sulfhydryl agents, e.g., p-hydroxymercuribenzoate, both inactivate and activate the Pseudomoms enzyme, depending on the presence of oxidized and reduced substrates, respectively ( 1 7 ) . Inactivated protein may be reactivated by mercaptoethanol suggesting that p-hydroxymercuribenzoate acts on sulfhydryl groups near or a t the active site. Reversible effects of sulfhydryl agents were also observed with the Chromatium enzyme ( 1 6 ) . Proteolytic enzymes such as trypsin did not inactivate the Pseudomonas transhydrogenase (8). Transhydrogenase obtained from Pseudomonas (2, 17) and Azotobacter (9, 10, 19) catalyzes a reversible reduction of NAD+ by NADPH, which is specific for the 4B hydrogen of N A D ( P ) H (6, 7, 18); both enzymes show a broad pH optimum with a maximum around 7 (1, 19). The degree of reversibility appears to vary depending on the source of the enzyme. The reaction catalyzed by the Pseudomonus enzyme is less readily reversible than that catalyzed by the Azotobacter enzyme, although this discrepancy may be diminished by specific activators. Thus, Kaplan et al. (3,21, 22) found that 2'-AMP and other nucleotides were specific and efficient allosteric activators of both the Pseudomoms and Azotobacter transhydrogenases ; compounds lacking a ribose 2-phosphate substituent did not activate (21,22). I n the presence of activators, high concentrations of NADP+ or PI were inhibitory, suggesting a competitive relationship between these inhibiting and activating agents. Rydstrom et al. (23) demonstrated that reduction of NADP' by NADH, catalyzed by a partially purified Pseudomonus transhydrogenase, was specifically activated by Ca2+even in the absence of 2'-AMP and analogous compounds. They found, in addition, that the specificity for the activating nucleotide was altered in the presence of low concentrations of Ca2+.A similar Caz+-dependentstimulation of the Azotobacter transhydrogenase is indicated by data reported by van den Broek and Veeger (9) and by van den Broek et al. (19). I n detailed investigations by van den Broek et al. (9,19) and by Chung (lo), using highly purified transhydrogenase preparations, the Azotobacter enzyme, in contrast to 23. J. Rydstrom, J. B. Hoek, and B. Hojeberg, BBRC 52,421 (1973).

2.

NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

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the Pseudomoms enzyme, was found to be activated by 2'-AMP only under certain conditions. Other reactions catalyzed by the Pseudomonas enzyme are transfer of hydrogen from NADPH to thio-NADP', thio-NAD+, pyridine-aldehyde-NAD+, deamino-NAD', acetyl-pyridine-NAD+, ethylpyridylketone, NMN (17, 24) and 3'-NADP+ (25, 26a). These reactions are activated by 2'-AMP provided that the concentration of NADPH is not saturating. However, when NADH serves as hydrogen donor 2'-AMP has to be present as activator. I n the case of the Azotobacter transhydrogenase thioNAD' and thio-NADP', but not acetyl-pyridine-NAD+ and pyridineNAD' (19, 26), may be used as hydrogen acceptors. Exchange of NADPH for NADH as hydrogen donor does not alter substantially the specificity for the hydrogen acceptor. Reactions catalyzed by transhydrogenases from Chromatium (16) and spinach (11) appear to be poorly reversible and 2'-AMP has only inhibitory effects in these cases. The stereospecificities and the sensitivities to Ca2+have not been tested with the Chromatium and spinach transhydrogenases. Likewise, the substrate specificities of these transhydrogenases were not investigated in detail, but appear to resemble those of the Pseudomoms enzyme (11, 16). The transhydrogenase activity of spinach ferredoxin-NADP reductase is subject to regulation by ferredoxin (13) which, depending on pH, affects both maximal velocity and affinities for the substrates. I n addition to transfer of hydrogen between various nicotinamide nucleotides, the transhydrogenases from Pseudomoms (17), spinach (11, IS), and Azotobacter (9, 19) also catalyze a diaphorase reaction, using either NADH or NADPH plus an artificial acceptor, e.g., potassium ferricyanide or dichlorophenolindophenol. As expected, 2'-AMP stimulates the NADH-linked diaphorase reaction catalyzed by the Pseudomonas enzyme ( 1 7 ) .

E. REACTION MECHANISM AND REGULATION The reaction mechanism of the Pseudornonas and Azotobacter transhydrogenases has been extensively investigated. Studies of the steady-state kinetics of Pseudomonas transhydrogenase by Cohen (97)and by Cohen 24. N. 0. Kaplan. Harvey Lect. 54, 105 (1972). 25. L. Shuster and N. 0. Kaplan, JBC 215, 183 (1955). 25a. Abbreviations as follows : thio-NAD', oxidized thionicotinamide adenine dinucleotide; thio-NADP', oxidized thionicotinamide adenine dinucleotide phosphate ; and 3'-NADP', oxidized nicotinamide adenine dinucleotide 3'-phosphate. 26. H. W. J. van den Broek, Ph.D. Thesis. H. Veenman and Zonen N. V., Wageningen, 1971.

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J. R Y D S T ~ M , J . B. HOEK, AND L. ERNSTER

and Kaplan ( 6 ) led to the proposal that the reaction proceeded by a ping-pong bi-bi mechanism. This conclusion was based on experiments with reduction of thio-NAD' by NADPH in the absence and in the presence of 2'-AMP, which gave parallel double reciprocal plots. The data presented by Cohen (27) and by Cohen and Kaplan ( 6 ) did not allow, however, an elimination of a ternary complex mechanism, with a dissociation constant for the first substrate-enzyme complex close to zero. I n fact, a ternary complex, Theorell-Chance mechanism, was later reported as an alternative mechanism (7). Similar results were obtained by van den Broek and Veeger (9, 28) and van den Broek (26) for the reduction of thio-NAD+ by NADH and NADPH, catalyzed by the Azotobacter transhydrogenase. The latter authors found, however, that inhibition of the above reactions by NADP', although complicated by substrate inhibition, appeared to give different patterns. Inhibition of the NADPHthio-NAD' reaction by NADP' was dependent on both donor and acceptor concentrations. I n the presence of high concentrations of NADPH and low concentrations of NADP+ and thio-NAD+, the inhibition was linear and noncompetitive with respect to NADPH. At increasing acceptor concentrations there appeared to be a competitive relationship between NADPH and NADP', which, at high concentrations of NADP' (60 was nonlinear and showed a second-order dependence on NADPH. Inhibition of the NADH-thio-NAD' reaction by NADP' was linear and showed a noncompetitive relationship between NAD' and NADH and a competitive relationship between NAD+ and thio-NAD'. Inhibition by NADP a t high acceptor concentrations was noncompetitive with respect to NADH, whereas lower acceptor concentrations resulted in nonlinear plots. At low NADP concentration there was an uncompetitive inhibition with respect to thio-NAD+, which, a t higher NADP' concentrations, became noncompetitive. Essentially the same inhibition patterns were obtained by Chung (10) who proposed the existence of separate hydrogen donor and hydrogen acceptor sites. On the basis of the primary plots and the inhibition patterns, van 'den Broek and Veeger (9, 28) and van den Broek (26) considered the ping-pong bi-bi mechanism unlikely and suggested an alternative ternary complex mechanism involving two donors, two acceptors, and a four-equivalent reduced state of the enzyme. Reduction of NADP+ and thio-NAD (P) was proposed to proceed according to a rapidequilibrium random bi-bi mechanism. Inhibition data on the Pseudomoms transhydrogenase reported by

a),

27. P. T. Cohen, Ph.D. Thesis, TJniversity of Michigan, Ann Arbor (University Microfilms, Ann Arbor, No. 67-16542), 1967. 28. H. W. J. van den Broek and C. Veeger, Eur. J . Biochem. 24, 72 (1971).

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Cohen (27) and by Cohen and Kaplan (6) appear to be similar to those obtained with the Azotobacter transhydrogenase. Thus, with NADH as hydrogen donor and thio-NAD+ as acceptor, inhibition by NAD' was competitive with respect to thio-NAD+ and noncompetitive with respect to NADH. As seen with the Azotobacter transhydrogenase, inhibition by NADP+ was more complicated than that by NAD'. At low concentrations NADP' appeared to be competitive with NAPH, whereas higher concentrations of NADP' gave a noncompetitive pattern. In the presence of 2'-AMP and with NADPH as donor and thio-NAD' as acceptor, inhibition by NADP' gave parabolic curves. These various types of inhibition by NADP+ were interpreted to indicate that NADP' is a less effective activator than 2'-AMP or NADPH and that NADP+ displaces either NADPH or 2'-AMP from a regulatory site. I n addition, the inhibitory effect of NADP+ was proposed by Cohn (27) and by Cohn and Kaplan ( 6 ) to depend partly on the formation of a dead-end complex with the oxidized form of the enzyme. In this connection i t should be pointed out that the pronounced activating effect of Ca2+on the reduction of NADP+ by NADH catalyzed by the Pseudomoms transhydrogenase, reported by Rydstrom et al. (B), occurred in the absence of 2'-AMP, and that saturation with Ca2+abolished completely the inhibitory action of NADP'. Moreover, in the presence of nonsaturating concentrations of Ca2+,activation by 2'-AMP or NADPH was considerably more efficient than in the absence of Ca2+.It seems likely that Ca2+may prove to be a useful tool in clarifying the mechanism of the reaction. Additional evidence for the existence of multiple binding sites, including a possible regulatory site, has been obtained from several independent experiments. In the case of the Pseudomoms transhydrogenase the parallel primary double reciprocal plots of reduction of thio-NAD' by NADPH revealed a second-order dependence on NADPH but a firstorder dependence with respect to thio-NAD+ (6, 27). Accordingly, two molecules of NADPH appeared to be bound to the enzyme in the course of the reaction, possibly one to an active site and one to a regulatory site. In the presence of 2'-AMP, reduction of thio-NAD' by NADPH showed a first-order dependence on NADPH suggesting that the hypothetical regulatory site now was occupied by 2'-AMP. The existence of a single binding site for both hydrogen donor and hydrogen acceptor is supported by experiments with 3'-NADP+ (see 8). With NADB as hydrogen donor and thio-NAD+ as acceptor, the enzyme was rather inactive in the absence of activators. Addition of 2'-AMP increased the maximal velocity, with a simultaneous pronounced increase of the affinity for NADH; the change of the affinity for thio-NAD+ was less dramatic (6, 27). After addition of 2'-AMP maximal stimulation of the velocity was

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J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

reached with a half-time of about 50 msec ( d l ) . The existence of two separate binding sites in Azotobacter transhydrogenase is strongly supported by data reported by van den Broek (26)and by van den Broek et al. (299). Spectral titrations indicated that the two NADP+-enzyme complexes, besides having different conformations (as evident from C D and ORD measurements), also have markedly different dissociation absorbing in the constants, the low affinity complex (Kd< 100 &) 300-400 nm region and the high affinity complex (& = 2-3 absorbing in the visible region (29). The thio-NADP+-enzyme complex has than the a much higher dissociation constant (&- 90 &) NADP+-enzyme complex ; this difference may provide an explanation for the weak inhibitory effect of thio-NADP+ as compared to that of NADP'. Neither thio-NADP+ nor NADP+ could be bound to the apoprotein. Boger (14) showed that the transhydrogenase activity catalyzed by ferredoxin-NADP reductase obtained from Bumilleriopsis filiformis, which is very similar to the spinach enzyme, is regulated by ferredoxin and that one common nicotinamide nucleotide binding site is involved in both the diaphorase and the transhydrogenase reactions.

a)

111. AB-Speciflc Transhydrogenares

A. HISTORICAL The discovery of nicotinamide nucleotide transhydrogenase in certain bacterial extracts by Kaplan and co-workers (1) led to a search for the enzyme in other bacteria as well as in various mammalian organs. I n 1953,Kaplan et al. (30) demonstrated the presence of transhydrogenase in beef heart homogenate. I n contrast to the soluble enzyme from Pseudom o m s aeruginosa ( 1 ) the beef heart enzyme appeared to be insoluble and firmly bound to the mitochondria1 membrane (31-35), although a partial solubilization was accomplished by treatment with digitonin, a nonionic detergent ( 3 0 ) .Furthermore, differences between the soluble and 29. H. W. J. van den Broek, J. S. Santema, and C. Veeger, Eur. J. Biochem. 24, 55 (1971). 30. N. 0. Kaplan, S. P. Colowick, and E. F. Neufeld, JBC 205, 1 (1953). 31. N. 0. Kaplan, M. N. Swartz, M. E. Frech, and M. M. Ciotti, Proc. N u t . Acad. Sci. U.S. 42, 481 (1956). 32. G. F. Humphrey, BJ 65, 546 (1957). 33. T. M. Devlin, JBC 234, 962 (1958). 34. W. W. Kielly and J. R. Bronk, JBC 230,521 (1958). 35. W. C. McMurray, G . F. Maley, and M. A. Lardy, JBC 230, 219 (1958).

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insoluble enzymes with respect to kinetic and regulatory properties (30) indicated that the enzymes were unrelated mechanistically. The first indication of the existence of an energy-linked NADP’ reduction was brought forward by Krebs (36) who found that NADPHdependent dehydrogenase reactions in pigeon heart homogenate were.sensitive to dinitrophenol. This finding formed the basis for the postulate (36) that NADP+ was reduced by an ATP-controlled flavoprotein. Klingenberg and Slenczka (20) measured the steady-state reduction levels of NAD and NADP in intact mitochondria and found that, indeed, NADP was always much more reduced than NAD. These observations led to the hypothesis (20) that mitochondria contain an asymmetric energy-linked transhydrogenase which uses oxidative energy for transfer of hydrogen from NADH to NADP’. Later findings by Estabrook and Nissley (37) and others ( 3 8 4 5 ) provided further support for this hypothesis. 36. H. A. Krebs, Bull. Johns Hopkins Hosp. 95, 34 (1954). 37. R. W. Estabrook and S. P. Nissley, in “Functionelle und morphologische Organisation der Zelle” (P. Karlson, ed.), p. 119. Springer-Verlag, Berlin and New York, 1963. 38. M. Klingenberg and P. Schollmeyer, Proc. Znt. Congr. Biochem., 6th, 1961 Vol. 5, p. 46 (1963). 39. E. C. Slater and J. M. Tager, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 97. Academic Press, New York, 1963. 40. J. M. Tager, BBA 77,258 (1963). 41. R. W. Estabrook, R. W. Hommes, and J. Gonze, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 143. Academic Press, New York, 1963. 42. J. M. Tager and E. C. Slater, BBA 77,227 (1963). 43. J. M. Tager, J. L. Howland, and E. C. Slater, BBA 77, 266 (1963). 44. M.Klingenberg, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 129. Academic Press, New York, 1963. 45. M. Klingenberg, H. van Haefen, and G. Wenske, Biochem. 2.343, 452 (1965). 46. M. Klingenberg, Biochem. Z. 343,479 (1965). 47. K. van Dam and M. F. ter Welle, Regul. Metab. Processes Mitochondria, Proc. Symp., 1966 BBA Libr., Vol. 7,p. 235 (1966). 48. K. van Dam, Ph.D. Thesis, University of Amsterdam, Jacob van Campen, Amsterdam, 1966. 49. E. J. de Haan, J. M. Tager, and E. C. Slater, BBA 131, 1 (1967). 50. S. Papa and A. Francavilla, in “Mitochondria1 Structure and Compartmentation” (E. Quagliariello et al., eds.), p. 363. Adriatica Editrice, Bari, 1967. 51. S. Papa, J. M. Tager, A. Francavilla, E. J. de Haan, and E. Quagliariello, BBA 131, 14 (1967). 52. D. G. Nicholls and P. B. Garland, BJ 114, 215 (1969). 53. J. M. Tager, S. Papa, E. J. de Haan, R. D’Aloya, and E. Quagliariello, BBA 172, 7 (1969). 54. S. Papa, J. M. Tager, A. Francavilla, and E. Quagliariello, BBA 172, 20 (1969). 55. S. Papa, in “Energy-Level and Metabolic Control in Mitochondria” (S.Papa et al., eds.), p. 401.Adriatica Editrice, Bari, 1969.

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J . B. HOEK, AND L. ERNSTER

More direct evidence for an energy-linked transhydrogenase reaction was furnished by Danielson and Ernster (66-68)who showed that respiratory energy or ATP markedly stimulated the rate of conversion of NADH and NADP+ into NAD’ and NADPH as catalyzed by submitochondrial particles. Thermodynamically, energization led to a shift of the apparent equilibrium toward an extensive reduction of NADPH (69). Later, a similar energy-linked transhydrogenase reaction was demonstrated with membrane fragments of both respiring (60-63) and photosynthetic bacteria (64, 66). The properties of these so-called AB-specific transhydrogenases (see Section I), particularly those of the beef heart enzyme, and their relationship to the energy-supplying systems, have attracted considerable attention (see 66-72). Because of the direct and striking kinetic and thermodynamic effects of energy on the transhydrogenase reaction, this reaction has been widely used as a probe for low energy levels in studies concerning energy coupling in both mammalian and bacterial systems (see 7 3 ) .The properties and function of mitochondrial transhydrogenase have been reviewed previously by Ernster and Lee (74). B. OCCURRENCE AB-Specific transhydrogenase was originally discovered in beef heart by Kaplan et al. (SO) but was also found in kidney, liver, other muscle 56. L. Danielson and L. Ernster, BBRC 10, 91 (1963). 57. L. Danielson and L. Ernster, Biochem. Z . 338, 188 (1963). 58. L. Danielson and L. Ernster, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 157. Academic Press, New York, 1963. 59. C. P. Lee and L. Ernster, BBA 81, 187 (1964). 60. P. S. Murthy and A. F. Brodie, JBC 239,4292 (1964). 61. R. J. Fisher and D. R. Sanadi, BBA 248, 34 (1971). 62. P. D. Bragg and C. Hou, Can. J . Biochem. 46,631 (1968). 63. A. J. Sweetman and D. E. Griffiths, BJ 121, 125 (1971). 64. D. L. Keister and N. J. Yike, BBRC 24,510 (1966). 65. D. L. Keister and N. J. Yike, Biochemistry 6, 3847 (1967). 66. L. Ernster and C. P. Lee, “Methods in Enzymology,” Vol. 10, p. 738, 1967. 67. J. Rydstrom, A. Teixeira da Cruz, and L. Ernster, Eur. J. Biochem. 17, 56 (1970). 68. A. Teixeira da Crua, J. Rydstrom, and L. Ernster, Eur. J. Biochem. 23, 203 (1971). 69. J. Rydstrom, A. Teixeira da Cruz, and L. Ernster, Eur. J. Biochem. 23, 212 (1971). 70. J. Rydstrom, Eur. J. Biochem. 31, 496 (1972). 71. J. Rydstrom, Ph.D. Thesis, University of Stockholm (Chem. Commun. No. VII), Stockholm, 1972. 72. J. Rydstrom, Eur. J. Biochem. 45,67 (1974). 73. F. Gibson and G. B. Cox, Essays Biochem. 9, 1 (1973). 74. L. Ernster and C. P. Lee, Annu. Rev. Biochem. 33,729 (1964).

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tissues (30, 32, 7 5 ) , and arterial tissue ( 7 6 ) . Different animals showed large discrepancies in distribution of transhydrogenase activity in various organs (30, 3 2 ) , although heart muscle generally contained the highest activity (30, 32, 7 5 ) . Tissues such as brain, prostate, seminal vesicle, spleen, and testis showed low or negligible activities (30, 32, 7 5 ) . Cultured cells from rat hepatoma contain a transhydrogenase that appears to be similar to that in beef heart ( 7 7 ) .Subcellular distribution studies have shown that transhydrogenase is localized in the mitochondria (31-35) where it is tightly bound t o the inner membrane (31-35, 56-68, 78) with its nicotinamide nucleotide binding site(s) exposed to the matrix. The latter is evident from the known impermeability of the mitochondrial inner membrane to nicotinamide nucleotides (79) and from the fact that “inside out” sonic submitochondrial particles (56-58), but not intact mitochondria ( 7 5 ) ,catalyze a rapid transhydrogenation with externally added nicotinamide nucleotides. Further support for a superficial location of the active site (s) of the transhydrogenase in submitochondrial particles is provided by the high sensitivity of the enzyme to trypsin (8 0) . As pointed out by Sweetman et al. (81) this does not exclude that in intact mitochondria part of the transhydrogenase molecule, different from the nicotinamide nucleotide binding site(s), may be exposed to the intermembrane space. Steroid-metabolizing mitochondria, e.g., those from beef adrenal cortex (75, 82-84) and porcine corpus luteum (86, 86), were recently shown to contain an active energy-linked transhydrogenase that appears to play a role in steroid hydroxylation reactions (see Section I V ) . Under proper assay conditions all mitochondria1 transhydrogenases hitherto described may be coupled to respiratory energy or ATP. Among respiring bacteria evidence for an energy-linked transhydrogenase was provided by Murthy and Brodie (60) and by Fisher and Sanadi (61) with membrane fragments from Escherichia coli. A similar but apparently non-energy-linked transhydrogenase was also found in MycoA. M. Stein, N. 0. Kaplan, and M. M. Ciotti, JBC 234, 979 (1959). V. K. Kalra and A. F. Brodie, BBRC 51, 414 (1973). C. DeLuca and R. P. Gioeli, Can. J. Biochem. 50,447 (1972). T. Kawasaki, K. Satoh, and N. 0. Kaplan, BBRC 17, 648 (1964). A. L. Lehninger, Harvey Lect. 49, 174 (1955). K. Juntti, U.-B. Torndal, and L. Ernster, i,n “Electron Transport and Energy Conservation” (J. M. Tager et al., eds.), p. 257. Adriatica Editrice, Bari, 1969. 81. A. d. Sweetman, A. P. Green, and M. Hooper, BBRC 58, 337 (1974). 82. B. JV. Harding and D. H. Nelson, Endocrinology 75,506 (1964). 83. W. Cammer and R. W. Estabrook, ARB 122,721 (1967). 84. S. B. Oldham, J. J. Bell, and B. W. Harding, ABB 123, 469 (1968). 85. J. Robinson and P. M. Stevenson, Eur. J . Biochem. 24, 18 (1971). 86. V. I. Uigiris, E. N. McIntosh, C. Alonso, and H. A. Salhanick, Biochemistry 10, 2916 (1971). 75. 76. 77. 78. 79. 80.

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J. RYDSTR~M,J. B. HOEK, AND L. ERNSTER

bacterium phlei (60). Photosynthetic bacteria, e.g., Rhodospirillum rubrum (64, 66), Rhodopseudomonas spheroides (87) , Rhodopseudomonas v i d i s (66))Rhodopseudomonas palustris (66),and Rhodospirillum molischiunum (66) contain a transhydrogenase that may be driven by light. Occurrence of an ATP-supported transhydrogenase was also reported in phosphorylating membrane fragments from Micrococcus denitrificans (88,89). Little is known about transhydrogenase in plants and yeast. Chloroplasts from spinach do contain a transhydrogenase, but this activity is most likely related to the ferredoxin-NADP’ reductase (11-13; see 66) in these chloroplasts. Hasson and West (90) reported on the interesting finding of a microsomal ATP-stimulated and 2,4-dinitrophenol-sensitive transhydrogenase in the endosperm of seeds of the immature cucumber Echinocystis macrocarpa. Sonic submitochondrial particles from Saccharomyces cerevisiae contain a very low transhydrogenase activity (91) that does not appear to be energy-linked (98). Harlow et al. (93) demonstrated the presence of energy-linked transhydrogenase in the protozoan Entamoeba histolytica. )

C. PREPARATIONS AND ASSAY Mitochondria1 nicotinamide nucleotide transhydrogenase can be estimated in the intact organelle either by removing aliquots and determining the individual oxidized and reduced nicotinamide nucleotides , (20) or by measuring changes of the intrinsic absorption (80, 37) or fluorescence (37) of endogenous reduced nicotinamide nucleotides. As a result of various interfering NAD (P)-dependent reactions and lack of suitable substrate-regenerating systems in intact mitochondria, these types of assays are often inadequate for determinations of absolute rates (see Section IV). With intact bacteria the assays are still more complicated since the levels of total NADP generally are very low (94). In fact, no attempt to estimate transhydrogenase activity in intact bacteria has so far been reported. 87. J. A. Orlando, D . Sabo, and C. Curnyn, Plant Physiol. 41, 937 (1966). 88. A. Asano, K. Imai, and R. Sato, Seikagaku 37,647 (1965). 89. A. Asano, K. Imai, and R. Sato, BBA 143,477 (1967). 90. E. P. Hasson and C. A. West, ABB 155,258 (1973). 91. J. Rydstrom and E. Ross, unpublished observation (1974). 92. G. Schatr and E. Racker, BBRC 22,579 (1966). 93. D. R. Harlow, E. C. Weinbach, and L. S. Diamond, Comp. Biochem. Biophy8.

(in press). 94. A. F. Brodie and D. L. Gutnick, “Electron Transfer in Biological Systems,” Vol. lB, p. 699. Dekker, New York, 1971.

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In order to circumvent the permeability barrier in mitochondria and bacteria to nicotinamide nucleotides, and make possible a reliable assay with added substrates as well as purification, the inner membrane and cell wall, respectively, must be broken. Thus, Kaplan e t al. (SO) employed mechanical disruption by means of a Waring blender to disrupt beef heart tissue and supposedly also mitochondria. Detergents, e.g., digitonin, and subsequently calcium gel adsorption were introduced by Kaplan et al. (SO) for further fractionation of the beef heart submitochondrial particles, resulting in a preparation that was about 10 times more active than the whole homogenate. More extensive purification failed because of the high sensitivity of the enzyme to lipid-removing agents such as acetone and bile salts (SO, 95). Extraction with digitonin followed by ammonium sulfate fractionation was later pursued by Humphrey (32) who did not achieve, however, significantly higher specific activities. I n more elaborate attempts, Kaufman and Kaplan (96) and Kaplan (97) reported on a 14-fold and a 25-fold purification, respectively, relative to digitonin-treated mitochondria. The latter preparation was obtained by repeated sucrose-density gradient centrifugation of the digitonin extract. Again, further purification was hampered by the high sensitivity of the enzyme to organic solvents, bile salts, and phospholipases (95, 9’7). Similar findings were reported by Kramar and Salvenmoser (98) and by Salvenmoser and Kramar (99; see also 100). Disruption of mitochondria by means of sonication (101) has proved to be an excellent method for preparing submitochondrial particles that carry out efficient phosphorylation (34, 35).As demonstrated by Danielson and Ernster (SS-SS), these particles catalyze a rapid reduction of NADP’ by NADH supported by either respiration or ATP. Depending on the composition of the sonication medium either nonphosphorylating or phosphorylating particles are obtained which drive respiration-supported transhydrogenase and respiration ; and ATP-supported transhydrogenase, respectively (102-104). By the addition of low amounts of oligomycin to the nonphosphorylating particles the capacity for phos95. N. 0. Kaplan, “Methods in Enzymology,” Vol. 2, p. 681, 1955. 96. B. Kaufman and N. 0. Kaplan, JBC 236,2133 (1961). 97. N. 0. Kaplan, “Methods in Enzymology,’’ Vol. 10, p. 317, 1967. 98. R. Kramer and F. Salvenmoser, Hoppe-Seyler’s Z. Physiol. Chem. 346, 310 (1966). 99. F. Salvenmoser and R. Kramar, Enzymologiu 40,322 (1971). 100. R. Kramar, M. Miiller, and F. Salvenmoser, BBA 162,289 (1968). 101. G. H. Hogeboom and W. C. Schneider, N u t w e (London) 166, 302 (1950). 102. C. P. Lee, G. F. Azzone, and L. Ernster, Nature (London) 201, 152 (1964). 103. C. P. Lee and L. Ernster. “Methods in Enzymology,” Vol. 10, p. 543, 1967, 104. R. J. Fisher, B. P. Sani, and D. R. Sanadi, BBRC 44, 1394 (1971).

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J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

phorylation as well as ATP-supported transhydrogenation is restored (106,106). Recently, Rydstrom et al. (107) demonstrated that transhydrogenase was solubilized efficiently and selectively from beef heart sonic submitochondria1 particles by lysolecithin, giving a preparation that was about seven times as active as particles. In contrast to earlier preparations this lysolecithin extract was devoid of all cytochromes except cytochrome c. A number of preparations of various bacterial membrane fragments exerting energy-linked transhydrogenase activities have been described. The use of sonication (SO-SS),French press (64,66),Ribi cell fractionator (108),and grinding with sand (109)give preparations that generally are less active with respect to both non-energy-linked and energy-linked transhydrogenase activities than submitochondrial particles. Of particular interest are chromatophores from Rhodospirillum rubrum from which a protein can be isolated that is necessary for both non-energylinked and energy-linked transhydrogenase activities of the chromatophores; the isolated factor itself has no measurable activity (109-113). Protein factors have also been isolated from both Rhodopseudomonas spheroides (114, 116) and Escherichia coli (116).However, both of these proteins appear to influence only the energy-linked transhydrogenase reaction and may therefore function as a general energy-coupling factor rather than a specific transhydrogenase factor (see 117). Assay of AB-specific transhydrogenase is principally identical to that of BB-specific transhydrogenase (see Section I1,C) ; i.e., with the natural nicotinamide nucleotides, an enzymic regenerating system is used to keep the concentration of one of the substrates constant (SO, 39, 6648,66) 105. C. P. Lee and L. Ernster, BBRC 18,523 (1965). 106. C. P. Lee and L. Ernster, Regul. Metab. Processes Mitochondria, Proc. Symp., 2966 BBA Libr., Vol. 7, p. 218 (1966). 107. J. Rydstrom, J. B. Hoek, and T. Hundal, BBRC 60, 448 (1974). 108. G. B. Cox, F. Gibson, L. M. McCann, J. D. Butlin, and F. L. Crane, BJ 132, 689 (1973). 109. R. R. Fisher and R. J. Guillory, FEBS (Fed. Eur. Biochem. Soc.) Lett. 3, 27 (1969). 110. R. R. Fisher and R. J. Guillory, JBC 244, 1078 (1969). 111. R. R. Fisher and R. J. Guillory, JBC 246, 4679 (1971). 112. R. R. Fisher and R. J. Guillory, JBC 246,4687 (1971). 113. A. W. T. Konings and R. J. Guillory, JBC 248, 1045 (1973). 114. J. A. Orlando, ABB 141, 111 (1970). 115. T. J. Berger and J. A. Orlando, ABB 159,25 (1973). 116. P. D. Bragg and C. Hou, FEBS (Fed. Eur. Biochem. Soc.) Lett. 28, 309 (1972). 117. A. W. T. Konings and R. J. Guillory, BBA 283, 334 (1972).

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and a suitable energy source is added. Nicotinamide nucleotide analogs of NADP', i.e., thio-NADP', and of NAD', i.e., acetyl-pyridine-NAD', may be used in the absence of a regenerating system (78, 96,118).Interactions between the transhydrogenase system and the energy pool of the membrane fragments used can be follwed by 8-anilinonaphthalene l-sulfonate fluorescence (72, 119) or by active transport of lipid-soluble anions (120).In Rhodospirillum rubrum chromatophores this type of interaction can also be monitored by spectral changes of carotenoids (121).

D. MOLECULAR PROPERTIES The transfer of hydrogen between NAD(H) and NADP(H), catalyzed by mitochondrial transhydrogenase, occurs without exchange with the hydrogen atoms of the surrounding water phase (106, 122, 123). The enzyme is stereospecific for the 4A hydrogen of NADH and the 4B hydrogen of NADPH (78, 106,122, 123). The same stereospecificity has been reported for the transhydrogenases of Escherichia coli ( 8 ) and Rhodospirillum rubrum (112). It has been proposed (8, 68-71) that AB-specific transhydrogenases have separate binding sites for NAD ( H ) and NADP(H) (see also Section II1,F). The natural nicotinamide nucleotides NADP+ and NAD' can be exchanged for various substrate analogs (118; see also 6 5 ) , e.g., thioNADP' and acetyl-pyridine-NAD+, respectively. The activities with these analogs vary substantially and 3'-analogs of NADP are virtually inactive (8, 7 0 ) .It should be pointed out that "transhydrogenation" (see Section I) between NADH and acetyl-pyridine-NAD', catalyzed by impure transhydrogenase preparations, is attributable to NADH dehydrogenase activity (124, 125) rather than to transhydrogenase activity (97, 100).

A variety of inhibitors of mitochondrial transhydrogenase have been described, some of which are relatively unspecific, such as various SH 118. R. R. Fisher and N. 0. Kaplan, Biochemistry 12,1182 (1973). 119. R. J. van de Stadt, F. J. R. M. Nieuwenhuis, and K. van Dam, BBA 234, 173 (1971). 120. L. L. Grinius, A. A. Jasaitis, Y. P. Kadziauskas, E. A. Liberman, V. P. Skulachev, V. P. Topali, L. M. Tsofina, and M. A. Vladimirova, BBA 216, 1 (1970). . 121. S. A. Ostroumov, V. D. Samuilov, and V. P. Skulachev, FEBS (Fed. Eur. Biochem. Soc.) Lett. 31, 27 (1973). 122. C. P. Lee, N. Simard-Duquesne, L. Ernster, and H. D. Hoberman, BBA 105, 397 (1965). 123. D. E. Griffiths and A. M. Roberton, BBA 118,453 (1966). 124. T. Cremona and E. B. Kearney, JBC 240,3645 (1965). 125. M. Gutman, H. Mersmann, J. Luthy, and T. Singer, Biochemistry 9, 2678 (1970).

70

J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

reagents (32, 96), triiodothyronine (41), Mg2+ (59,67, 186, 1271, Ca2+ (127), Mn2+(127),or D,O (72,128). Specific transhydrogenase inhibitors include various adenine nucleotides which compete with the substrates of the enzyme. I n the case of the AB-specific transhydrogenases those inhibitors display a site specificity in the sense that 2’- or 3’-substituted adenine nucleotides are competitive with respect to NADP(H) , and adenine nucleotides without such a substituent are competitive with respect to NAD(H) (70,129, 130). I n addition, it is found (70,129, 130) that inhibitors competitive with NADP (H) show increasing potency with increasing hydrophobicity. For example, palmityl-CoA, which is a competitive inhibitor of AB-specific transhydrogenases with respect to NADPH(H), is considerably more potent than CoA (70,129, 130). I n contrast, palmityldephospho-CoA, which is a competitive inhibitor with respect to NAD ( H ), is only slightly more potent than dephospho-CoA (70).This pattern is suggestive of a hydrophobic environment of the NADP ( H )-binding site, and a relatively hydrophilic environment of the NAD (H)-binding site of the enzyme. A survey of different site-specific inhibitors of mitochondrial transhydrogenase-which seems to be valid for other Al3-specific transhydrogenases as well (8)-is shown in Table 11. Further characterization of AB-specific transhydrogenase has so far been hampered by the lack of a purified enzyme and most of the available information on the properties of AB-specific transhydrogenase has therefore been obtained using various preparations of membrane fragments. The partially purified transhydrogenase from beef heart was reported by Kaufman and Kaplan (96) and by Kaplan (97)to be a complex lipoprotein with a molecular weight of about 250,000; no flavin could be detected (97).Being a membrane-bound protein and presumably a lipid-dependent enzyme, it is not surprising that mitochondrial transhydrogenase is highly sensitive to lipid-removing agents such as detergents (30, 96,98,107),organic solvents (30,96), and phospholipases (30, 131, 132). Direct evidence for a lipid dependence of transhydrogenase is lacking as yet. However, lecithin has been shown by Pesch (13.9) to be stimu126. A. Hommes, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 39. Academic Press, New York, 1903. 127. T. E. Andreoli, R. L. Pharo, and D. R. Sanadi, BBA 90, 16 (1964). 128. S. A. Margolis, H. Baum, and G. Lenaz, BBRC 25, 133 (1966). 129. J. Rydstrom, A. V. Panov, G. Paradies, and L. Ernster, BBRC 45, 1389 (1971). 130. J. Rydstrom, J. B. Hoek, R. Alm, and L. Ernster, in “Mechanisms in Bioenergetics” (G. F. Azzone et al., eds.), p. 579. Academic Press, New York, 1973. 131. L. A. Pesch and J. Peterson, BBA 06, 390 (1965). 132. V. N. Luzikov, V. V. Kupriyanov, and T. A. Makhlis, Bioenergetics 4, 521 (1973). 133. L. A. Pesch, BBA 81, 229 (1964).

2.

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NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

SITE-SPECIFIC INHIBITORS

TABLE I1 MITOCHONDRIAL TRANSHYDROOENASE'

OF

Inhibitor

Specificity

Non-energylinked

Energylinked

Adenosine 5'-AMP ADP Dephospho-CoA Acetyl-dephospho-CoA 2'-AMP 3'-AMP CoA Acetyl-CoA Palmityl-CoA 3':5'-AMP

NAD(H) NAD(H) NAD(H) NAD(H) NAD(H) NADP(H) NADP(H) NADP(H) NADP(H) NADP(H) NADP(H)

500 300 300 9 11 700 700 200 200 0.15 400

500 700 400 40 40 1200 1200 700 700 0.15 500

From Rydstrom (70).

latory with a transhydrogenase preparation obtained by extraction of beef heart mitochondria with tert-amylalchol. Obviously, the transhydrogenase protein is very labile and difficult to isolate, and it appears that a new technique for isolation of this type of hydrophobic protein is required. I n this respect the use of lysolecithin, as recently reported by Rydstrom et al. (lor), may be promising. An interesting 'development is the isolation of a transhydrogenase factor from Rhodospirillum rubrum by Fisher and Guillory (109-115). Its molecular weight is about 70,000, it contains essential sulfhydryl groups and at least two subunits, and it is heat-inactivated and trypsinsensitive (113). Later, Hoek et al. (8) showed that the membrane component is also trypsin-sensitive. The factor apparently contains no flavin and has no known enzymic activity (113). Both non-energy-linked (112) and energy-linked (including light-driven) transhydrogenase activities of the depleted chromatophores are absent unless transhydrogenase factor is added (109-113). Binding of the factor to the membrane component is strongly influenced by the substrates of the transhydrogenase reaction (111). In contrast to the transhydrogenase factor isolated from Rhodopseudomonas spheroides (114, 116), the Rhodospirillum rubrum factor cannot be replaced by thiol reagents (117).

E. RELATIONSHIP TO

THE

ENERGY-COUPLING SYSTEM

AB-Specific transhydrogenases are functionally coupled to the energytransfer system of the membrane in which they are located. This coupling

72

J.

RYDSTRBM,

J . B. HOEK, AND L. ERNSTER

is manifested by an energy-dependent increase in both the rate (57, 58, 60, 64, 89) and the extent (59, 67) of reduction of NADP’ by NADH. I n mitochondria (56-58, 106) and respiring bacteria (6043), the transhydrogenase reaction can be driven by energy generated either by electron transport through any of the coupling sites of the respiratory chain or by ATP hydrolysis. It can also be driven by a potassium ion gradient across the mitochondria1 inner membrane in the presence of valinomycin (134). In photosynthetic bacteria, energy can be generated either by light-induced electron transport or by the hydrolysis of inorganic pyrophosphate or ATP (64, 65,87), Inhibitors of electron transport do not affect the ATP-driven transhydrogenase reaction (5648, 65) whereas energy-transfer inhibitors, e.g., oligomycin or N,N’-dicyclohexylcarbodiimide, do not inhibit the respiration-, light-, or pyrophosphate-driven reactions, but inhibit the ATPdriven reaction (56, 57, 61, 65, 78, 108). I n so-called nonphosphorylating submitochondrial particles, oligomycin stimulates and may even be obligatory for the energy-linked transhydrogenase reaction driven by either respiration or ATP (103,106,135-137). Uncouplers of oxidative and light-induced phosphorylation abolish the energy-linked transhydrogenase reaction (47, 58, 60-65, 89, 106). Similarly, the reaction is abolished by the combined effects of valinomycin and nigericin in the presence of potassium iops (138). I n submitochondrial particles it has been found that both uncouplers and oligomycin inhibit the ATP-driven transhydrogenase less efficiently than ATP-driven reduction of NAD’ by succinate (106, 139). On the other hand, the two reactions were equally sensitive (139, 140) to the mitochondrial ATPase-inhibitor protein of Pullman and Monroy ( 1 4 1 ) , and it has been suggested (139, 140) that the transhydrogenase and ATPase interact in a direct molecular fashion. Interestingly, in bacterial membrane particles the ATP-driven transhydrogenase reaction has been reported to be more uncoupler-sensitive than the succinate-linked NAD’ reduction (62, 6 3 ) . 134. E. Conover, in “Energy Transduction in Respiration and Photosynthesis” (E. Quagliariello, S. Papa, and C. S.Rossi, eds.), p. 999. Adriatica Editrice, Bari, 1971. 135. C. P. Lee and L. Ernster, BBRC 23, 176 (1966). 136. C. P. Lee and L. Ernster, “Round Table Discussion on Mitochondria1 Structure and Compartmentation” (E. Quagliariello et al., eds.), p. 353. Adriatica Editrice, Bari, 1967. 137. C. P. Lee and L. Ernster, Eur. J . Biochem. 3, 385 (1968). 138. M. Montal, B. Chance, C. P. Lee, and A. Azzi, BBRC 34, 104 (1969). 139. L. Ernster, K. Juntti, m d K. Asami, Bioenergetics 4,351 (1972). 140. K. Asami, K. Juntti, and L. Ernster, BRA 205, 307 (1970). 141. M. E. Pullman and G. L. Monroy, JBC 238,3762 (1963).

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73

NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES Substrate

Oxyqen

-K

ow ’ Oliqomycin

I

NAN

-

+ NADP+

“F“

NADH+ NAD+

“ATP

FIQ.1. Relationship of the energy-linked transhydrogenase reaction to oxidative phosphorylation. From Ernster et al. (166).

The available information thus suggests that the energy-linked transhydrogenase reaction and electron transport-linked phosphorylation utilize a common energy pool. A consequence of this assumption would be that the two processes are competing for the energy available in this pool. Experiments carried out with the purpose of demonstrating such a competition in submitochondrial particles were not successful initially (47, 48). However, it was later shown by Lee and Ernster (135-137) that under favorable conditions, i.e., limited respiration and the presence of saturating concentrations of NADH and NADP’, a competition indeed occurred between energy-linked transhydrogenation and oxidative phosphorylation. It was found that oxidative phosphorylation caused an increase in K , of the energy-linked transhydrogenase for NADH and that the energy-linked transhydrogenase reaction caused an increase in K , of the oxidative phosphorylation system for Pi (135-137). A schematic relationship of the energy-treated transhydrogenase reaction to oxidative phosphorylation in mitochondria is depicted in Fig. 1. The energy expenditure of the mitochondria1 energy-linked transhydrogenase reaction has been estimated to one high energy bond per NADP’ reduced (57, 58, 106, 137, 142?-1&). The same stoichiometry was reported for the energy-linked transhydrogenases in Escherichia coli (63), Rhodospirillum rubrum (66), and Micrococcus denitrifioans (89). The overall ATP-driven transhydrogenase reaction may thus be written as NADH

+ NADP+ + ATP 6NAD+ + NADPH + ADP + P,

Since the equilibrium constant of the non-energy-linked transhydrogenase reaction is 0.79 (30) and that of ATP hydrolysis is about lo5 M 142. 9. Papa, A. Alifano, J. M. Tager, and E. Quagliariello, BBA 153, 303 (1968). 143. D. W. Haas, BBA 82,200 (1984). 144. J. M. Tager, G . S. P. Groot, D. Roos, 8. Papa, and E. Quagliariello, in “The Energy Level and Metabolic Control in.Mitochondria” (S. Papa et al., eds.), p. 453. Adriatica Editrice, Bari, 1969.

74

J. RYDSTRbf, J. B. HOEK, AND L. ERNSTER

(based on a AGO of 7.3 kcal/mole; see 146),the equilibrium constant of the above overall reaction is also of the order of lo6 M . In spite of the very unfavorable equilibrium, van de Stadt et al. (119) have succeeded in demonstrating a reversal of the reaction, leading to net ATP synthesis by using a very high nicotinamide nucleotide potential (defined as [NADPH] [NAD+]/ [NADH] [NADP+]) and efficient regenerating systems for NADPH, NAD+, and ADP. Additional evidence for the reversibility of the energy-linked transhydrogenase reaction was provided by Skulachev and associates in both submitochondrial particles (120, 146-148) and chromatophores derived from Rhodospirillum rubrum (IN), using transport of lipophilic anions as a probe for the high energy state. The mechanism by which energy is transferred between the various energy-generating systems and the transhydrogenase is not known. The problem is intimately related to the more general problem of the mechanism of energy conservation in biological membranes (for reviews, see 149, 160). Attempts have been made to explain the energy-linked transhydrogenase reaction in terms of one of the three main current hypotheses of energy conservation, namely, the chemical (161, 168) conformational (163), and chemiosmotic (164) hypotheses. Mechanisms related to the chemical hypothesis involve energized forms of the substrates, i.e., NADH (or NADP -) as considered a t an early stage by Ernster and associates (6648,106, 136, 137). Protonated species of the substrates traversing the mitochondria1 inner membrane down a transmembrane pH gradient according to the chemiosmotic hypothesis have been proposed by Mitchell (166,166; see also 167).Later, this hypothesis provided the basis for more elaborate mechanisms postulated by Skulachev and coH

J. Rosing and E. C. Slater, BBA 267,275 (1972). V. P. Skulachev, FEBS (Fed. Bur. Biochem. Soc.) Lett. 11, 301 (1970). V. P. Skulachev, Curr. Top. Bioenerg. 4, 127 (1971). A. E. Dontsov, L. L. Grinius, A. A. Jasaitis, I. I. Severina, and V. P. Skulachev, Bioenergetics 3,277 (1972). 149. L. Ernster, Fed. Eur. Biochem. SOC.Symp. 35,257. 150. H. Baltscheffsky and M. Baltacheffsky, Annu. Rev. Biochem. 43, 871 (1974). 161. F. Lipmann, “Currents in Biochemical Research,” p. 137. Wiley (Interscience), New York, 1946. 152. E. C. Slater, Nature (London) 172, 975 (1953). 153. P. D. Boyer, in “Oxidases and Related Redox Systems” (T. E. King, H. 9. Mason, and M. Morrison, eds.), Vol. 2, p. 994. Wiley, New York, 1965. 154. P. Mitchell, Nature (London) 191,144 (1961). 155. P. Mitchell, “Chemiosmotic Coupling in Oxidative and Photosynthetic Phosphorylation.” Glynn Res., Bodmin, Cornwall, England, 1966. 156. P. Mitchell, Bioenergetics 3, 5 (1972). 157. J. Moyle and P. Mitchell, BJ 132,571 (1973). 145. 146. 147. 148.

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NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

75

workers (1.20,146-148),derived from their demonstration of membrane potential changes accompanying the transhydrogenase reaction. Based on the conformation hypothesis of Boyer (153),a principally different mechanism was put forward by Rydstrom e t al. (67,69, 71; see also 1681, who proposed that energization involves a conformational change of the transhydrogenase molecule per se. Such a mechanism, in contrast to the previous ones, provides a satisfactory explanation for both the kinetic and the thermodynamic features of the energy-linked transhydrogenase reaction. The experimental basis of this mechanism is provided by comparative studies of the steady-state kinetics of the energy-linked a,nd non-energy-linked transhydrogenase reactions (see Section II1,F). A conformational change is also included in the so-called electromechanochemical model proposed recently by Green and J i (169).According to this hypothesis, energy-linked transhydrogenation involves a respiration- or ATP-dependent polarization of a “transhydrogenase supermolecule,” which is discharged in the course of the transhydrogenase reaction. Probably, the final understanding of the mechanism of the energy-linked transhydrogenase reaction will have to await the elucidation of the mechanism of energy conservation in biological membranes. An the other hand, the energy-linked transhydrogenase reaction may provide a valuable tool for reaching this goal.

F. KINETICSAND REACTIONMECHANISM Teixeira da Cruz e t al. (68) and Rydstrom e t al. (69, 158) have investigated the steady-state kinetics of the non-energy-linked and energy-linked transhydrogenase reactions catalyzed by sonic submitochondrial particles from beef heart. The data obtained seem to establish clearly that the reaction proceeds by way of ternary complex of very short lifetime, i.e., a Theorell-Chance mechanism (Fig. 2 ) . This conclusion was based on linear and convergent double reciprocal plots of initial velocities vs. substrate concentrations, as well as on product inhibition patterns that revealed competitive relationships between the oxidized and reduced forms of the same nicotinamide nucleotide and noncompetitive relationships between NAD+ and NADP+ and between NADH and NADPH. This pattern of product inhibition indicated that the transhydrogenase has separate binding sites for NAD(H) and N A D P ( H ) . Studies with site-specific inhibitors (70,71) suggested, furthermore, that NAD(H) is the first substrate bound by the enzyme. As pointed out by 158. J. Rydstrom, A. Teixeira da Cruz,and L. Emster, in “Biochemistry and Biophysics of Mitochondria1 Membranes” ( G . F. Azzone e t al., eds.), p. 177. Academic Press, New York, 1972. 159. D. E. Green and S. Ji, Proc. N u t . Acud. Sci. U. S . 69, 726 (1972).

76

J. 1

RYDSTRBM,

J . B. HOEK, AND L. ERNSTER

P

)H

h 1

I

5 1.4

(5.3)

3

I

10. a

II

(-0)

I

I

5 0.3

3.0

(10.2)

(2.1)

I

I

I

I

I I I/ E+E-NADH+NADP+-

5

5.1

20.6 (166.0)

I I I \I

E*NADHF=: NADPH-E*NAD++

NAD+- E

e~

FIG.2. Reaction mechanism of mitochondrial transhydrogenase. Rate constants within brackets refer to the energy-linked hydrogenase; kl, k,, k,, and ks are expressed as pM-' min-', where kl and ka are expressed as min-'. Fast reactions are indicated by dashed lines. From Rydstrom et al. (69).

Cleland (160), steady-state kinetics of a Theorell-Chance mechanism can generally apply also to a rapid-equilibrium random mechanism with two dead-end complexes. However, in view of the data obtained with sitespecific inhibitors this latter mechanism is unlikely in the case of the transhydrogenase (70,7 1 ). The proposed mechanism is also consistent with the observation of Fisher and Kaplan (118)that the breakage of the C-H bonds of the reduced nicotinamide nucleotides is not a ratelimiting step in the mitochondrial transhydrogenase reaction. At neutral pH, the maximal initial velocities of the two directions of the non-energy-linked transhydrogenase reaction differ by a factor of about five, the reduction of NADP+ being the slower reaction (SO, 69, 68, 71, 127). Reduction of NADP+ by NADH is maximally active a t about pH 5.5., whereas reduction of NAD+ by NADPH shows a pH optimum a t about 7.0 (30,67, 71, 7.2; see also 3.2). When the reduction of NADP+ by NADH approaches equilibrium, the rate constant of the reaction is increased (67), indicating an activation of the transhydrogenase that is related to the accumulation of the products NAD+ and NADPH. It has been proposed (67, 69, 71) that this activation may involve a conversion of the enzyme from an inactive to an active conformational state, similar to that proposed to occur upon energization (see below). Michaelis constants of the non-energy-linked beef heart transhydrogenase reaction are 9 rJM for NADH, 40 yN for NADP+, 28 p M for NAD', and 20 for NADPH (68); these values are similar to those found with other AB-specific transhydrogenases (66,89; see also 8).Dissociation constants for the E-NADH and E-NAD+ complexes, derived from 160. W. W. Cleland, "The Enzymes," 3rd ed., Vol. 2, p. 1, 1970.

2.

NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASES

77

the steady-state kinetic data (68),are 6.8 and 10.0 @ respectively. I , The individual rate constants for the partial reactions that were computed from the experimentally determined Michaelis and dissociation constants (68)are indicated in Fig. 2. The equilibrium constant of the transhydrogenase reaction, calculated from the Haldane relationship and based on these rate constants, showed good agreement with the directly determined equilibrium constant (68), especially if the value was corrected for the difference in maximal initial velocities of the two directions of the reaction. Addition of an energy source in the form of a respiratory substrate or ATP leads to a 5- to 10-fold increase in maximal initial velocity of the reduction of NADP' by NADH (66-68, 69, 71, 127) as well as to alterations of both the Michaelis constants for NAD(H) and NADP(H) and the dissociation constants of the E-NADH and E N A D + complexes (69, 71). The relative increase in maximal initial velocity is dependent on the energy level (72, 106,155-137), the substrate concentrations (69, 7 1 ) , pH (67, 72), and is also influenced by Mg2+ (69, 67, 126).Energylinked changes in the Michaelis constants are particularly apparent with the oxidized nicotinamide nucleotides, giving values of 6.5 p M and 43.5 UJM for NADP' and NAD+, respectively (69). The dissociation constants of the energy-linked reaction are approximately 0 for the E-NADH complex and 172 pLM for the E-NAD+ complex, i.e., energization promotes the binding of NADH and the release of NAD' (69). On the basis of the above kinetic data the rate constants of the energy-linked reaction were calculated (Fig. 2, values in parentheses). A comparison of these rate constants with those derived for the non-energy-linked reaction reveals that k, and k, are the rate constants that are most influenced by energization. Product inhibition patterns (69) and the order of substrate binding (70, 71) for the energy-linked reaction are the same as for the non-energy-linked reaction. As pointed out previously (Section III,E), the mechanism by which the transhydrogenase reaction is linked to energy is unknown. However, since the substrates of the transhydrogenase reaction are highly water soluble and therefore may be assumed to interact with the enzyme in the aqueous phase, it appears likely that the observed energy-linked changes in dissociation constants reflect a structural alteration of the transhydrogenase molecule per se from an inactive to an active form, rather than alterations of the nicotinamide nucleotides. Recently, evidence was presented (72) indicating that an increased proton concentration mimics the effect of energy on the kinetics of the mitochondria1 transhydrogenase, and it was suggested (72; see also 118) that the energy-linked conversion of the inactive into the active form of the en-

78

J. RYDSTR~M, J. B. HOEH, AND L. ERNSTER

zyme may involve a protonation of a specific group (or groups) of the enzyme. A functional relationship between Complex I (NADH-ubiquinone reductase) and transhydrogenase was suggested by Hatefi (161) and by Hatefi and Hanstein (168),on the basis of the oxidation of NADPH by submitochondrial particles and Complex I. These workers postulated the presence of a specific acceptor site for NADPH in NADH dehydrogenase which is associated with transhydrogenase (see also 163).Subsequently, however, Rydstrom et al. (164) pointed out that NADPH in the absence of NAD’ could be oxidized unspecifically by NADH dehydrogenase (see 166) in a palmityl-CoA insensitive manner (see 70) via the NADH-binding site of the enzyme. This explanation is consistent with the finding (97,100, 107) that transhydrogenase may be readily separated from NADH dehydrogenase. An additional palmityl-CoA sensitive pathway for the oxidation of NADPH by artificial acceptors in submitochondrial particles, possibly constituting a partial reaction of transhydrogenase, has been demonstrated by Ernster et al. (166).

G. RECONSTITUTION Because of its functional relationship to the energy-conserving system and its ready response to low energy levels (see Section III,E), the energy-linked transhydrogenase reaction has been recognized as a valuable tool for studying reconstitution, particularly in bacterial systems. The studies reported have as a rule concerned reconstitution of the energy-coupling system, with the transhydrogenase being present in the particles before the addition of coupling factors. There are two exceptions from this generalization, one of which is the transhydrogenase factor isolated from Rhodospirillum rubrum chromatophores (109-118).This factor is obligatory for both energy-linked and non-energy-linked transhydrogenation ; its properties and function have already been reviewed (see Section 111,D). Butlin (167;see also 73) re161. Y. Hatefi, BBRC 50, 978 (1973); Ann. N . Y . Acad. Sci. 227, 504 (1974). 162. Y. Hatefi and W. G . Hanstein, Biochemistry 12,3575 (1973). 163. C. I. Ragan, W. R. Widger, and T. E. King, BBRC GO, 894 (1974). 164. J. Rydstiom, J. B. Hoek,and L. Ernster, BBA 303,694 (1973). 165. C. Rosai, T. Cremona, J. M. Machinist, and T. P. Singer, JBC 240, 2634 (1965). 166. L. Ernster, C. P. Lee, and U.-B. Torndal, in “The Energy Level and Metabolic Control in Mitochondria” (S. Papa, et al., eds.), p. 439. Adriatica Editrice, Bari, 1969. 167. J. D. Butlin, unpublished observation.

2.

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79

ported on a mutant of Escherichia coli K12 (nut-) that was lacking both non-energy-linked and energy-linked transhydrogenase. Enhancement of both ATP-driven and respiration-driven transhydrogenase in Escherichia coli by a protein factor was reported by Bragg and Hou (116). This factor restored both ATPase activity as well as energy-linked transhydrogenase activity in factor-stripped membrane fragments. An energy-transfer factor from rat liver mitochondria was found to exert a similar stimulation on respiration-driven transhydrogenation in Escherichia coli (168). A different factor was isolated from Rhodopseudomoms spheroides by Orlando (114) which only stimulated the light-driven transhydrogenase reaction and could be replaced by thiols (116). Various mutant strains of Escherichiu coli are lacking Mgz+-Ca2+-activated ATPase and ATP-driven transhydrogenase but retain respirationdriven transhydrogenase (16‘9-172). Washing of membrane fragments obtained from an ATPase-deficient Escherichiu coli strain and subsequent addition of purified Mg2+-Caz+-stimulatedATPase reconstituted the ATP-driven transhydrogenase (171) . Butlin et al. (1‘73) concluded that two proteins specified by the unc A and unc B genes in Escherichia coli K12 were essential for ATP-driven transhydrogenase. A mutant deficient in cytochromes was found to have an unimpaired ATP-driven transhydrogenase reaction (17 4 ) . IV. Physiological Roles of Nicotinamide Nucleotide Transhydrogenases

As pointed out by Krebs and Veech (1‘76), the relationship between the redox states of NAD and NADP in mammalian cells would be governed to a large extent by the substrate levels of NAD- and NADPdependent dehydrogenases, interlinked by shared reactants. The coordination of these systems of interlinked dehydrogenases and, in particular, energy-linked transhydrogenase has been a matter of controversy. Funda168. R. J. Fisher, K . W. Lam, and D. R. Sanadi, BBRC 39, 1021 (1970). 169. D. L. Gutnick, B. I. Kanner, and P. W. Postma, BBA 283, 217 (1972). 170. B. I. Kanner and D. L. Gutnick, FEBS (Fed. Eur. Biochem. Soc.) Lett. 22, 197 (1972). 171. G. B. Cox, F. Gibson, L. M. McCann, J. D. Butlin, and F. L. Crane, BJ 132, 689 (1973). 172. P. D. Bragg and C. Hou, BBRC 50, 729 (1973). 173. J. D. Butlin, G. B. Cox, and F. Gibson, BBA 292, 366 (1973). 174. A. P. Singh and P. D. Bragg, BBRC 5‘7,1200 (1974). 175. H. A. Krebs and R. L. Veech, in “The Energy Level and Metabolic Control in Mitochondria” (S. Papa et el., eds.), p. 329. Adriatica Editice, Bari, 1969.

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J. RYDSTR~M, J. B. HOEK, AND L. ERNSTER

mental to this argument is the question of the redox states of mitochondrial NAD and NADP in vivo, and we shall devote some attention to this problem in one of the following subsections. The majority of synthetic reactions in mammalian cells takes place in the cytosol. The intramitochondrial localization of transhydrogenase excludes a direct participation in these anabolic processes. Substrate shuttle mechanisms (176, 177) are required to allow for the interaction between intra- and extramitochondrial nicotinamide nucleotide-dependent reactions, In the first instance transhydrogenase can be regarded to be functionally related to intramitochondrial NADP-linked reactions. A number of studies on isolated mitochondria have elaborated these relationships in some detail, in particular with regard to mitochondria1 monooxygenation reactions and to the metabolism of glutamate and isocitrate. Whereas these studies primarily emphasize the role of energy-linked transhydrogenase in the supply of reducing equivalents at the expense of energy, other authors (I@, 166) regard the enzyme as a component of a fourth coupling site of the respiratory chain, which, when catalyzing the reduction of NAD+ by NADPH, may drive ion translocations (147, 167) and ATP synthesis (119). Even less satisfactory than in the case of mammalian transhydrogenase is the experimental evidence suggesting specific metabolic functions for the enrymes occurring in bacterial systems. Keister and Hemmes (16) pointed out that the inhibitory effect of NADP+ on transhydrogenase from Chromatium would indicate a controlling function of the enzyme in the transfer of reducing equivalents from NADPH (produced in the glyoxylate cycle in this organism) to NAD+, thus providing a regulatory site for carbohydrate synthesis. The transhydrogenase from Pseudomoms and Azotobacter, being subject to a complex allosteric regulation (see above), could well play a comparable role in controlling the pathway of reducing equivalents from NADPH. The close physical association of transhydrogenase with enzymes involved in pyruvate oxidation in Azotobacter vinelandii led van de Broek et al. (19) to suggest that the enzyme is involved in regulating the transfer of reducing equivalents from pyruvate to either N, or 0,.Bragg et al. (178) observed that the transhydrogenase activity in Escherichia coli was suppressed by the presence of amino acids in the growth medium, and they suggested the enzyme to be involved in amino acid synthesis in this organism. The mechanism 178. A. J. Meijer and K. van Dam, BBA 346,213 (1974). 177. P. Borst, in “Functionelle und morphologische Organization der Zelle,” p. 137. Springer-Verlag, Berlin and New York, 1963. 178. P. D. Bragg, P. L. Davies, and C. Hou, BBRC 47, 1248 (1972).

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was not elaborated in detail, but more information on this effect may be expected with the recently announced isolation of a mutant of Escherichia coli deficient in both energy-linked and non-energy-linked transhydrogenase [ i.e., presumably a mutant in the transhydrogenase enzyme (73,167)1.

A. REWX STATE OF MITOCHONDRIAL NICOTINAMIDE NUCLEOTIDES By their careful analyses of oxidized and reduced nicotinamide nucleotides in isolated mitochondria under a variety of metabolic conditions, Klingenberg and co-workers (20, 38,179; see also 37, 40, 47) established that under energized conditions mitochondrial NADP is highly reduced compared to NAD. These findings form the main experimental support for the existence of an energy-linked transhydrogenase functioning in intact mitochondria. However, the ratio of oxidized and reduced NAD and NADP as measured after extraction of the mitochondria is not necessarily equal to their thermodynamic redox state (l79a); any factor that influences the activity coefficient of the oxidized and reduced forms of the nicotinamide nucleotides to a different extent would result in a difference between these entities. The question arises to what extent the relatively high reduction level of mitochondrial NADP reflects a true difference in redox state between NAD and NADP. Moyle and Mitchell (157)suggested that the reduction level of free NADP in isolated rat liver mitochondria is much higher than that of total NADP to the extent that the potential difference between NAD and NADP is sufficient to provide the free energy for phosphorylation of ADP to ATP. A similar conclusion is indicated by the tentative calculations of Williams (181) and Greenbaum et al. (182),who estimated the redox state of mitochondrial NADP in the intact liver from the total tissue levels of metabolites of isocitrate dehydrogenase by the metabolite indicator method (see 175, 180). 179. M. Klingenberg, “Zur Bedeutung der freien Nukleotide,” Vol. 11, p. 82. Springer-Verlag, Berlin and New York, 1961. 179a. The following terms are used throughout this section: “total NAD and NADP” refers to nicotinamide nucleotides as measured after appropriate extraction ; “free NAD and NADP” refers to activities of nicotinamide nucleotides in a specific cellular environment; “redox state of NAD and NADP” is used in its thermodynamic sense and denotes the ratio of activities of oxidized and reduced NAD and NADP, respectively (see 176, 180). 180. T. Biicher and M. Klingenberg, Angew. Chem. 70,552 (1958). 181. J. R. Williams, in “The Energy Level and Metabolic Control in Mitochondria” (S. Papa et al., eds.), p. 385. Adriatica Editrice, Bari, 1969. 182. A. L. Greenbaum, K. A. Gumaa, and P. McLean, ABB 143, 617 (1971).

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Although valuable from a methodological point of view, these calculations are based on a large number of assumptions, some of which lack sufficient experimental support (see 183) (this point is partially illustrated by the large variations in results found by Greenbaum et al. (182) when applying different methods of calculation). In fact, preliminary results of Hoek and Ernster (184) indicate that there exist only minor differences between the reduction level of free and total mitochondria1 NADP. I n contrast, Krebs and co-workers (176,186, 186) concluded that, in vivo, the redox state of mitochondrial NADP is close to that of NAD. These conclusions were based on the estimation of glutamate and 8-hydroxybutyrate redox couples in freeze-clamped rat liver as indicators of the redox state of mitochondrial NAD and NADP, respectively (see also 187-189).Hoek and Tager (1YO) made similar calculations on isolated rat liver mitochondria, equilibrated in the presence of metabolites of glutamate and p-hydroxybutyrate dehydrogenases. They observed that these metabolic couples were at the same redox potential under conditions where total NADP was considerably more reduced than total NAD as a result of the energy-linked transhydrogenase. Later work (191) indicated that glutamate dehydrogenase (which was shown to be reactive with both NAD and NADP in rat liver mitochondria) can maintain a nearequilibrium condition with NAD, but not with NADP, suggesting that the enzyme is unable to compete with the efficient reduction of NADP by the energy-linked transhydrogenase under these conditions. These findings indicate that caution is warranted in the interpretation of calculations based on indicator metabolite levels. Thus, no definite conclusion can be drawn as yet concerning the redox state of mitochondrial NADP in vivo and, in this situation, little can be said about the possible role of transhydrogenase in maintaining a high reduction level of this coenzyme. This point should be kept in mind when the results of studies on isolated mitochondria that will be discussed in the following sections are to be extrapolated to the in vivo situation. 183. J. B. Hoek, Ph.D. Thesis, University of Amsterdam, Mondeel Offsetdrukkery, Amsterdam, 1972. 184. J. B. Hoek and L. Ernster, in “Alcohol and Aldehyde Metabolizing Systems” (R. G . Thurman et al., eds.), p. 351. Academic Press, New York, 1974. 185. D. H. Williamson, P. Lund, and H. A. Krebs, B j 103, 514 (1967). 186. J. T. Brosnan, H. A. Krebs, and D. H. Williamson, BJ 117, 91 (1970). 187. K. 8. Henley and E. G . Laughrey, BBA 201,9 (1970). 188. R. A. F. M. Chamalaun and J. M. Tager, BBA 222, 119 (1970). 189. B. Willms, J. Kleineke, and H. D. Soling, BBA 215,438 (1970). 190. J. B. Hoek and J. M. Tager, BBA 325, 197 (1973). 191. J. B. Hoek, L. Ernster, E. J. de Haan, and J. M. Tager, BBA 333, 540 (1974).

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B. ROLEOF TRANSHYDROGENME IN MITOCHONDRIAL MONOOXYGENATION REACTIONS Several monooxygenation reactions have now been demonstrated to have an intramitochondrial localization in different tissues, including llp- and 18-hydroxylation of steroids in adrenal cortex (192-194) ; side chain cleavage of cholesterol or its sulfate ester in adrenal cortex (195, 196), ovary (197-199) , and other steroidogenic tissues (200-202) ; and la-hydroxylation of 25-hydroxycholecalciferol in kidney (203, 204). These reactions are all catalyzed by an electron transport chain containing cytochrome P-450 as the terminal electron acceptor (see 205-207). A P-450 cytochrome has been suggested to be involved in the 26-hydroxylation of cholesterol in liver mitochondria (208).NADPH is required as a specific hydrogen donor for these processes (for reviews, see 205, 206, 209). Enzymes potentially involved in the supply of NADPH to steroid hydroxylation in adrenocortical mitochondria include transhydrogenase (84, 192, 210, a l l ) , NADP-linked “malic” enzyme 192. A. C. Brownie and J. K. Grant, BJ 57, 255 (1954). 193. A. C. Brownie, J. K. Grant, and D. W. Davidson, BJ 58, 218 (1954). 194. P. Greengard, S. Psychoyos, N. H. Tallan, D. Y . Cooper, 0. Rosenthal, and R. W. Estabrook, ABB 121, 298 (1967). 195. I. D. K. Halkerston, J. Eichorn, and 0. Hechter, JBC 236, 374 (1961). 196. K. D. Roberts, L. Bandi, and S. Lieberman, BBRC 29, 741 (1967). 197. B. Tamaoki and G. Pincus, Endocrinology 69, 527 (1961). 198. P. F. Hall and S. B. Koritz, Biochemistry 3, 129 (1964). 199. S. Sulmovici and G. S. Boyd, Eur. J. Biochem. 3,332 (1968). 200. P. F. Hall and K . B. Eik-Nes, BBA 63, 411 (1962). 201. D. Toren, K. M. J. Menon, E. Forchielli, and R. I. Dorfman, Steroids 3, 381 (1964). 202. G. Morrison, R. A. Meigs, and K. J. Ryan, Steroids, Suppl. 2, 177 (1965). 203. D. R. Fraser and E. Kodicek, Nature (London) 228,764 (1970). 204. R. W. Gray, J. L. Omdahl, J. G. Ghazarian, and H. F. DeLuca, JBC 247, 7528 (1972). 205. C. J. Sih, Science 163, 1297 (1969).

206. M. Hamberg, B. Samuelsson, I. Bjorkhem, and H. Danielsson, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 29. Academic Press, New York, 1974. 207. J. G. Ghaaarian, C. R. Jefcoate, J. C. Knutson, W. H. OrmeJohnson, and H. F. DeLuca, JBC 249, 3026 (1974). 208. I. Bjorkhem and J. Gustafsson, JBC 249, 2528 (1974). 209. P. F. Hall, in “The Testis” (A. D. Johnson, W. R. Comes, and N. L. Van Denmark, eds.), Vol. 2, p. 1. Academic Press, New York, 1970. 210. M. L. Sweat and M. D. Lipscomb, JACS 77,5185 (1955). 211. B. W. Harding, L. D. Wilson, S. H. Wong, and D. H. Nelson, Steroids, Suppl. 2, 51 (1965).

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J. RYDSTR~M, J . B. HOEK, AND L. EBNSTER

($l8-214), and NADP-linked isocitrate dehydrogenase (212, 216). Numerous studies have been carried out with isolated mitochondria to identify the source of NADPH for llp-hydroxylation by studying the effect of inhibitors and uncouplers of oxidative phosphorylation in the presence of different hydrogen donors (83,192, 210, 212, 214-222). Part of this work was inconclusive since several complicating features in the metabolism of adrenocortical mitochondria were insufficiently taken into account such as secondary inhibitory effects of the substrates, inhibitors, or uncouplers used (83,214, 220, 222) ; the requirement of transport of substrates across the mitochondrial membrane (223, 224) ; and the possibility of intramitochondrial dismutation reactions (265). A role for energy-linked transhydrogenase in the supply of NADPH was advocated by several authors (83,84,211,216,218-222). I n contrast, Simpson and Estabrook ($14) suggested that malate is the primary hydrogen donor for llp-hydroxylation. These authors proposed that, in vivo, intra- and extramitochondrial “malic” enzyme would cooperate, ensuring a continuous shuttling of reducing equivalents to mitochondrial NADP from the cytosol (see also 217, 226). However, their experimental evidence supporting a predominant role of malic enzyme has been disputed (219-223). At present, the bulk of the evidence indicates that either of the NADP-linked enzyme systems can, under suitable conditions, provide the reducing equivalents required for llp-hydroxylation in isolated adrenocortical mitochondria. To what extent this conclusion is valid in vivo remains an open question: Little information is available yet on the factors that regulate the contributions of the different pathways of NADPH supply to 1lp-hydroxylation. Reports on the effect of different hydrogen donors and respiratory chain inhibitors on the side chain cleavage of cholesterol in adrenal cortex J. K. Grant, BJ 64,559 (1956). E. R. Simpson, W. Cammer, and R. W. Estabrook, BBRC 31, 113 (1968). E. R. Simpson and R. W. Estabrook, ABB 125,384 (1969). J. L. Purvia, G. R. Battue, and F. G. Peron, in “Functions of the Adrenal Cortex” (K. McKerns, ed.), Vol. 11, p. 1007. Appleton, New York, 1968. 216. F. Guerra, F. G. Peron, and J. L. McCarthy, BBA 117, 433 (1966). 217. D. Foneo, B. W. Harding, and D. H. Nelson, Endocrinology $1, 605 (1967). 218. L. A. Sauer and P. J. Mulrow, ABB 134,486 (1969). 219. Y. Harano and J. Kowal, ABB 153, 68 (1972). 220. L. A. Sauer, ABB 139, 340 (1970). 221. K. 0. Klein and B. W. Harding, Biochemistry 9, 3653 (1970). 222. L. A. Sauer, ABB 149,42 (1972). 223. L. A. Sauer and R. Park, Biochemistry 12,643 (1973). 224. A. N. Launay, J. M. Michejda, and P. V. Vignais, BBA 347, 60 (1974). 226. F. G. Peron and B. V. Caldwell, BBA 164,396 (1968). 212. 213. 214. 215.

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and ovary mitochondria are inconsistent. Uigiris et al. (86) observed an inhibition of the reaction in beef corpus luteum mitochondria by rotenone, antimycin, or cyanide with different added hydrogen donors. They suggested that the major pathway for generation of NADPH involves energy-linked transhydrogenase (see also 209), although a minor contribution of NADP-linked isocitrate dehydrogenase could not be ruled out when isocitrate was added as the hydrogen donor. [No indications of malic enzymic activity in corpus luteum mitochondria have been found (86, 2267.1 In contrast to these observations, a stimulation of the cleavage reaction by inhibitors of respiratory chain oxidation was noted by Hall (227) in beef adrenocortical mitochondria and by Robinson and Stevenson (86) in porcine corpus luteum mitochondria. These findings were interpreted to indicate the involvement of non-energy-linked transhydrogenase in the supply of reducing equivalents (85, 927).Hall observed a correlation between the reduction level of mitochondria1 NAD and the rate of side chain cleavage. However, the reduction level of NADP was much higher than that of NAD and showed little variation under different reaction conditions. When succinate or fatty acids were used as the hydrogen donor, energy was found to stimulate the rate of side chain cleavage (86, 86, 228-250) , suggesting the requirement for reversed electron transport coupled to transhydrogenation (85, 86). Robinson and Stevenson (230) proposed a direct, energy-dependent reduction of NADP' by succinate not involving transhydrogenase, but more experimental evidence is needed to substantiate this proposal. Hochberg et al. (231) found that succinate addition did not stimulate cholesterol side chain cleavage in adrenal mitochondria over the rate sustained by endogenous substrates.

C. ROLEOF TRANSHYDROGENASE IN MITOCHONDRIAL GLUTAMATE AND ISOCITRATE METABOLISM Considerations on the role of transhydrogenase in glutamate metabolism in rat liver mitochondria have been dominated largely by a postulate 226. P. M. Stevenson and P. 1,. Taylor, FEBS (Fed. Eur. Biochem. SOC.)Lett. 19, 251 (1971). 227. P. F. Hall, Biochemistry 11, 2891 (1972). 228. P. F. Hall, Biochemistry 6, 2791 (1967). 229. J. Robinson and P. M. Stevenson, FEBS (Fed. Eur. Biochem. SOC.)Lett. 17, 53 (1971). 230. J. Robinson and P. M. Stevenson, FEBS (Fed. Eur. Biochem. SOC.) Lett. 23, 327 (1972). 231. R. B. Hochberg, S. Ladany, M. Welch, and S. Lieberman, Biochemistry 13, 1938 (1974).

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of Klingenberg and co-workers (20, 179) that glutamate dehydrogenase in intact mitochondria reacts preferentially with NADP and not with NAD, although the isolated enzyme is known to react almost equally well with both coenzymes (932). An effect of the mitochondria1 energy level on reactions involving glutamate dehydrogenase was demonstrated by several authors (37-40, 45, 46, 49-50), and it was suggested that the reduction level of NADP+, as set by the energy-linked transhydrogenase reaction, is the main factor regulating the degree of domination of glutamate in isolated mitochondria and in vivo (46, 4 9 ) . This conclusion was challenged by Krebs and co-workers (175,185,186) who interpreted their observations on metabolite levels of the glutamate and p-hydroxybutyrate redox couples in freeze-clamped liver as evidence that glutamate dehydrogenase reacts with both NAD and NADP in the intact tissue (see above). A recent reinvestigation of the work on isolated mitochondria by Hoek et al. (191) led to a reinterpretation of the experimental evidence on the nicotinamide nucleotide specificity of glutamate dehydrogenase. These studies indicated that glutamate dehydrogenase in intact isolated rat liver mitochondria can react with both NAD and NADP, the degree of reactivity with either coenzyme being determined by their relative concentrations. Therefore, in the presence of appropriate concentrations of its substrates the enzyme can catalyze a transhydrogenation that counteracts the energy-linked reduction of NADP+ by NADH via transhydrogenase (see 175). Two different isocitrate dehydrogenases are present in mammalian mitochondria, one NAD-linked, the other NADP-linked (233, 234). Thus, two pathways are available for the oxidation of isocitrate by oxygen (cf. 235). I n one of these, the NADP-linked pathway, transhydrogeBase is involved in the transfer of reducing equivalents from NADPH to NAD+. Nicholls and Garland (52) compared the rate of isocitrate oxidation in intact rat liver mitochondria with the maximal rate of the reverse transhydrogenase reaction and concluded that in the absence of other reactions for NADPH oxidation the NADP-linked pathway can contribute only to a minor extent under energy-rich conditions (see 75, 236). This conclusion is in agreement with several other lines of evidence indicating that the NAD-linked pathway for oxidation of isocitrate by 232. J. A. Olson and C. B. Anfinsen, JBC 202,841 (1953). 233. L. Ernster and F. Navazio, E z p . Cell Res. 11,483 (1956). 234. L. Ernster and F. Navazio, BBA 26, 408 (1957). 235. G. W. E. Plaut, “The Enzymes,”2nd ed., Vol. 7, p. 106, 1963. 236. A. M. Stein, J. H. Stein, and S. R. Kirkman, Biochemistry 6, 1370 (1967).

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the respiratory chain is the prevalent one (for reviews, see 52, 74, 237, 238). Moyle and Mitchell (157) recently reemphasized the NADP-linked pathway for isocitrate oxidation and argued that NAD-linked isocitrate dehydrogenase is absent in rat liver mitochondria. Their experimental evidence has, however, been criticized (239) and presently there are no convincing arguments to doubt the predominant role of NAD-linked isocitrate dehydrogenase in the mitochondrial oxidation of isocitrate under physiological conditions. I n view of the high reduction level of NADP in energized mitochondria, the possibility suggests itself that NADP-linked isocitrate dehydrogenase may function primarily in isocitrate synthesis. Under appropriate conditions, the reductive carboxylation of a-ketoglutarate was shown to be catalyzed by mitochondria from rat liver and kidney in an NADPH-dependent reaction (54, 55, 184, 2 3 9 ~ ) . The synthesis of isocitrate wiih NAD-linked substrates required a supply of energy,’indicating the involvement of the energy-linked transhydrogenase (53, 184). The NADPH-dependent intramitochondrial synthesis of isocitrate is of particular interest in connection with a proposed a-ketoglutarate-isocitrate shuttle, functioning in the transfer of reducing equivalents from mitochondrial to cytosolic NADP (24, 5 5 ) . This substrate shuttle would involve the intra- and extramitochondrial NADP-linked isocitrate dehydrogenases and the translocators for a-ketoglutarate and isocitrate in the mitochondrial inner membrane. Recently, the a-ketoglutarate-isocitrate shuttle system was reconstituted with isolated rat liver mitochondria plus added isocitrate dehydrogenase and NADP (184). Under these experimental conditior,s, mitochondrial NADP-linked substrates could be used as hydrogen donors for the reduction of extramitochondrial NADP’ and, with appropriate enzyme systems added, for driving extramitochondrial NADPH-linked reactions such as glutathione reduction and microsomal hydroxylation reactions (240).Although these studies indicate that, in principle, the a-ketoglutarate-isocitrate shuttle can operate, thus providing a functional link between energy-linked transhydrogenase and extramitochondrial NADPH-dependent reactions, the relevance of these findings to the physiological role of transhydrogenase is questioncbble as yet. In particular, it remains to be established why, under various condi237. G. W. E.Plaut, Cun.Top. Cell. Regul. 2, 1 (1970). 238. G. D. Greville, “Citric Acid Cycle,” p. 1. Dekker, New York, 1969. 239. J. B. Hoek, J. Rydstrom, and L. Ernster, BBA 305,669 (1973). 239a. C. R. Mackerer in “Energy Metabolism and the Regulation of Metabolic Processes in Mitochondria” (M. Mehlman and R. W. Hanson, eds.), p. 271. Academic Press, New York, 1972. 240. J. B. Hoek and L. Ernster, unpublished observations (1974).

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tions, the need for cytosolic NADPH cannot be met entirely by cytosolic reactions (see 2.41-246).

D. ROLEOF TRANSHYDROGENASE IN FATTY ACID SYNTHESIS The inhibitory effect of CoA-thioesters of long chain fatty acids, in particular palmityl-CoA, on transhydrogenase activity in submitochondrial particles from beef heart (70, 71, 129,130) was suggested to indicate a role of the enzyme in lipogenesis. Palmityl-CoA has earlier been shown to inhibit several enzymes involved in fatty acids synthesis (246-261 ) , and it has been proposed (S@) that this compound may act as a regulator of fatty acid metabolism in vivo. Intramitochondrial fatty acid elongation was reported to utilize preferentially NADH (269-964) or both NADH and NADPH (266,266). Podack and Seubert (267)isolated an NADPH-specific enoyl-CoA reductase from rat liver mitochondria. This enayme was found to react preferentially with fatty acids of medium chain length. Recently, the authors suggested (268) that the enzyme is involved primarily in the mitochondria1 elongation of unsaturated fatty acids. Transhydrogenase could well play a role in the supply of NADPH for this process, but experimental evidence substantiating this proposal is lacking as yet. 241. C. M. Nepokroeff, M. R. Lakshmanan, G. C. Ness, R. A. Muesing, D. A. Kleinsek, and J. W. Porter, ABB 162, 340 (1974). 242. S. J. Wakil, in “Lipid Metabolism” (S. J. Wakil, ed.), p. 1. Academic Press, New York, 1970. 243. R. G. Thurman and R. Scholz, Eur. J. Biochem. 10,459 (1969). 244,W. Cleland, Annu. Rev. Biochem. 36, 77 (1967). 246. P. E. Sluse, A. J. Meijer, and J. M. Tager, FEBS (Fed. Eur. Bwchem. Soc.) Lett. 18, 149 (1971). 246. J. W. Porter and R. W. Long, JBC 233,20 (1958). 247. P. K. Tubbs, BBA 70,608 (1963). 248. 0.Wieland and L. Weiss, BBRC 13, 26 (1963). 249. P. K. Tubbs and P. B. Garland, BJ 93,550 (1964). 250. P. A. Srere, BBA 108,445 (1965). 251. J. A. Dorsey and J. W. Porter, JBC 243,3512 (1968). 252. J. V. Dahlen and J. W. Porter, ABB 127,207 (1968). 253. W. Colli, P. C. Hinkle, and M. E. Pullman, JBC 244, 6432 (1969). 254. E. M. Wit-Peeters, Ph.D. Thesis, University of Amsterdam, Mondell Offsetdrukkery, Amsterdam, 1971. 255. 9. J. Wakil, J . Lipid Res. 2,1 (1961). 256. E. Quagliariello, C. Landriscina, and P. Coratelli, BBA 164, 12 (1968). 257. E.R.Podack and W. Seubert, BBA 280,235 (1972). 258. W. Seubert and E. R. Podack, Mol. Cell. Biochem. 1,29 (1973).

Flavin-Containing Dehydrogenases CHARLES H . WILLIAMS. JR .

I . Introduction . . . . . . . . . . . . . . . . I1. Pyridine Nucleotide-Disulfide Oxidoreductasea . . . . . . A . The Reactions Catalyzed-Chemical Similarities and Cross-Reactivity . . . . . . . . . . . . B. Similarities and Contrasts in Mechanism . . . . . C . Similarities and Contrasts in Structure . . . . . . I11. Lipoamide Dehydrogenase . . . . . . . . . . . . A . Metabolic Functions . . . . . . . . . . . B . Review of the Mechanism of Massey and Veeger . . . C . Properties of the 2-Electron-Reduced Enzyme, EH1 . . D . Kinetic Studies . . . . . . . . . . . . . E . Role of NAD+ as a Modifier . . . . . . . . . F. Structural Studies . . . . . . . . . . . . G. Summary and Conclusions. . . . . . . . . . IV . Glutathione Reductase . . . . . . . . . . . . . A . Metabolic Functions . . . . . . . . . . . B . Properties of the 2-Electron-Reduced Enzyme, EHp . . C . Kinetic Studies . . . . . . . . . . . . . D . Thiol Groups . . . . . . . . . . . . . V . Thioredoxin Reductase . . . . . . . . . . . . . A . Metabolic Functions . . . . . . . . . . . B . Specificity of Thioredoxin Reductase . . . . . . . C . General Properties of the E. coli Enzyme . . . . . D . Reduced States of the Enzyme-Mechanism . . . . E . Light-Activated Reduction-Neutral Semiquinone . . . VI . Microsomal Electron Transport . . . . . . . . . . A . The NADPH-Cytochrome P-450 Reductase-Containing System . . . . . . . . . . . . . . . B. The NADH-Cytochrome bs Reductase System . . . . C . Possible Synergism between the Two Microsomal Systems . 1). NADPH-Dependent Mixed Function Amine Oxidase . . 89

90 92 92 94 99 106 107 111 111

115 117 120 126 129 129 133 138 141 142 142 144 144 145 147 148 149 150 151 153

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VII. NADH-Cytochrome bs Reductaae . . . . . . . . . . A. Molecular Properties of the Amphipathic and Soluble Forms of the Reductme, . . . . . . . . . B. Review of the Mechanism of Strittmatter . . . . . C. Mechanism of the Reductase Bound to the Microsome . D. Structural Studies . . . . . . . . . . . . E. The Functional Methemoglobin Reductase System of the Mature Erythrocyte. . . . . . . . . . . VIII. NADPH-Cytochrome P-450 Reductaae . . . . . . . . A. General Properties . . . . . . . . . . . . B. Catalytic Activities of the Reductme . . . . . . C. Mechanism . . . . . . . . . . . . . .

154 154 156 161 162 164 165 166 167 169

I. Introduction

The five enzymes lipoamide dehydrogenase, glutathione reductase, thioredoxin reductase, NADH-cytochrome b, reductase, and NADPHcytochrome P-450 reductase are all flavin-containing dehydrogenases that do not contain metals. The terms “dehydrogenase” and “reductase,” when applied to these nonmetalloflavoproteins, usually connote electron transferases which, following reduction by their donor substrate, react only slowly with oxygen. They are thus distinguished from the oxidases which react rapidly with oxygen and from the hydroxylases which react more rapidly with oxygen in the presence of their other substrates. This rather qualitative distinction among simple flavoproteins on the basis of oxygen reactivity correlates rather well with four other properties :

1. Most reductases and dehydrogenases form neutral (blue) semiquinones while most oxidases form anionic (red) semiquinones upon reduction by dithionite or anaerobic photoirradiation in the presence of EDTA ( 1 ) . 2. Only oxidases react with sulfite to give an addition complex (W). 3. Reductases and dehydrogenases catalyze the rapid reduction of 1-electron acceptors such as ferricyanide or cytochrome c whereas oxidases do not ( 8 ) . 4. Those reductases and dehydrogenases thus far tested form superoxide anion upon reoxidation of their reduced forms by oxygen; if oxidases 1. V. Massey and G.Palmer, Biochemistry 5,3181 (1966). 2. V. Maasey, F. Muller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G . Howell, 9. G . Mayhew, R. G. Matthew, and G . P. Foust, JBC 244, 3999 (1969).

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91

form superoxide anion upon reoxidation, it is so firmly bound as to be undetectable (3).

It will be seen, as each enzyme is discussed in detail, that only one of the five covered in this chapter possesses all of the properties considered above as characteristic of reductases. The pyridine nucleotide-disulfide oxidoreductases, lipoamide dehydrogenase (4),glutathione reductase ( 5 ) , and thioredoxin reductase (6-8) share so many properties in common that they will be compared and contrasted before being considered separately. As their group name implies, they catalyze the transfer of electrons between pyridine nucleotides and disulfides. In spite of their similarities they function in widely divergent metabolic roles. NADH-cytochrome b, reductase (9) and NADPH-cytochrome P-450 reductase (10,11) are microsomal enzymes. The latter has been referred to until very recently as NADPH-cytochrome c reductase, since that is how it is assayed, but there is no cytochrome c in microsomes and its physiological acceptor seems to be cytochrome P-450.It is thus distinguished from NADH-putidaredoxin reductase (la), NADPH-adrenodoxin reductase (13),and NADH-rubredoxin reductase (14).The adrenodoxin reductase and the rubredoxin reductase, together with their respective iron-sulfur protein acceptors, each constitute a cytochrome P-450 reductase system. Lipoamide dehydrogenase (16,16) and NADH-cytochrome b, reductase (17)were covered in detail in the second edition of “The Enzymes.” The material in these chapters has stood the test of time remarkably well and will only need to be summarized here. 3. V. Mqssey, S. Strickland, S. G. Mayhew, L. G. Howell, P. C. Engel, R. G. Matthews, M. Schuman, and P. A. Sullivan, BBRC 36,891 (1969). 4. V. Massey, BBA 37, 314 (1960) ; 30, 205 (1958). 5. R. E. Asnis, JBC 213, 77 (1955). 6. S. Black, E. M. Harte, B. Hudson, and L. Wartofsky, JBC 235,2910 (1960). 7. T. Asahi, R.. S. Bandurski, and L. G . Wilson, JBC 236, 1830 (1961). 8. E. C. Moore, P. Reichard, and L. Thelander, JBC 239, 3445 (1964). 9. P. Strittmatter and S. F. Velick, JBC 221,253 and 277 (1956). 10. C. H. Williams, Jr. and H. Kamin, JBC 237, 587 (1962). 11. A. H. Phillips and R. G. Langdon, JBC 237,2652 (1962). 12. I. C. Gunsalus, P. W. Trudgil, and R. DuBus, JBC 241, 1194 (1966). 13. T. Omura, E. Sanders, R . W. Estahrook, D. Y. Cooper, and 0. Rosenthal, ABB 117,660 (1966). 14. T. Ueda and M. J. Coon, JBC 247, 5010 (1972). 15. V. Massey “The Enzymes,” 2nd ed., Vol. 7, p. 276, 1963. 16. D. R. Sanadi, “The Enzymes,” 2nd ed., Vol. 7, p. 307, 1963. 17. P. Strittmatter, “The Enzymes,” 2nd ed., Vol. 8, p. 113, 1963.

92

CHARLES H. WILLIAMS, JR.

II. Pyridine Nucleotide-Disulfide Oxidoreductares

A. THE REACTIONS CATALYZED-CHEMICAL SIMILARITIES AND

CROSS-REACTIVITY

Electron transfer between pyridine nucleotides and disulfide compounds is catalyzed by several flavoproteins and three of these are well characterized. Lipoamide dehydrogenase functions in the oxidative decarboxylation of a-keto acids catalyzing the reoxidation of reduced lipoate by NAD' (18, 19). Glutathione reductase catalyzes electron transfer between NADPH and glutathione (20-22). Thioredoxin reductase catalyzes the reduction of thioredoxin by NADPH (8) ; thioredoxin is a protein of 12,000 molecular weight containing a single cystine residue which is the electron acceptor (23). It is not surprising that enzymes catalyzing such similar chemical reactions should bear striking similarity to one another both structurally and mechanistically. Lipoamide dehydrogenase (24-28), glutathione reductase (g9),and thioredoxin reductase (30,31) contain, in addition to FAD, a reactive disulfide which is functional in catalysis. These flavoproteins consist of two identical or near identical polypeptide chains, each with a reactive cystine residue, and two molecules of FAD ( 3 1 3 6 ) . The specificity of these enzymes toward their disulfide substrates is quite remarkable: There is virtually no cross-reactivity. Since it is quite difficult to separate glutathione reductase from thioredoxin reductase it 18. I. C. Gunsalus, Fed. Proc., Fed. Amer. Soc. E x p . Biol. 13, 715 (1954). 19. L. J. Reed, Advan. Enzymol. 18,319 (1957). 20. N.U. Meldrum and H. L. A. Tarr, BJ 29, 108 (1935). 21. E.E.Conn and J. W. Vennesland, JBC 192,17 (1951). 22. T. W. Rall and A. L. Lehninger, JBC 194,119 (1952). 23. T. C. Laurent, E. C. Moore, and P. Reichard, JBC 239, 3436 (1964). 24. V. Maasey, Q.H. Gibson, and C. Veeger, BJ 77,341 (1960). 25. R. L.Searls and D. R. Sanadi,BBRC 2,189 (1960). 26. R.L. Searls and D. R. Sanadi, JBC 235, PC32 (1960). 27. V. Masaey and C. Veeger, BBA 48,33 (1961). 28. R. L. Searle, J. M. Peters, and D. R.. Sanadi, JBC 236, 2317 (1961). 29. V. Massey and C.H. Williams, Jr., JBC 240, 4470 (1965). 30. G. Zanetti and C. H. Williams, Jr., JBC 242,5232 (1967). 31. L. Thelander, Bur. J. Biochem. 4, 407 (1968). 32. V. Massey, T. Hofmann, and G. Palmer, JBC 237,3820 (1962). 33. B. D.Burleigh, Jr. and C. H. Williams, Jr., JBC 247,2077 (1972). 34. R.D.Mavis and E. Stellwagen, JBC 243,809 (1968). 35. E.T. Jones and C. H. Williams, Jr., JBC 250,3779 (1975). 36. C. H.Williams, Jr., G. Zanetti, L. D. Arscott, and J. K. McAllister, JBC 242, 5226 (1967).

3.

93

FLAVIN-CONTAINING DEHYDROGENASES

is not possible, a t very low levels, to distinguish cross-reactivity from cross-contamination (37). The glutathione reductase activity of thioredoxin reductase preparations is less than 0.01% (8, 37, 38),while the reverse measurement is about 0.4% ( 3 7 ) . These levels of activity are of the same order of magnitude as the oxidase activity of thioredoxin reductase so that they must be considered upper limits (37, 38). The activity of erythrocyte glutathione reductase toward lipoate is quite high being about 3% that with glutathione as acceptor (38-40).The important point is the very high degree of specificity of these enzymes. The specificity of glutathione reductase toward mixed disulfides will be discussed later. The three disulfide substrates are different in many respects. Dihydrolipoamide (6,8-dimercaptooctanoamide) forms a five-membered ring of high stability upon oxidation. I n its physiological form, in the pyruvate and a-ketoglutarate dehydrogenase complexes, the amide nitrogen is the r-amino group of a lysine residue of dihydrolipoyl transacetylase (41-43) or transsuccinylase; thus, it is a protein bound substrate. The three-enzyme complex composed of the transacetylase, the a-keto acid oxidative decarboxylase, and lipoamide dehydrogenase has been the subject of extensive study and review ( 4 4 ) . Oxidized glutathione (GSSG), unlike lipoamide, forms two molecules of 7-glutamylcysteinylglycine (GSH) upon reduction. Its reoxidation is therefore entropically less favorable. As mentioned above, thioredoxin is a protein in which the disulfide is a cystine residue. The two halves of the cystine are separated in the polypeptide chain by only two other residues. The sequence in the region of the disulfide of the Escherichia coli protein is (46-47) -Gly-Pro-ps

-Trp-Ala-Glu-Trp-ps

-Lys -Met

-

s The fluorescence of the tryptophan residues changes markedly upon reduction and reoxidation of the disulfide ( 4 8 , 4 9 ) . S

37. L. D. Arscott and C. H. Williams, Jr., unpublished observations. 38. L. Thelander, JBC 242, 852 (1967). 39. A. Icen, Scund. J. CZin. Lab. Invest. 20, 96 (1967) ; 27, Suppl. 116,5 (1971). 40. E. M. Scott, I. W. Duncan, and V. Ekstrand, JBC 238, 3928 (1963). 41. V. Massey, BBA 38,447 (1960). 42. H. Nawa, W. T. Brady, M. Koike, and L. J. Reed, JACS 82, 896 (1960). 43. K. Daigo and 1,. J. Reed, JACS 84,666 (1962). 44. L. J. Reed and D. J. Cox, “The Enzymes,” 3rd ed., Vol. 1, p. 213, 1970. 45. A. Holmgren, Eur. J. Biochem. 6,474 (1968). 46. D. E. Hall, A. Baldesten, A. Holmgren, and P. Reichard, Eur. J . Biochem. 23, 328 (1971). 47. B. M. Sjoberg and A. Holmgren, JBC 247,8063 (1972). 48. L. Stryer, A. Holmgren, and P. Reichard, Biochemistry, 6, 1016 (1967). 49. A. Holmgren, JBC 247, 1992 (1972).

94

CHARLES H. WILLIAMS, JR.

The three enzymes are quite specific for their respective pyridine nucleotide substrates. Under conditions normally used for assay, lipoamide dehydrogenase is less than 1% as active with NADPH as with NADH (16) and thioredoxin reductase is less than 1% as active with NADH as with NADPH (36, 36). Lipoamide dehydrogenase can transfer electrons to a number of NAD' analogs (9'7).Yeast glutathione reductase is quite specific for NADPH (60),but the erythrocyte enzyme is 20% as active with NADH as with NADPH under the conditions of the standard assay (39,4O, 61).

B. SIMILARITIES AND CONTRASTS IN MECHANISM I n this section on mechanism and in the section to follow on structure, the comparisons will show that the relationship between lipoamide dehydrogenase and glutathione reductase is more marked than is the relationship of either to thioredoxin reductase. Thus, in catalysis, lipoamide dehydrogenase and glutathione reductase cycle between the oxidized state and a spectrally characteristic state in which the enzyme has accepted two electrons and these are shared between the FAD and the active center disulfide. This intermediate does not seem to be operative in thioredoxin reductase, and in this enzyme the FAD and disulfide interact in a different way. The oxidized forms of these enzymes can then be represented as

The spectra of oxidized glutathione reductase and of the 2-electronreduced enzyme are shown in Fig. 1. This red intermediate, which has been shown to be functional in catalysis (299))will be referred to as EH, designating a half-reduced active center; it has also been referred to as F (62, 65), but this can be confused with oxidized flavin in other nomenclatures. Its spectral characteristics are virtually identical with those of the analogous species of lipoamide dehydrogenase (24, 27, 64). It has 50. V. Massey, C. H. Williams, Jr., C . Zanetti, and G. Foust, h o e . Znt. Congr. Biochem., i'h, 1967 Abstracts, Vol. 1, p. 165 (1968). 51. G. E. J. Staal and C. Veeger, BBA 185,49 (1969). 52. J. E. Bulger and K. G. Brandt, JBC 246,5570 (1971). 53. J. E. Bulger and K. G. Brandt, JBC 246, 6578 (1971). 54. N. Savage, BJ 67, 146 (1957).

3.

FLAVIN-CONTAINING DEHYDROGENASES

95

Wavelenqth (nm)

FIG.1. Yeast glutathione reductase. Spectra of the oxidized and 2-electron-reduced oxidized enzyme, (---I forms of the enzyme recorded anaerobically (29): (-) 526 moles GSH, and (0) 1 mole T P N H in the presence of DPNase.

relatively high absorbance in the 450-nm region and a shoulder extending to 650 nm. Two-electron reduction of free FAD results in almost complete bleaching a t wavelengths greater than 440 nm; thus, the flavin of EH, is not fully reduced and a second electron acceptor must be operating. The second electron acceptor in these enzymes is the disulfide of a cystine residue. The disulfide in EH, is also partially reduced. Several lines of evidence support this, but one crucial experiment with pig heart lipoamide dehydrogenase should be described in some detail (27, 66). Arsenite complexes quite specifically with vicinal dithiols as shown in Eq. (1).

The spectrum of the oxidized enzyme is unaffected by arsenite. If one equivalent of NADH is added anaerobically to enzyme in the presence of arsenite, EH, is produced immediately. In the absence of arsenite this species would be stable indefinitely. However, with arsenite present, the flavin is seen to reoxidize almost completely; at the same time the pyridine nucleotide is fully oxidized (Fig. 2). Thus two electrons have been taken up by the enzyme, but the FAD is oxidized. A second electron 55. V. Massey and G. Palmer, JBC 237,2347 (1962).

96

CHARLES H. WILLIAMS, JR.

-

-

0

500

600

700

800

Wavelength (nm)

Fro. 2. Anaerobic reduction of pig heart lipoamide dehydrogenase in the presence of arsenite (97): 1, oxidized enzyme plus 1 mM arsenite; 2, after 1.0 equivalent NADH; 3, after 2.2 equivalents NADH; 4, after 3.3 equivalents NADH; and 5, after NADase.

acceptor is clearly indicated, and the known specificity of arsenite suggests that i t is a disulfide-dithiol. Furthermore, the initial rate of catalysis by lipoamide dehydrogenase is unchanged following preincubation with arsenite alone, or NADH alone, but is strongly inhibited if NADH is included in the preincubation with the arsenite. Addition of more than one equivalent of NADH in the presence of arsenite leads initially to EH, followed slowly by extensive reduction of the flavin as evidenced by loss of absorbance at 455 nm; the enzyme takes on a green color with a very broad absorption centered about 700 nm. (Free FADH, in dilute solution is pale yellow with no absorption beyond 500 nm.) The extinction of this species is enhanced by excess NAD' and by low temperature and indeed is totally dependent on the presence of NAD+. This is demonstrated in Fig. 2 by the loss of the long wavelength absorption upon addition of NADase (Neurospora crassa NAD' glycohydrolase) which hydrolyzes NAD' and NADP+ but is inactive with NADH and NAPH. These properties have led to the suggestion that this form of the enzyme is

3.

97

FLAVIN-CONTAINING DEHYDROGENASES

a charge transfer complex between FADH, and NAD’, and this is further reinforced by the observation of a red shift in the extinction maximum when an acceptor of more positive potential is used ( 5 5 ) . The complex is not formed upon reduction with NADPH in the presence of arsenite and excess NADP’. On the other hand, when glutathione reductase is reduced in the presence of arsenite and excess NADP’ (its natural cofactor) the charge transfer band is formed while it is not formed when NADH and NAD’ are used ( 2 9 ) . Under these conditions the enzymes have accepted four electrons per FAD as a result of the reaction of the nascent dithiol with arsenite. The charge transfer complex is formed too slowly to be significant in catalysis. To reiterate: lipoamide dehydrogenase and glutathione reductase accept only two electrons upon reduction with excess substrate to form EH,, but in the presence of arsenite this intermediate is not stable and 4-electron reduction results. The exact chemical nature of EH2,i.e., the status of the two electrons shared between the FAD and the disulfide, has been much debated and perhaps cannot be adequately described by any single nomenclature. It has variously been referred t o as a biradical (11) (24, 26, 28), as a charge transfer complex in which thiolate is the donor and FAD the acceptor (111) (28, 56,56a),and as a covalent bond between FAD and sulfur (IV) (57). The roman numerals refer t o the structures below i n which only

bm (1)

Ey (n)

p p p? SH

(rn)

(Iv)

(V)

one of the two active centers of the dimeric enzyme is shown; (I) is the oxidized enzyme and (V) is the 4-electron-reduced enzyme which is catalytically inactive. None of these formulations reflects accurately the properties of EH,. It is now clear that it is not a simple semiquinone. Two types of simple semiquinones have been observed i n flavoproteins (1). Spectra of these are shown i n Figs. 3a (58) and 3b. The neutral semiquinone (FADH’) is blue and its spectrum has peaks at 580, 490, 390, and 340 nm with a broad shoulder extending t o 700 nm. The anion semiquinone (FAD’) is red and its spectrum has peaks at 480,400, and 360 nm. Each type is associated with a distinct electron spin resonance 56. E. M. Kosower, in “Flavins and Flavoproteins” (E. C. Slater, ed.), Vol. 1, p. 1. Elsevier, Amsterdam, 1966. 56a. V. Maeaey and S. Ghisla, Ann. N.Y. Acad. Sci. 227, 446 (1974). 57. G. Palmer and V. Massey, in “Biological Oxidations” (T.P. Singer, ed.), p. 263. Wiley (Interscience), New York, 1968. 58. G. Zanetti, C. H. Williams, Jr., and V. Massey, JBC 243, 4013 (1968).

98

CHARLES H. WILLIAMS, JR.

Ii

10 09

as

P O0 67

X

I 0 1 04

03 02

01

00

m

0

x . 3 W*Iqm Inml

1.

Ibl

Fxo. 3. (8) Typical neutral (blue) flavin semiquinone produced upon anaerobic oxidized irradiation of thioredoxin reductase in the presence of EDTA (68): (-) thioredoxin reductase in 2 x lo-' M EDTA, (---) 1 hr light a t Oo, .) 2 hr light at O", (--) 2 hr 40 min light at 0", and ( 0 )4 hr light at 0". (b) Typical anionic (red) flavin semiquinone produced upon anaerobic irradiation of oxynitrilase in the presence of EDTA (I). ( a

-

(ERS) signal ( 1 ) . The spectral characteristics of E H 2 are clearly different from either of these, especially in the 450-600-nm region, and indeed must arise from a n interaction between the FAD and another group, namely, the redox active disulfide. No ESR signal is observed with E H 2 (1, 88, 69). Its long wavelength absorption is only slightly temperature sensitive, and the response is in the opposite direction to that which would be predicted for a charge transfer complex (55).The weak covalent interaction of (IV) implies that the isoalloxazine ring is reduced, and this is clearly inconsistent with the spectrum. The problem seems t o rest with the molecular orbital theorist. The importance of EH, in catalysis by lipoamide dehydrogenase and glutathione reductase has been demonstrated by rapid reaction spectrophotometry. It is produced upon reduction with NADH or NADPH, respectively, in the dead time of the instrument (ca. 3 msec) and is rapidly reoxidized by lipoamide or glutathione at rates commensurate with catalysis (g4, 60, 5 4 ) . A species similar to EH, has not been observed with thioredoxin reductase (30,31). Its anaerobic titration with NADPH is shown in Fig. 4. Two equivalents are required to reach a stable spectrum which is char59. V. Massey, G. Palmer, C. H. Williams, Jr., B. E. P. Swoboda, and R. H. Sands, in "Flavins and Flavoproteins" (E. C. Slater, ed.), Vol. 1, p. 133. Elsevier, Amsterdam, 1968.

3.

FLAVIN-CONTAINING DEHYDROGENASES

99

Wavelength (nm)

FIQ. 4. Anaerobic reduction of thioredoxin reductase with NADPH in 0.05 M phosphate, pH 7.6, 5" (SO): (-1 oxidized thioredoxin reductase, (---) 0.5 mole TPNH/mole FAD, .) 1.0 mole TPNH/mole FAD, (*--.) 2.0 moles TPNH/ .-. ' .) 4.0 moles TPNH/mole FAD. mole FAD, and ( a

(

-

0

acterized by virtually complete bleaching in the 450-nm region and absorption of very low extinction extending beyond 700 nm (SO, 31). Thus, this enzyme can accept four electrons without apparent complex formation between its nascent dithiol and the FAD. Initial rates of catalysis are not inhibited by preincubation with mercurial alone or with NADPH alone, but preincubation with both results in complete loss of activity (8, SO). If a second aliquot of enzyme (not preincubated) is added to the inhibited mixture the inhibited enzyme is reactivated and double the control activity is observed. The uninhibited enzyme in the time of mixing has produced sufficient reduced thioredoxin (dithiol form) to reverse the inhibition of the preincubated enzyme (SO). The enzyme is also sensitive in the reduced state to arsenite and Cd*+ (8).

C. SIMILARITIES AND CONTRASTS IN STRUCTURE The gross structure of the pyridine nucleotide-disulfide oxidoreductases is the same, i.e., two polypeptide chains each containing a redox active

100

CHARLES H. WILLIAMS, JR.

disulfide (and no other disulfide) and two molecules of FAD (61).Glutathione reductase (34, 39) and lipoamide dehydrogenase (32) have molecular weights of about 100,000 while the molecular weight of thioredoxin reductase is about 70,000 (31,36,37,62). The FAD is very tightly bound and exhibits an extinction coefficient (at the wavelengths of the visible maxima, Table I) (29, 31, 32, 36, 37) of 11.3 mM-l cm-1 which is equal to that of free FAD (at its maximum, 448 nm) (69). The tightly bound flavin can be related to the amino acid composition to give the minimum molecular weight. Table I gives the amino acid composition of the three enzymes as isolated from a prokaryote and from a eukaryote. The molecular weights calculated from amino acid analysis do not agree well with some of the estimates made by physical measurements, but, on the other hand, neither do measurements made by different physical means agree with one another. In one case an exact comparison can be made between molecular weight of a flavoprotein from amino acid analysis data (70) and the molecular weight subsequently obtained when the protein had been sequenced ( 7 1 ) . The agreement is to less than 1%. The quality of such measurements is subject to two types of error. The error resulting from losses in transfer can be eliminated by making the spectral estimation of flavin directly on the solution used for amino acid analysis (of course, the extinction coefficient of the flavin must be known). The other error results from the presence of apoenzyme; this is seldom serious with flavoproteins such as these where the FAD is tightly bound. Data from amino acid analysis of a number of simple flavoproteins are now available: reductases (31, 33, 36, 60, 62, 63, 72-74) and oxidases 60. L. D. Arscott and C. H. Williams, Jr., unpublished observations. 61. C. H. Williams, Jr., B. D. Burleigh, Jr., S. Ronchi, L. D. Arscott, and E. T. Jones, in “Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 295. Univ. Park Press, Baltimore, Maryland, 1971. 62. S. Ronchi and C. H. Williams, Jr., JBC 247,2083 (1972). 63. R. G.Matthews, L. D. Arscott, and C. H. Williams, Jr., BBA 370, 26 (1974). 64. J. E.Wilson, ABB 144, 216 (1971). 65. M. L.Cohn and I. R. McManus, BBA 276,70 (1972). 66. M. L. Speranea, S. Ronchi, and L. Minchiotti, BBA 327, 274 (1973). 67. R.L.Spencer and F. Wold, Anal. Biochem. 32, 186 (1969). 68. P. M. Harrison and T. Hofmnnn, BJ 80, 38P (1961). 69. H. Beinert, “The Enzymes,” 2nd ed., Vol. 2,Part A, p. 339,1960. 70. S. G.Mayhew and V. Massey, JBC 244, 791 (1969). 71. M. Tanaka, M. Haniu, K. T. Yasunobu, S. Mayhew, and V. Massey, JBC 248, 4354 (1973). 72. G.Forti and E. Sturani, Eur. J. Biochem. 3,461 (1968). 73.P. C. Engel and V. Massey, BJ 125,879 (1971). 74. L.Spate and P. Strittmatter, JBC 248,793 (1973).

3.

FLAVIN-CONTAINING DEHYDROGENASES

101

(76-80).I n comparing reductases with oxidases (and hydroxylases) only one trend is apparent: The tryptophan content of dehydrogenases is low (1-6 per flavin) and of oxidases is high (7-14 per flavin) . (Normalization of the data to a constant molecular weight does not change the conclusion.) This fact is reflected in the ratios of absorbance in the 280-nm region to that in the 450-nm region with reductases showing a lower ratio than oxidases. The relationship between this ratio and the tyrosine plus tryptophan content has been used (Table I) to predict a tryptophan value for yeast thioredoxin reductase. The data in Table I do not show any other obvious trends. The fact that the prokaryotes have markedly lower serine contents than do their eukaryote pair-mates is very probably an anachronism. Using average extinction coefficients a t 280 nm for tryptophan (5.6 mM-' cm-l) and tyrosine (1.3 mM-' cm-l) in proteins (81), the extinction coefficient for enzyme-bound FAD a t that wavelength can be estimated and is found to range from 34 to 40 mM-I ern-'. The extinction coefficient for free FAD a t 280 nm is 21 (69),indicating a marked red shift upon binding. This is important in the interpretation of modification studies of flavoproteins which involve monitoring in the ultraviolet. It has been pointed out, for example, that thiol determinations by the Boyer (82) procedure are often inaccurate because of the dissociation of flavin during the titration (8s). The free carboxyl groups of pig heart lipoamide dehydrogenase have been determined as 56 per FAD (6'4). The glutamine plus asparagine content is, by difference, 35. It can then be calculated that the net charge on the protein will depend on the effective pK values of the histidine residues and will vary with pH from +5 to -6 per FAD. Amide data for thioredoxin reductase (31) allow a similar calculation: The charge can vary from +2 to -8 per FAD. The special reactivity as well as the specificity of the active center disulfides in these enzymes is determined by their environment in the proteins. Crucial components in this environment are the near neighbors of the half-cystines in the primary structure. Peptides have been isolated 75. J. H. Paeur, K . Kleppe, and A . Cepure, ABB 111,351 (1965). 76. A. DeKok and A. B. Rawitch, Biochemistry 8, 1405 (1969). 77. M. Schuman and V. Massey. BBA 227,500 (1971). 78. S. C. Tu, S. J. Edelstein, and D. B. McCormick, ABB 159, 889 (1973). 79. B. Curti, S.Ronchi, U. Branzoli, G . Ferri, and C. H. Williams, Jr., BBA 327, 266 (1973). 80. M. I. S. Flashner and V. Massey, JBC 249, 2579 and 2587 (1974). 81. H. Edelhoch, Biochemistry 6, 1918 (1967). 82. P. D. Boyer, JACS 76, 4331 (1954). 83. G. Palmer and V. Massey, BBA 58, 349 (1962).

K

TABLE I AMINOACID ANALYSIS OF PYRIDINE NUCLEOTIDE-DISULFIDE OXIDOREDFCTASES~

Amino acid Cysteic acid* Aspartic acid Threonine Serine Glutsmic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lysine Histidine Arginine Tryptophan' FAD Total amino acid residues Minimum molecular weight/ FAD (including FAD) - 4 2 0 0 I A m s x via'

Visible maximum (nm) NHpterminal' COOH-terminalm

Lipoamide dehydrogenase (pig heart)*

Lipoamide dehydrogenase (E. eoli)"

Glutathione reductase (yeast)d

Glutathione reductase (E. coZi)*

Thioredoxin reductase (yeast)'

Thioredoxin reductase (E. wZi)~

10 44 26 24 47 18 53 46 44 9 36 31 8 15 36 11 14 2 1 474 51,000

5 41 27 15 49 21 52 52 45 9 38 36 15 42 13 16 4 1 488 53 000

6 49 23 27 45 13 43 33 41 7 29 31 17 16 39 14 16 5 1 454 50,600

7 49 30 16 43 22 45 47 43 11 34 30 16 15 26 12 14 3 1 464 50 800

5 29 28 20 33 13 29 35 19 7 20 26 9 11 24 6 11 4i 1 327 35 900

5 37 25 15 35 10 37 34 21 8 24 31 9 11 14 10 16 1 344 37,700

5.3 455

6.7 455

8.5 460

7.3 462

6.8 456

5.0 456

Ala

8

)

Ser Ala-Lys-L ys

GlY

)

)

1

G~Y

d

F F

2!

x

dE r

"5 4

*r

0

Data are based on spectral measurement of FAD in the solutions used for amino acid analyses. Corrections have been made for

losses in serine and threonine upon hydrolysis by extrapolation of data a t various times of hydrolysis to zero time and for the slow hydrolysis of some peptide bonds involving valine and isoleucine by using only those values obtained at later times of hydrolysis. hIatthews el al. (63). Agreement with other analyses (32, 64,65) is excellent. The high value for arginine (32) has not been borne out in subsequent analyses (63-65).

Williams et a[. (33, 36). Arscott and Williams (60). Numerous errors are apparent in the previous analysis (29). " Arscott and Williams (60). f Speransa el a/. (66). 0 Williams el al. (36, 62). Agreement with another analysis is excellent (31).The Trp value is taken from (31). I, Determined on separate samples hydrolyzed in the presence of dimethylsulfoxide (67). ' Determined on separate samples by a modification of the method of Spies and Chambers (68). Value assumed on the basis of ultraviolet absorption, see text. Ratio of the absorbance a t 280 nm to that a t the visible maximum given in the next line. I Thelander (31), Rlassey et al. (32),Burleigh and Williams (3.3, and Jones and Williams (35). Burleigh and Williams (33). J

+;.r

7 8

2k 5

z

0

U

x" 3

T10 PI

2

01

H m

104

CHARLES H. WILLIAMS, JR.

Pig heart lipoamide dehydmgenase: Glu-Lye -Asx-Glu-Thr-

Lsu- Gly- Gly- Thr-

8

B

Cys- Lsu-Am-Val-

I

Cly -Cye -1le -Pro- Ser -Lys-Ala-Leu

Yeast glutathione reductase:

s Lys-Ala-G4-Lys

-Ala-Leu-Gly-Gly

s

I

I

-Thr -Cys-Val- Asn-Val- Gly-Cye-Val-Pro-Lys

-Val-Val-Met

E s c k e r i c k i a c o 1 f lipoamlde dehydrogenase:

s

s Tyr- Asn -Thr --Leu

- Gly -Gly-Val-

I

Cys- Leu-Am-Val-

Gly

I - Cys-

Ile-Pro-

Sar- Lys

E s c h e r fc h l a c o Ii thioredoxln reductase:

s A l a - 4 8 -Ala-Thr-

s I

Cys -Asp-Gly- Phe

FIG.5. Sequences around the active center disulfide of the pyridine nucleotidedisulfide oxidoreductases.

from E . coli (33, 61, 84) and pig heart (63, 86, 86) lipoamide dehydrogenase, from yeast glutathione reductase (561, and from E. coli thioredoxin reductase (61, 62,87) containing both halves of the reactive cystine residue. The sequences of these peptides are shown in Fig. 5 . A feature common to the disulfides is that the half-cystines are close to one another in the primary sequence, being separated by four other residues in lipoamide dehydrogenase and glutathione reductase and by two other residues in thioredoxin reductase, thus conforming tight loops in the polypeptide chain. This structural arrangement had been predicted for lipoamide dehydrogenase (28). The E. coli and pig heart lipoamide dehydrogenase sequences (a prokaryote-eukaryote pair) are identical in 14 of 17 overlapping residues. It is suggested that these proteins have been derived by divergent evolution from a common ancestor (86, 867, but a final judgment on this point will have to await total sequencing. A very high degree of homology also exists between the lipoamide dehydrogenases and glutathione reductase in this analogous region (36).The two substitutions in the immediate disulfide area, valine for leucine and isoleucine, are conservative both chemically and genetically. It may be of interest that the prokaryote 84. J. P. Brown and R. N. Perham, FEBS (Fed. Eur. Biochem. Soc.) Lett. 26, 221 (1972). 85. C. H. Williams, Jr. and L. D. Arscott, 2. Naturforsch. B 27, 1078 (1972). 86. J. P. Brown and R. N. Perham, BJ 137,505 (1974). 87. L. Thelander, JBC 245, 6026 (1970).

3.

FL AVIN -CONTAI N I N G DEHYDROG E N ASES

105

E. coli lipoamide dehydrogenase has a valine residue preceding the first half-cystine, whereas the eukaryote enzymes have a threonine residue. Chemically this is a relatively conservative change since the side chains of these amino acids are virtually identical in volume. The lack of homology between lipoamide dehydrogenase and thioredoxin reductase, and the apolar nature of the residues around the active center disulfide in lipoamide dehydrogenase, an enzyme with an apolar substrate, led to the postulation that the disulfide region contained important determinants for interaction with the respective substrates (33, 62, 85, 88).The very high degree of homology between lipoamide dehydrogenase and glutathione reductase suggests that some modification of that postulation is in order; clearly, glutathione and lipoamide are very different molecules. It is possible that the apolar region is important in the binding of lipoamide while the determinants for binding the larger glutathione molecule are elsewhere. One would predict that ionic interactions play a role in the binding of anionic glutathione and the nearby lysine residues may function in this way. Since the two enzymes share a common intermediate in catalysis, it seems reasonable to suggest that the structure of the disulfide region confers a special reactivity on that disulfide. The resolved spectrum of FAD when bound to glutathione reductase and lipoamide dehydrogenase indicates that the flavin is bound in a hydrophobic milieu. It has been suggested for lipoamide dehydrogenase (33) that the tight loop in the polypeptide chain, imposed by the’proximity of the half-cystines in the sequence, forms part of this milieu. The tight loop would favor a flat conformation accommodating the planar isoalloxazine ring and allowing multiple van der Waals contacts. Electron sharing between the flavin and the disulfide demands that they be close. There is a growing list of enzyme families in which a high degree of homology has been demonstrated around a common active site residue (89). These include the serine proteases, the cysteine proteases, the carboxypeptidases, the ATP-guanidine phosphotransferases (creatine, arginine, and lombricine kinases) , and the aldolases. The pyridine nucleotidedisulfide oxidoreductases can now be added to this list. A parallel can be drawn between this group and the aldolases where the closely related aldolases A and B show extensive homology but do not show homology with transaldolase. Such is the case with glutathione reductase and lipoamide dehydrogenase having a high degree of homology but lacking homology with thioredoxin reductase around the active center cystine residue. 88. A. L. Fluharty, G. I. Adelson, and B. P. Gaber, ABB 134, 346 (1969). 89. M. 0. Dayhoff, “Atlas of Protein Sequence and Structure,” Vol. 5 , p. 56. Nat. Biomed. Res. Found., Washington, D. C., 1972.

106

CHARLES H. WILLIAMS, JR.

111. Lipoamide Dehydrogenase

Lipoamide dehydrogenase has been isolated from many species and these are listed in Table I1 (4, 56, 90-115). The mammalian enzymes are of mitochondria1 origin. Where tested, the enzyme from eukaryotic sources is isozymic, while that from prokaryotes is a single species (105). There are marked differences in the sensitivity of the enzyme as isolated from various sources to inhibition by excess NADH in the NADH + lipS,NH, reaction and in the relief of that inhibition by NAD’ (27, 99, 105, 106, 108, 109, 116, 117). Attempts to make phylogenetic 90. F. B. Straub, BJ 33,787 (1939). 91. C. J. Lusty and T. P. Singer, JBC 239,3733 (1964). 92. T. C. Linn, J. W. Pelley, F. .H. Pettit, F. Hucho, D. D. Randall, and L. J. Reed, ABB 148, 343 (1972). 93. C. J. Lusty, JBC 238,3443 (1963). 94. S. A. Millard, A. Kubose, and E. M. Gal, JBC 244, 2511 (1969). 95. J. K. Reed, JBC 248, 4834 (1973). 96. S. Ide, T. Hayakawa, K. Okabe, and M. Koike, JBC 242, 54 (1967). 97. C. A. Eberhard, A. H. Guindon, C. Kepler, V. Massey, and C. Veeger, Biol. Bull. 123, 480 (1962). 98. D. K. Basu and D. P. Burma, JBC 235,509 (1960). 99. J. Matthews and L. J. Reed, JBC 238,1869 (1963). 100. L. L. Poulsen and R. T. Wedding, JBC 245,5709 (1970). 101. A. Wren and V. Massey, BBA 110,329 (1965). 102. E. Misaka, Y. Kawahara, and K. Nakanishi, J . Biochem. ( T o k y o ) 58, 436 (1965). 103. C. H. Williams, Jr., unpublished observations. 104. Y. Kawahara, E. Misaka, and K. Nakanishi, J . Biochem. ( T o k y o ) 83, 77 (1988). 105. W. H. Scouten and I. R. McManus, BBA 227, 248 (1971). 106. H. Fehrmann and C. Veeger, BBA 350, 292 (1974). 107. 0. Vogel and U. Henning, Eur. J. Bioehem. 35,307 (1973). 108. C. H. Williams, Jr., JBC 240,4793 (1965). 109. M. Koike, P. C. Shah, and L. J. Reed, JBC 235, 1939 (1960). 110. G. W. Notani and I. C. Gunsaius, Abstr. Pap., 134th Meet., Amer. Chem. SOC.Div. Biol. Chem., p. 3C (1958). 111. D. S. Goldman, BBA 32, 80 (1959). 112. M. L. Baginsky and F. M. Huennekens, ABB 120, 703 (1967). 113. S. M. Klein and R. D. Sagers, JBC 242,297 and 301 (1967). 114. C. Veeger, J. Krul, T. W.Bresters, H. Haaker, J. H. Wassink, J. S. Santema,

and A. DeKok, in “Enzymes, Structure and Function” (J. Drenth, R. A. Osterbaan, and C. Veeger, eds.), Proc. 8th FEBS Meet., p. 217. North-Holland Publ., Amsterdam, 1972. 115. I. C. Gunsalus, in “The Mechanism of Enzyme Action” (W. D. McElroy and B. Glass, eds.), p. 545. Johns Hopkins Univ. Press, Baltimore, Maryland, 1954. 116. R. L. Searls and D. R . Sanadi, JBC 238,680 (1961). 117. A. Wren and V. Massey, BBA 122,436 (1966).

3.

107

FLAVIN-CONTAINING DEHYDROGENASES

TABLE I1 SOURCESFROM WHICHLIPOAMIDE DEHYDROGENASE HAS BEENISOL~~TSD Eukaryotes

Ref.

Pig heart Beef heart Beef liver Pig brain Beef kidney Rat liver Human liver Dogfish liver Spinach leaves Cauliflower floral heads Saccharomyces eerevisiae Saccharomyces oviformis Torula Candida krusei Ncurospora crassa Pythium ultimum Phytophtora erythroseptica

4190 91,98 93 94 92 96 96 97 98,99 100 101 102 10s 104 106 106 106

1

Prokaryotes

Ref.

Escherichia coli B Escherichia coli K12 Escherichia coli M191-6 Escherichia coli Crookes Leuconostoc mesenteroides Mycobacterium tuberculosis Peptococcus glycinophilus Azotobacter vinelandii Proteus vulgaris Streptococcus faecalis Serratia marcescens Pseudomonas jluorescens Bacillus subtilis Azotobacter agilis

36 107 108 109 110 111 118,113 114 116 116 106 106 106 106

distinctions based on these differences are perhaps somewhat premature (105).The complications in such attempts lie in the fact that the sensitivity and its relief are dependent on the pH and in the fact that lipoamide dehydrogenase from some species binds NAD' very tightly SO that the NAD' present in most commercial NADH may be sufficient to activate the enzyme. There are also marked differences in the stability of the 2-electron-reduced enzyme (EH,, the catalytically important intermediate) in the presence of excess NADH, anaerobically in the absence of an acceptor. Three categories can be distinguished: very stable, moderately stable, and unstable. Enzyme isolated from mammals falls into the first category (24, 26, 93) ; enzyme from lower eukaryotes (99, 101, 106) and from an anaerobic bacterium (11%)falls into the second category; while enzyme from other prokaryotes, facultative (108) and aerobic (114) bacteria, falls into the third category. Differentiation between the first and second categories has been made difficult by more recent findings (106, 118), but spectral details will be discussed in a later section.

FUNCTIONS A. METABOLIC Lipoamide dehydrogenase, as its more proper name dihydrolipoamide dehydrogenase implies, functions physiologically in the reoxidation of 118. C. Veeger and V. Massey, BBA 67,679 (1963).

108

CHARLES H. WILLIAMS, JR.

dihydrolipoic acid, bound in amide linkage to the r-amino group of a lysine residue in the transacetylase or transsuccinylase (41, 43) ; the electron acceptor is NAD+ (18, 19). The lipoic acid is reduced as a consequence of the thiamine pyrophosphate-dependent oxidative decarboxylation of a-keto acids, pyruvate and a-ketoglutarate, yielding acetyl-CoA and succinyl-CoA, respectively, as shown below for pyruvate. CH,-CO-COO-

+ E,-TPP'-

E,-TPP'--CO-CH,+

-

t\-ip NH-E,

E,-TPPa-

s-s lip-NH-E, I \

HS S-CO-CH,

+ CoA-SH

+ CO~

E,-TPPS--CO-CH,

lip-NH-E, I \

+

d

ip-NH-E,

(2) (3)

8-co-CH,

+ CoA-S-CO-CH,

(4)

HS SH

Lipoamide dehydrogenase can be isolated from the multienzyme complexes that carry out these oxidative decarboxylations : from the a-ketoglutarate dehydrogenase complex of heart (41,119-181) or E. coli ( l d b ) , or from the pyruvate dehydrogenase complex of heart (98, 183, 184) or E. coli (107, 186). The complexes have recently been reviewed (186'). It is possible that lipoamide dehydrogenase also functions in the complexes that oxidatively decarboxylate the a-keto acids resulting from the transamination of valine, isoleucine, and leucine but these have proved difficult to resolve (1a7). Lipoamide dehydrogenase also functions in the pyridoxal phosphate and tetrahydrofolate-dependent oxidative decarboxylation of glycine in the anaerobic bacterium Peptococcus glycinophilus. The reaction in which the protein-bound lipoic acid is reduced is very complex and not yet fully understood; the ultimate electron acceptor is NAD+ (118, 113,128). Lipoamide dehydrogenase is quite easily dissociated from the a-keto 119. R. L. Searls and D. R. Sanadi, JRC 235,2485 (1960). 120. N. Tanaka, K. Koike, M. Hamada, K. Otsuka, T. Sueniatsu, and M. Koike, JBC 247, 4043 (1972). 121. F. H. Pettit, L. Hamilton, P. Munk, G. Namihira, M. H. Eley, C. R. Willms, and L. J. Reed, JBC 248,5282 (1973). 122. B. B. Mukherjee, J. Matthews, D. L. Homey, and L. J. Reed, JBC 240, PC2268 (1965). 123. T. Hayakawa and M. Koike, JBC 242, 1356 (1967). 124. T. Hayakawa, T. Kanzaki, T. Kitamura, Y. Fukuyoski;Y. Sakurai, K. Koike, T. Suematsu, and M. Koike, JBC 244,3660 (1969). 125. M. Koike, L. J. Reed, and W. R. Carroll, JBC 238,30 (1963). 126. L. J. Reed, Accounts Chem. Res. 7,40 (1974). 127. Y. Namba, K. Yoshizawa, A. Ejima, T. Hayaski, and T. Kaneda, JBC 244, 4437 (1969). 128. J. R. Robinson, S. M. Klein, and R. D. Sagers, JBC 248, 5319 (1973).

3.

FLAVIN-CONTAINING DEHYDROGENASES

109

acid dehydrogenase complexes (129) ; indeed, even when the complexes are carefully prepared, free flavoprotein is present (91). It is felt that this free enzyme arises chiefly from the pyruvate dehydrogenase complex (199,130).The physiological significance, number, and origin of the electrophoretically separable forms of heart and liver lipoamide dehydrogenase have been topics of considerable controversy since their original observation (131). The most appealing (i.e., the simplest) theory is that there are two major forms, one arising from each of the major a-keto acid dehydrogenase complexes (132, 133), and that other forms come from proteolytic degradation during isolation (64, 134) ; but this hypothesis may be too simple to explain all the facts (65, 135) and has been sharply criticized (130).While numbers of isozymes as high as thirteen have been reported (136),the most commonly quoted number is six (65, 93, 131, 134, 135, 137). Chromatography on anion exchangers resolves the mixture into two major bands (65, 93, 132, 133, 136) with further resolution of the more anionic band if shallow gradients are employed (93).The more anionic species seem to be associated with the a-ketoglutarate dehydrogenase complex and the less anionic species with the pyruvate dehydrogenase complex (65,130,132,133,135).Attempts have been made using a wide variety of physical and chemical methods, other than electrophoretic mobility, to demonstrate differences between the isozymes; with the possible exception of the 280:455-nm ratios of 5.5 for the less anionic species and 4.9 for the more anionic species (661,the differences fall within the limits of experimental error (130-136, 138, 139). Despite the sizable differences in effective charge, there is a generally 129. V. Massey, “Methods in Enzymology,” Vol. 9,p. 272, 1966. 130. W.C.Kenney, D. Zakim, P. K. Hoguem, and T. P. Singer, Eur. J. Biochem. 28, 253 (1972). 131. M. R. Atkinson, M. Dixon, and J. M. Thornber, BJ 82, 29P (1962). 132. Y. Sakurai, Y. Fukuyoski, M. Hamada, T. Hayakawa, and M. Koike, JBC 245, 4453 (1970). 133. T. Hayakawa, Y. Sakurai, T. Aikawa, Y. Fukuyoski, and M. Koike, in “Flavins and Flavoproteins” (K. Yagi, ed.), Vol. 2, p. 99. Univ. Park Press, Baltimore, Maryland, 1908. 134. J. E. Wilson, “Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 313. Univ. Park Press, Baltimore, Maryland, 1971. 135. M. L.Cohn, L. Wang, W. Scouten, and I. R. McManus, BBA 159, 185 (1968). 130. A. M.Stein and J. H. Stein, Biochemistry 4, 1491 (1965). 137. V. Massey and Q,. H. Gibson, Proc. Int. Congr. Biochem., 6th, 1961 Vol. 1, p. 157 (1963). 138. Y. Sakurai, T.Hayakawa, Y. Fukuyoski, and M. Koike, J. Biochem. (Tokyo) 65, 313 (1909). 139. A. M. Stein, B. Wolf, and J. H. Stein, Biochemistry 4, 1500 (1965).

110

CHARLES -H. WILLIAMS, JR.

held opinion that a single structural gene codes for lipoamide dehydrogenase. I n mammals this assumption is based on the fact that the enzyme isolated from either complex can be used to reconstitute the other complex and on the lack of differences referrred to above (130, 13%).It has been suggested that the differences observed electrophoretically are induced when lipoamide dehydrogenase binds to the transacetylase or transsuccinylase (which serve structural as well as catalytic functions in the complexes) since the optical rotary dispersion and circular dichroism of the flavoprotein differ in the two complexes (132).These changes must be reversible, however, since they cross-react in the reconstitution of the complexes. In E . coli, lipoamide dehydrogenase isolated from one complex can serve in the reconstitution of the other and a mixture of the enzyme isolated from the two complexes is electrophoretically homogeneous ; on this basis a single structural gene was hypothesized (140). This has now been confirmed by chromosomal mapping in an extensive series of lipoamide dehydrogenase mutants of E . coli K12 (141-143). Very recently another physiological function has been suggested for lipoamide dehydrogenase in addition to the reoxidation of lipoic acid in a-keto acid oxidation. The observations (14.4)are these: Metabolic conditions which result in an increase in intramitochondrial GTP levels lead to the almost complete inhibition of oxidation of NAD+-linked substrates, while having no effect on FAD-linked substrates. This effect can be mimicked by inhibitors of lipoamide dehydrogenase such as arsenite and 5-methoxyindole-2-carboxylic acid. The latter compound is a very poor inhibitor, having a K, of about 3 mM (146).It must be emphasized that no direct effect of GTP on isolated lipoamide dehydrogenase can he demonstrated (14.4).It is hypothesized (144) that lipoamide dehydrogenase serves as a transhydrogenase between two pools of pyridine nucleotide, i.e., NADH produced by NAD+-linked substrates and NAD' available to the electron transport chain. The transhydrogenase activity of this enzyme has been well demonstrated (146, 147). This hypothesis has been criticized on metabolic grounds, but the results have not been refuted (148, 149). It should perhaps be treated as a very interesting but tentative hypothesis. 140. F. H. Pettit and L. J. Reed, Proc. Nut. Acad. Sci. U S . 58, 1126 (1967). 141. J. R. Guest and I. T. Creaghan, J . Gen. Microbiol. 75, 197 (1973). 142. J. R. Guest, J . Gen. Microbiol. 80,523 (1974). 143. J. R. Guest and I. T. Creaghan, J. Gen. Microbiol. 81, 237 (1974). 144. M. S. Olson and T. T. Allgyer, JBC 248,1582 and 1590 (1973). 145. J. Reed and H. A. Lardy, JBC 245, 5297 (1970). 146. M. M. Weber and N. 0. Kaplan, JBC 225,909 (1957). 147. A. M. Stein, B. R. Kaufman, and N. 0. Kaplan, BBRC 2, 354 (1960). 148. C. M. Smith, J . Bryla, and J . R. Williamson, JBC 249, 1497 (1974). 149. E. I. Walajtys, D. P. Gottesman, and J. R. Williamson, JBC 249, 1857 (1974).

3.

FLAVIN-CONTAINING DEHYDROGENASES

B. REVIEWOF

THE

111

MECHANISM OF MASSEY AND VEEGER

The mechanism proposed by Massey and Veeger (15,27), which was based in large measure on the work of Massey et al. (24), has served as the working hypothesis in the intervening 12 years. Chief feature of the mechanism was the 2-electron-reduced intermediate, EH,, with the electrons shared between the flavin and the reactive disulfide. I n catalysis the enzyme was reduced to EH, and reoxidized once in each catalytic cycle. An equilibrium of EH, (at a second site on EH, referred to as the “Y” site) with NAD’ was proposed as a side reaction because of the requirement for NAD’ in the oxidation of NADH by lipoamide a t low p H ; however, in the discussion the authors emphasized that while they had made this a side reaction in the scheme, it was likely that NAD’ was bound to the Y site in the catalytic cycle per se. Thus, the proposed mechanism was of the ping-pong type but with provision for the role of NAD+ (in NADH oxidation) in protecting the enzyme against overreduction when NAD’ was bound a t the Y site. I n addition to the catalytic mechanism, the scheme also proposed side reactions to account for the inhibited 4-electron-reduced state. The five lines of evidence which led to this scheme can be summarized as follows: 1. The two-electron stoichiometry of the reduction of the enzyme to an intermediate in which the flavin had spectral properties intermediate between those of FAD and FADH,. 2. The fact that the intermediate was formed and reoxidized in halfreactions at rates commensurate with the kinetics observed in the overall reaction. 3. The inhibition by arsenite preincubation in the presence of NADH and the spectral effects in the presence of arsenite (Section 11,B). 4. The requirement for NAD’ in the oxidation of NADH at low pH together with the four-electron reduction of the enzyme by NADH in the presence of NADase to an inactive form. 5. The parallel line kinetics found when one substrate was varied a t several levels of the second substrate. The data will be discussed briefly by way of introduction to the next three sections.

c. PROPERTIES OF

THE

2-ELECTRON-REDUCED ENZYME, EH,

The spectral characteristics of pig heart lipoamide dehydrogenase reduced anaerobically a t pH 7.6 and 2 5 O under the following conditions are virtually identical: 1 equivalent (1 mole/mole of enzyme FAD) of

112

CHARLES H. WILLIAMS, JR.

NADH with NADase present (66, Fig. 5) or excess dihydrolipoamide (24, Fig. 1 ) . The extinction coefficient of EH, is 3.05 mM-l cm-l a t 530 nm and 8.7 - 9.2 mM-' cm-l a t 445 nm. A large excess of NAD+ (100 equivalents) with a small excess of NADH (2 equivalents) leads to the formation of a distinct spectral species shown in Fig. 6 (118).Based on its high absorbance in the 450-nm region, it is postulated that this is a complex of NAD+ with EH, (118).Rapid reaction measurements show that it is formed in less than 3 msec and thus represents a potential catalytic intermediate (118). Large excesses of NADH lead to the formation of a complex with EH,. This complex was originally observed in glutathione reductase (62,63), but retrospective examination of spectra of lipoamide dehydrogenase indicates that it is present in this enzyme also (118, Fig. 2, control curve, 19 equivalents NADH and 160, Fig. 5, curve 2). The available data indicate that a tight complex of this type is formed in lipoamide dehydrogenase of Pythium ultimum (106), Peptococcus glycinophilus (112), and perhaps yeast (101) and spinach (99). The characteristics of this complex will be discussed in connection with glutathione reductase, but the diag-

NADHI-NAD rotio(fin0l)-

-

700

320 Wavelength (nm)

FIQ.6. Complex formation between the 2-electron-reduced form of pig heart liposmide dehydrogenase and NAD' (118).

3.

FLAVIN-CONTAINING DEHYDROGENASES

113

nostic sign of its formation is a drop in extinction in the 450-nm region without a proportional drop in the 530-nm region upon the addition of large excesses of NADH to EH,. I n some cases there is an increase in extinction a t 530 nm (150, Fig. 5 ) , and this is the case with glutathione reductase (52, 53). A number of conditions lead to the further reduction of EH,. I n the presence of NADase, excess NADH reduces the enzyme completely (four electrons) ( 9 7 ) . It has been argued that the protection of the enzyme by NAD' against four-electron reduction is the result of its binding a t the same site as would the second NADH molecule (27).An alternative explanation is that the NAD+ acts to reverse an already unfavorable equilibrium (151).The effective potential (reflecting true potential and substrate binding) a t pH 7.6 of the 2-electron-reduced enzyme-4-electron-reduced enzyme couple must be more negative than that of dihydrolipoamide-lipoamide (-325 mV, p H 7.6) ( 4 ) since EH, is stable in the presence of 20 equivalents of dihydrolipoamide (24). The potential of the NADH-NAD' couple a t this p H is -340 mV (152), i.e., only about 15 mV more negative than that of lipoate; thus, the removal of the product, NAD', is crucial if extensive reduction of EH, is to be observed a t pH 7.6. This is illustrated in a direct comparison of the rate of reduction of EH, by excess NADH or NADPH in the presence of NADase in lipoamide dehydrogenase and glutathione reductase, respectively. NADP' is much more tightly bound to the 4-electron-reduced glutathione reductase than is NAD+ to 4-electron-reduced lipoamide dehydrogenase ; and the rate of reduction of EH, in the former is much slower since NADP' or NAD' bound to their respective enzymes are not available to the hydrolytic action of NADase ( 2 9 , 5 2 , 5 3 ) .At pH 6.3 the separation of potential between dihydrolipoamide and NADH is much greater (ca. 55 mV). At this pH, about 20% reduction of EH, to the fully reduced enzyme is observed (24, Fig. 2) in a slow reaction, and this overreduction is prevented by NAD+ (137).This degree of overreduction implies that the effective potential of the 2-electron-reduced enzyme-4-electron-reduced enzyme couple is about 7 mV more negative than the NADHNAD' couple at pH 6.3 (-298 mV). In the presence of lipoate, i.e., in turnover, overreduction of EH, is rapid (137). Formation of EH, upon reduction with 1 equivalent of dithionite is only about 85% complete because of overreduction, and the enzyme is fully reduced by 2 equivalents of dithionite (59). Reduction of lipoamide dehydrogenase by NADH with arsenite present 150. L. Casola and V. Massey, JBC 241, 4985 (1966). 151. D. R. Sanadi, Proc. Int. Congr. Biochem., 6th, 1961 Vol. 5, p. 172 (1963). 152. F. L. Rodkey, JBC 213,777 (1955).

114

CHARLES H. WILLIAMS, JR.

has been discussed in Section I1,B. It leads to the production of a charge transfer complex between FADH, and NAD' that is characterized by a broad absorption band centered a t 700 nm imparting a green color to the enzyme (27). Similar results are observed if cadmium ion (26, 28, 163) or mercurials (160) are present or if the enzyme has been pretreated with cupric ion (164); however, the mechanisms by which these three reagents effect the conversion to the charge transfer complex are quite different: Cadmium ion acts like arsenite bridging the nascent thiols (26, 28, 153) ; mercurials react with one or both of the nascent thiols of the reactive disulfide (160) ; and cupric ion causes the oxidation of 2 thiols to a disulfide, causing sufficient strain to weaken (but not totally prevent) the interaction between the FAD and the sulfur in the 2-electron-reduced enzyme (166, 156). Details of the modifications by cupric ion, by cadmium ion, and by mercurials will be discussed in Section II1,F. This same species can be formed from EH,, which has been produced by the addition of 1 equivalent of NADH, simply by cooling the enzyme to 4 O ; the dismutation is slow, requiring more than 16 hr to form the equilibrium mixture of oxidized enzyme and the charge transfer complex (65). The stability of EH2 is very species dependent. All of the above results refer to the pig heart enzyme and, where tested, t o other mammalian species. It was initially reported that no long wavelength absorption was observed upon reduction of E. coli enzyme with NADH (109), but reduction by 1 equivalent of NADH or dihydrolipoamide leads to the formation of 25% of the maximal 2-electron-reduced species (108) and similar results are obtained with the Azotobacter enzyme (114). That this species is the catalytically important one in the E. coli enzyme as well as in the mammalian enzyme has also been demonstrated (60).Reduction with dihydrolipoamide in the rapid reaction spectrophotometer a t 2O results in the full formation of EH, followed by the slow (k = 13 min-l, 1 mM dihydrolipoamide) four-electron reduction. The spectrum of EH, generated in this way is shown in Fig. 7 and is identical with that of the pig heart enzyme. The 2-electron-reduced form, EH2 of lipoamide dehydrogenase of spinach (99) may be somewhat unstable; however, spectrally it is difficult to distinguish between instability and formation of the EH,NADH complex (see above) on the basis of available spectral data. Either phenomenon could lead to inhibition by excess NADH. I n glutathione reductase it is possible that the complex can be rapidly reoxidized by glutathione (65), 153. 154. 155. 156.

A. M. Stein and J. H. Stein, JBC 248,670 (1971). C. Veeger and V. Massey, BBA 64, 83 (1962) ; 37, 181 (1960). L. Casola, P. E. Brumby, and V. Massey, JBC 241,4977 (1988). R. G. Matthews and C. H. Williams, Jr., BBA 370, 39 (1974).

3.

FLAVIN-CONTAINING

DEHYDROGENASES

115

Wavelength (nm)

FIG.7. Escherichia coli lipoamide dehydrogenase. The spectrum of the oxidized enzyme ( A ) . The spectrum of the 2-electron-reduced enzyme (0) was generated from rapid reaction spectrophotometry as described in the text. The spectrum of the 4-electron-reduced enzyme (0) was produced by anaerobic reduction by 12 moles dihydrolipoamide/mole FAD.

D. KINETICSTUDIES The kinetics of the half-reactions for pig heart lipoamide dehydrogenase, i.e., the conversion of enzyme to EH, by NADH or dihydrolipoamide and the reoxidation of EH, by NAD' or lipoamide derivatives, have been measured by rapid reaction spectrophotometry (24, 137). Reduction of the enzyme by NADH and reoxidation of EH, by NAD' are complete in the dead time of the instrument which is 3 msec. The rate of reduction of the enzyme by dihydrolipoamide is rate determining in the overall reaction and is 33,000 min-I a t infinite reductant conceptration; the same rate is determined by conventional kinetics a t infinite concentration of both substrates ( 2 4 ) . Initial velocity patterns obtained for the reduction of NAD' by dihydroplipoamide give a series of parallel lines (reciprocal plots). The K,,,

116

CHARLES H. WILLIAMS, JR.

for NAD- is 0.2 mM, and the K,, for dihydrolipoamide is 0.3. mM. On the basis of these data the authors proposed what would now be referred to as a bi-bi ping-pong mechanism (94). This can be represented as follows, where E is the oxidized enzyme, EH, is the 2-electron-reduced enzyme, lip (SH) ,NHz is dihydrolipoamide, and lipS,NH, is lipoamide: ki

E

+ lip(SH)zNH2ekr [E-lip(SH)gNHZ

EHz-~~~SZNHZ]

(5)

kr

+ lipS2NHz E-NADH] e E + NADH kn

[E-lip(SH)nNHze EHrlipS~NH~] e EHz kr

kr

EHz

+ NAD+ e [EHz-NAD+ ks

(6)

kr

(7,8)

The pH optimum of the pig heart lipoamide dehydrogenase in the direction of NAD+ reduction by dihydrolipoamide is 7.9 ( 4 ) .I n the direction of NADH oxidation by lipoamide the pH optimum is 6.5 ( 4 ) . I n this latter direction there is an absolute requirement for NAD+ a t the pH optimum (971, but this requirement disappears as the pH is raised (116). It is therefore crucial to be aware of the pH of the measurements in comparing kinetic data. A more recent examination of the kinetics of this enzyme by initial rate measurements has included product inhibition patterns and has led to the conclusion that a t least under some conditions an ordered bi-bi mechanism applies which involves a ternary complex of enzyme, NAD+, and dihydrolipoamide (167).Clear spectral evidence is presented for the existence of a complex between NAD+ and the oxidized enzyme and this will be discussed in Section II1,E. The product inhibition pattern for NAD+ tended toward that expected for this mechanism only at high NAD+ concentration. A third study of the kinetics of lipoamide dehydrogenase has utilized the enzyme isolated from rat liver (96). At 25O, the temperature of the two previous studies, when dihydrolipoamide was varied a t fixed levels of NAD+, the double reciprocal plots were concave down. At 3 7 O this behavior was not observed. The detailed studies were carried out a t the higher temperature. Rates were measured in both directions a t p H 8.0, the pH optimum for the reduction of NAD+. Under these conditions, initial velocity patterns for the forward and reverse reactions were a series of parallel lines. The K, for NAD+ was 0.52 mM, for dihydrolipoamide was 0.49 mM, for NADH was 0.062 mM, and for lipoamide was 0.84 mM. The maximum rate for NAD+ reduction was 20,700 min-*/FAD 157. J. Viaser, H. Voetberg, and C. Veeger, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H.Sund, ed.), p. 359. Springer-Verlag, Berlin and New York, 1970.

3.

FLAVIN-CONTAINING DEHYDROGENASES

117

and for NADH oxidation was 7,500 min-’/FAD. These data were consistent with a bi-bi ping-pong mechanism. Further support for this mechanism came from rates of exchange between 14C-NAD+and NADH; the time course of this exchange was unaltered by the addition of a mixture of lipoamide and dihydrolipoamide. The substrate inhibition patterns were complex and a t variance with the previous study (157).Dependence of pattern type on the concentration of the fixed substrate was observed. The overall pattern was consistent with a “dead end” inhibition product of EH,-NADH. This may prove to be a most interesting result in view of the spectral indication of such a complex (see Section II1,C). The kinetics of the yeast lipoamide dehydrogenase in the direction of NAD’ reduction indicate a bi-bi ping-pong mechanism is operative in this species also (117).If the enzyme from yeast indeed proves to have a tighter EH,-NADH complex than does the mammalian enzyme, product inhibitions studies should show impressive dependence on the fixed substrate (96). The kinetic studies cited above have assumed that in lipoamide dehydrogenase the two active centers are independent of one another. A recent paper has indicated homotropic regulation is operative under some conditions (157,158).

E. ROLEOF NADt AS A MODIFIER The requirement for NAD+ in the oxidation of NADH by lipoic acid derivatives a t low pH led to the proposal of a second binding site, referred to as the ‘‘Y site,” for pyridine nucleotide in lipoamide dehydrogenase (97).The argument that NADt acted in preventing overreduction simply by reversing the unfavorable equilibrium between EH, and the 4-electron-reduced enzyme (161) rather than by combining a t a second site has been discussed above (see Section 111,C). Several other lines of evidence have been advances which are consistent with two pyridine nucleotide sites, but none of these leads unequivocally to the conclusion that both sites are involved in catalysis. The matter is complicated by the fact that the binding constant of a second site might be high on EH2 but low on the oxidized enzyme. Some of the experiments to be outlined have dealt only with the oxidized enzyme. Binding of NAD+ to the oxidized enzyme causes small changes both in the visible (157,159,160) and in the fluorescence spectra (161). These 158. C. S.Tsai, ABB 159,453 (1973). 159. A. M. Stein and G . Czerlinski, Fed. Proc., Fed. Amer. SOC.Exp. Biol. a, 842 (1967). 180. H. Muiswinkel-Voetberg and C. Veeger, Eur. J . Biochem. 33, 285 (1973). 161. G. Su and J. E. Wilson, ABB 143,253 (1971).

118

CHARLES H. WILLIAMS, JR.

changes are highly temperature-dependent, and in this connection the temperature dependence of the kinetics (96) (Section II1,D) should be recalled. Although the fluorescence changes are small, the sensitivity of the method allowed usable measurements down to NAD' concentrations of 125 p M (161). At 5" and a t 25O plots of reciprocal fluorescence change vs. reciprocal NAD+ concentration are clearly biphasic while a t 37O they are monophasic. At 25O K D values of 0.2 and 0.9 mM were estimated; a t 37O the K D was 1.7 mM. The data were interpreted as indicating two sites for NAD+a t the lower temperatures. Changes in the visible spectrum upon NAD+ binding are not sufficiently large to allow measurements a t low NAD' concentrations (167, 160). Maxima (positive) in the difference spectrum were observed a t 505 and 387 nm and minima (negative) at 477,450,430, and 370 nm. The relative extinction changes varied with temperature. The largest change was a t 450 nm and was about 0.34 mM-' cm-l. The binding curves were not regular hyperbolas, and dissociation constants were evaluated from Stockell plots. These gave dissociation constants of 50 and 200 pM a t 25O for the changes a t 430 nm and indicated more than one site. The lowest concentration used in these experiments was 100 ~LM. The disparity between the two sets of dissociation constants has been discussed by both groups (160, 161).

Alkylation of native pig heart lipoamide dehydrogenase by concentrated iodoacetamide results in a 95% decrease in the NADH-lipoamide reductase activity and in the loss of the ability to form the charge transfer complex between NAD' and 4-electron-reduced enzyme (163). Since the formation of this complex is absolutely dependent on the presence of NAD', the authors interpreted their results as indicating that the alkyation had destroyed the NAD' binding site by reaction with a thiol. The groups alkylated were not characterized. It was indicated that the modified enzyme could be reduced to EH, but that further reduction resulted with excess NADH (153). Subsequently, enzyme modified by concentrated iodoacetamide has been characterized more fully (166). Two thiols and one methionine were alkylated. The more slowly reacting of the two thiols could be protected by high concentrations of NAD'. This thiol was identified; of the 7-8 thiols in the pig heart enzyme, 7 have been identified by association with unique sequences (63, 86) and these will be discussed in the next section. It was shown that all of the modified enzyme could be reduced to EH, by NADH thus indicating that the 5% residual activity was not resulting from 5% unmodified enzyme but rather from an inherent activity of the modified enzyme (166). Further, this suggested that the substrate binding site was not the site affected by alkylation. Since the modified enzyme could no longer form the charge

3.

FLAVIN-CONTAINING DEHYDROGENASES

119

transfer complex with NAD' in the 4-electron-reduced state, the possibility was presented that a second pyridine nucleotide binding site was that affected by alkylation (156).In view of the marked lowering of activity these data do not preclude modification of the substrate binding site. If indeed there is but one pyridine nucleotide binding site, the modification may only prevent the exact alignment necessary for charge transfer. Recent findings have clarified the role of NAD' in the activation of NADH oxidation by lipoic acid (160). It was found that NAD' shifted the pH optimum of this reaction from 6.2 to 5.0 and that NAD+ binding was associated with the release of a proton. It was concluded that NAD' binding lowered the pK of a group essential for activity in its deprotonated form, and the suggestion was made that the group might be a thiol (160). If this is the case, i t is almost certainly the thiol which, when alkylated, prevents charge transfer between NAD' and the 4-electronreduced enzyme. The apparent pK (6.6) of the group(s) giving rise to the high pH arm of the pH vs. activity curve did not change upon "AD+ binding (160).The authors speculated on the identity of this latter group suggesting that it might be the lysine residue near the active center disulfide (Fig. 5 ) , pointing out that the pK was low for a lysine residue but not without precedent. This apparent pK should be ascribed to factors having to do specifically with the reaction in the direction of NADH oxidation when lipoic acid is the acceptor. If lipoamide is the acceptor, this pK shifts to a higher value, ca. 7.5. Furthermore, in the direction of NAD' reduction, a still higher but indeterminant pK is seen ( 9 5 ) . A requirement for a lysine residue (or the terminal amino group) in the charged configuration would be very interesting since electrophilic catalysis can be an enhancement factor in thiol-disulfide interchange (162) which is the first step in the reoxidation of the enzyme. I n this role i t could, a t the same time, stabilize the increasing negative charge on the substrate and the decreasing negative change on the enzyme. A representation of a possible transition state is shown in Fig. 8A. Electrophilic catalysis is not necessary in the direction of dihydrolipoamide oxidation as shown in Fig. 8B, since the nascent thiolate anion is stabilized in EH,. But the same residue, a t the higher pH optimal for the reaction in this direction, might be expected to be largely uncharged and in this state serve as an excellent general base in the formation of a thiolate anion on the substrate (Fig. 8B). Lysine residues have been found near the disulfide in E. coli (33) and pig heart (63, 85, 86) lipoamide dehydrogenase, in yeast glutathione reductase (35), and in thioredoxin ( 4 5 , 4 7 ) . 162. D. S. Garwood and D. C. Garwood, J. Org. Chem. 37, 3804 (1972).

CHARLES H. WILLIAMS,

120 (A1

JR.

(8)

0

-S -FAD)

pH 6.3

pH 8.0

FIG.8. (A) Hypothetical transition state in the electrophilic catalysis of thiol-disulfide interchange. (B) General acid-base catalysis of thiol-disulfide interchange by a hypothetical amino group.

Thus far a modifier role for NAD+has been discussed only in the direction of NADH oxidation. Recent studies suggest that NAD+ may also have a double role in the direction of NAD+ reduction by dihydrolipoamide (163).

F. STRUCTURAL STUDIES The total half-cystine content of pig heart lipoamide dehydrogenase is 10 per FAD (63). The basis for the protein quantitation has been discussed in Section I1,C. Older data suggested that there were 2 cystine residues and 6 cysteine residues (83),but more recent data (61,63) give strong evidence that the active center cystine residue is the only cystine residue. Only 7 thiols react with DTNB under denaturing conditions (63); however, recalculation of data of Brown and Perham (86),taking 51,000 as the molecular weight, indicates that under reducing and denaturing conditions 10 thiols are alkylated by iodoacetate. Thus it would appear that the enzyme contains 8 thiols (one of which is very unreactive) and the active center disulfide. Combining the data of two laboratories, Table I11 shows that unique compositions (and in some cases sequences) are associated with 7 of the presumed 8 thiols. The thiol con163. C. Veeger, H. Voetberg, J. Pronk, and A. J. W. G. Visser, in “Structure and Function of Oxidation Reduction Enzymes” (A. Akeson and A. Ehrenberg, eds.), p. 476. Pergamon, Oxford, 1972.

TABLE I11 SEQUENCESAROUND THE RE.4CTIVE THIOLS" Peptide 51 18

0.00

DTC4a

0.00

0.00

20

DTC2n2 55

DTC2b

Sequence

RI

7

(3

-

Leu(Val,Cys(Cm),Ile,Gly)Arg 1 Val-Q@Cm)-IlyGly-Ar$ V+-Cy~(Cm)-IlfGly-Ar$

+0.14 4-0.15

Val(Cys(Cm),His,Ala,His,Pro, Thr,Ser,Glx,Ala,Leu,Phe)Arg

-0.23 -0.27

Thr (Vtl,Cy!

~-C~(Cm)-HL-Al~-H~-Pro(Thr,Ser,Glx,Ala,Leu,Phe)Arg

(Cm),Ile, Glx )Lys, Th-Val-Cys (Cm)-Ile-Glu-Lys

cc 0 D

m

DTC2a2

-0.51 -0.52

Tyr(Ser,Glu,Ala,Leu,Gln,Gly,Asn,Gly,Ala,Ser,Cys(Cm),Glu,Asp,Ile,Ala)Arg 2 (Gln,Gly,Asn,Gly,Ala,Ser,Cys(Cm),Glu,Asp,Iie,Ala)Arg Gly,Ala-Ser,Cys(Crn)-Gl~-Aslj-Il~-Al~-Ar~

52

-0.30

(Cys(Cm),Asp,Ser,Pro,VaL,Ile,Tyr)

10

8

7777

No corresponding peptide DTCla

-0.87

Ala-Glx-Asx-Glx-Gly-Ile(Cys(Cm),Glx,Gly,Val,Met) 777777

No corresponding peptide DTClb3

-0.63

Ala-Gly-Val-Ile-Thr-Cys (Cm)-Asp-Val-Leu-Leu 7 7 7 7 7 -

1 7 7 1

No corresponding peptide Composition and sequence data (63,86).Simple numbers for peptides refer to Matthews et al. (63),while alpha-numeric designations refer to Brown and Perham (86). -,represents a residue placed by the dansyl-Edman or the subtractive-Edman procedures. R, refers to peptide mobility on electrophoresis at pH 6.5 relative to the mobility of aspartic acid (-1.0). 0

CL

r?

122

CHARLES H. WILLIAMS, JR.

tained in peptides 18, 51, and DTC4a is the one thought to be protected by NAD' from alkylation by concentrated iodoacetamide (156) (see Section 111,E). The thiols in the native enzyme are remarkable unreactive except with mercurials (150, 164) and with cupric ion (154, 155, 165). Two thiols react rapidly with phenyl mercuric acetate and 2 more slowly. Reduction of mercurial treated enzyme (after removal of excess reagent) by NADH results in the migration of phenyl mercury to the nascent thiols of the active center (150). Treatment of native pig heart lipoamide dehydrogenase with cupric ion leads to loss of lipoate-linked activities and to a marked increase (10- to 30-fold) in the NADH-DCI activity. Concomitantly there is a drop of 2 in the number of titratable thiols. The action of cupric ion is catalytic (154, 155). Amperometric titration in the presence of urea before and after addition of Eulfite indicates that the cupric ion-treated enzyme contains one disulfide in addition to the active center disulfide (155).Sulfite reacts with disulfides as follows: RS-S-R'

+ SOa*-

F! RSSOa-

+ R'S-

Thus, an increase in the thiol titer of one upon treatment with sulfite indicates the reaction of one disulfide. The thiols involved in the formation of this disulfide are contained in peptides designated 20 and 8/10 in Table 111 (156). It is of possible interest that peptide 20 contains 2 histidine residues, and it has been suggested that one or both of these bind cupric ion prior to its catalysis of the formation of the disulfide bond (156).Formation of the disulfide bond and the changes in the catalytic activities can be reversed by dialysis against cysteine provided the cupric ion treatment is not prolonged (155). If cupric ion is removed after the rapid changes have occurred, and the enzyme stored a t Oo for long periods (up to 7 months) , further oxidation of thiols appears to take place. These changes are also reversed by treatment with cysteine and are thus distinct from those observed upon prolonged reaction with cupric ion which are not reversible by cysteine (155).This result may indicate that a cluster of thiols exists and that these can oxidize sIowly in enzyme already containing a disulfide between thiols 8/10 and 20. The properties of the enzyme treated briefly with cupric ion, other than the marked changes in catalytic activities, would indicate that the formation of the disulfide bond does not markedly alter the enzyme. Thus, in the optical spectrum the peak in the visible is blue-shifted only 3 nm; 164. M. Nakamura and I. Yamazaki, BBA 267,249 (1972). 165. E. Misaka and K. Nakanishi, J. Biochem. (Tokyo) 80, 17 (1966);59, 545 (1966).

3.

FLAVIN-CONTAINING DEHYDROGENASES

123

the fluorescence excitation spectrum of the enzyme-bound FAD is unaffected; and the fluorescence emission spectrum of the aromatic amino acid residues is unaffected (155). Moreover, the enzyme is rapidly reduced to EH, (the 2-electron-reduced form) by NADH and by dihydrolipoamide, though reduction with the latter agent is not as rapid as in the native enzyme. However EH, is not stable in the presence of excess reductant and slow further reduction ensues leading, when NAD' is present, to the charge transfer interaction (154). Prolonged treatment with cupric ion leads to secondary changes as evidenced by the loss of the high NADH-DCI activity and large decreases in the fluorescence exitation spectrum of enzyme-bound FAD and to large increases in the fluorescence emission spectrum of the aromatic amino acid residues (155). The thiols in native lipoamide dehydrogenase are remarkably unreactive with other reagents; only one thiol is a t all reactive with DTNB or iodoacetate (61). Formation of the TNB-enzyme mixed disulfide is greatly increased by low concentrations (0.7 M ) of guanidine hydrochloride (166). Its modification is associated with the destabilization of the enzyme in 1 M guanidine hydrochloride which results in the slow reaction of 6 additional thiols. If the denaturant and excess DTNB are removed when the single thiol has reacted, the spectrum of enzyme-bound FAD is unmodified and the enzyme retains almost full activity. It is concluded that the thiol and the FAD are remote from one another in the protein (166).

Another facet of the reversible denaturation of the pig heart enzyme by concentrations of guanidine hydrochloride up to 1 M is the effect of the perturbant on activity and on the FAD spectrum (166). The difference spectrum in the visible and near ultraviolet between enzyme in guanidine hydrochloride and enzyme in an equal concentration of sodium chloride arises from a generalized red shift and a drop in extinction a t 455 nm. The changes are complete within the 3-msec dead time of the rapid reaction spectrophotometer. The difference spectra are reminiscent of, but some 5-fold larger than, those effected by temperature (167). The activity in the direction of lipoamide reduction falls to 20741,while in the direction of dihydrolipoamide oxidation it falls to 10% a t 1.0 M denaturant; the NADH-DCI activity rises slightly. All of the changes are reversible (166). Slightly higher concentrations of guanidine hydrochloride lead to the dissociation of the FAD (168). The effect of sodium dodecyl sulfate on the activity, fluorescence spectrum, fluorescence polarization, circular dichromisms, and sedimentation coefficient have been 166. C. Thorpe and C. H. Williams, Jr., Biochemistry 13, 3263 (1974). 167. F. Muller, S. G . Mayhew, and V. Massey, Biochemktry 12, 4654 (1973). 168. A. H. Brady and S. Beychok, JBC 244,4634 (1969).

124

CHARLES H. WILLIAMS) JR.

reported (169).Binding of this agent to the protein takes place in two phases, the first of which can be reversed by cooling to Oo. The effects on activity in the NADH-lipoate reaction are dependent on a group with pK of 6.6. At higher concentrations dissociation of the FAD and dissociation to the monomer are observed (169). Lipoamide dehydrogenase is quite stable in 6.5 M urea provided it is not frozen or reduced; full activity is regained upon dilution (32, 170). However, if reduced with NADH in 6.5 M urea, EH,, which is initially formed, is further reduced over a period of a few hours. The activity remaining upon dilution during this period is directly proportional to the amount of EH, remaining a t the time of dilution. NAD’ stabilizes EH, in 6.5 M urea just as it does in the absence of urea. When reduction in urea is allowed to go to completion, the spectrum upon reoxidation is that of free FAD indicating dissociation as a result of reductive denaturation (82, 170). The apoenzyme of pig heart lipoamide dehydrogenase has been prepared by two quite different procedures. Both forms of the apoenzyme can be reactivated by FAD, but they differ from one another in several respects. Apoenzyme prepared by precipitation once with ammonium sulfate at low pH in the presence of a high concentration of monovalent anions (171)is a monomer, i.e., molecular weight 52,000 (172-174).This preparation is moderately stable at Oo. It has less than 10% of the original FAD, less than 5% of the original NADH-lipoate activity, and about 90% of the original NADH-DCI activity ; the latter activity probably results from the residual FAD turning over a t a rapid rate. The yield of apoenzyme in this preparation is inversely dependent on the protein concentration. The fluorescence yield of the aromatic amino acids is enhanced in the apoenzyme (relative to that in the holoenzyme) , and excitation maxima are exhibited a t 284 and 290 nm (173-176).In the second procedure, apoenzyme is prepared by dialysis against 1.5 M guanidine hydrochloride, pH 7.6 to which F M N is added a t a concentration equal to that of the eneyme-bound FAD. The guanidine hydrochloride is removed by dialysis against F M N and finally the F M N is removed by 169. H.van Muiswinkel-Voetberg and C. Veeger, Eur. J . Biochem. 33, 279 (1973). 170. V. Massey, JBC 235, PC47 (1960). 171. P. Strittmatter, JBC 236, 2329 (1961). 172. C. Veeger, D. V. Dervartanian, J. F. Kalse, A. DeKok, and J. F. Koster, in “Flavins and Flavoproteins” (E. C. Slater, ed.), Vol. 1, p. 242. Elsevier, Amsterdam, 1966. 173. J. F.Kalse and C. Veeger, BBA 159,244 (1968). 174. C.Veeger, in “Flavins and Flavoproteins” (K. Yagi, ed.), Vol. 2, p. 252. Univ. Park Press, Baltimore, Maryland, 1908. 175. J. Visser and C. Veeger, BBA 206,224 (1970).

3.

FLAVIN-CONTAINING DEHYDROGENASES

125

dialysis against buffer. The F M N stabilizes the aproprotein in the presence of the gunnidine hydrochloride but binds only very loosely, if a t all (see below) (168).The apoenzyme prepared by this method seems to be dimeric (17 6 ) . The recombination of FAD with apoenzyme prepared by the acid ammonium sulfate method (monomer) is a multistep process (172-175). It can be divided into two phases though each is complex. I n the first phase FAD combines rapidly (half-time less than 2 min, 5O) with the monomeric apoprotein ; the product, still monomeric, initially gains (half-time also less than 2 min, 5 O ) a very high NADH-DCI activity and this decreases about 25% over a 5-min period to a level about 12 times that of the native enzyme. This species has very low NADH-lipoate activity. At 5O no second phase ensues; a t higher temperatures, however, dimerization leads to a fully active holoenzyme. The half-time for this process a t 2 5 O and a t apoprotein concentrations of 0.5-1.0 mg/ml is about 5 min. The return of NADH-lipoate activity is roughly parallel with the decrease in NADH-DCI activity to levels near those of native enzyme. The activation energy of the process is 15-20 kcal/mole. As would be expected, the rate of the second phase is dependent on protein concentration; it is optimal a t pH 7.2 and a t 0.2 M phosphate concentration. Careful examination of the properties of the reconstituted enzyme is possible only after removal of denatured protein by chromatography on calcium phosphate gel which also serves to remove excess FAD. Reconstitution of holoenzyme is inhibited by FMN, ADP, ATP, NAD’, and pyrophosphate, indicating that multiple linkages are involved in the very tight binding. Of the several flavin derivatives tested, only 3-methyl-FAD binds to give enzyme with appreciable NADH-lipoate activity (173-175, 177). Detailed curve fitting indicates that binding of FAD in lipoamide dehydrogenase introduces no new transitions, i.e., the same set of six Gaussian bands can be accounted for in free and bound FAD (178). A reversible dimer-monomer transition has been reported independent of FAD dissociation (174, 179-182). This transition seems t o be potentiated by the exhaustive removal of phosphate ion or by freezing of dilute 176. 177. 178. 179.

V. Massey, personal communication. J. Visser, D. B. McCormick, and C. Veeger, BBA 159, 257 (1968). A . H. Brady and S.Beychok, JBC 246,5498 (1971). J. Visser and C. Veeger, BBA 159, 265 (1968). 180. C. Veeger, H. Voetberg, J. Visser, G. E. J. Staal, and J. F. Koster, in ‘‘Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 261. Univ. Park Press, Baltimore, Maryland, 1971. 181. H. van Muiswinkel-Voetberg, J. Visser, and C. Veeger, Eur. J . Biochem. 33, 265 (1973). 182. H. van Muiswinkel-Voetberg and C. Veeger, Eur. J . Biochem. 33, 271 (1973).

126

CHARLES H. WILLIAMS, JR.

enzyme at low ionic strength; dimer is converted to monomer upon dilution following either treatment. The monomer is characterized by high NADH-DCI activity and low NADH-lipoate activity. Immediately following thawing of dilute enzyme frozen a t low ionic strength, the NADHlipoate activity is 75% of normal, the NADH-DCI activity is raised, and the peak in the visible is shifted from 455 to 452 nm. These three parameters return to normal overnight. The presence of bovine serum albumin, EDTA, or ammonium sulfate during freezing of dilute enzyme protect against the changes seen upon thawing. Enzyme potentiated toward dissociation to monomer can be stabilized by NAD+, by heating to 70°, by alkaline pH, or by 0.2 M phosphate at pH 7.2 (174,179-182). Lipoamide dehydrogenase from E . coli has been crystallized but there appears to be more than one dimer per asymmetric unit making X-ray structure determination difficult (183,184). The yeast enzyme has also been crystallized as long yellow needles approximately 20 p in width (104, 186). Measurements of fluorescence energy transfer have allowed estimation of the intramolecular distance between one of the two tryptophan residues of pig heart lipoamide dehydrogenase and the enzyme-bound FAD. This distance is found to be 13-16 A (186).Another very interesting estimation of distance by fluorescence energy transfer is that between the FAD of lipoamide dehydrogenase and the thiamine pyrophosphate of pyruvate dehydrogenase in the complex of E . coli (187).This distance is between 30 and 60 A. Lipoic acid, bound through amide linkage to the €-amino group of a lysine residue in the transacetylase, must make contact with three groups in the course of its reductive acetylation, acetyl transfer, and reoxidation [see Eqs. (2), ( 3 ) , and ( 4 ) ] . The lipoic acid itself, together with the side chain of the lysine residue, is about 14 A long (126). Thus, it would appear that small changes are required in the juxtaposition of the three enzymes within the complex as the dithiolane ring moves from one site to another (187).

G. SUMMARY AND CONCLUSIONS The chief feature of the mechanism of lipoamide dehydrogenase proposed by Massey and Veeger (16,27) is the catalytic intermediate in 183. D. DeRosier, personal communication. 184. J. Hainfeld, Ph.D. Dissertation, University of Texas, Austin, 1974. 185. E. Misaka and K. Nakanishi, J. Biochem. (Tokyo) 53, 465 (1963). 186. A. J. W. G. Viwer, H. J. Grande, F. Muller, and C . Veeger, Eur. J . Biochem. 45, 99 (1974). 187. 0. A. Moe, Jr., D. A. Lerner, and G. G. Hammes, Biochemistry 13, 2552 (1974).

3.

127

FLAVIN-CONTAINING DEHYDROGENASES

which the enzyme has accepted two electrons and these are shared between the FAD and the reactive disulfide. Furthermore, this intermediate, BH,, turns over once in each catalytic c,ycle accepting two electrons from dihydrolipoamide and donating them to NAD'. Figure 9 shows this cycle; I is the oxidized enzyme and I11 is EH,, while I1 and IV are complexes of EH, with the substrates. The enzyme catalyzes a readily reversible reaction, but the physiological direction is clockwise. Very simple representations such as this one do not adequately describe all that is known about the enzyme, and they force a choice of one form over others which may be equally good. For example, EH, is depicted as a charge transfer complex between thiolate anion and FAD ; but, as was discussed in Section II,B, this may be an inadequate description of the sharing of two electrons between the sulfur and FAD. On the other hand, such representations act as working hypotheses and suggest features of the mechanism to be tested. One such feature that has long eluded investigators is the possibility of intermediates in the very rapid reduction of the enzyme by NADH. It has been suggested that the FAD is reduced to FADH, followed by intramolecular electron rearrangement (98).It can be argued that without such a step the flavin does not play a true redox function when EH, is a charge transfer complex. If FADH, is an intermediate I

II

NADp f$ ~

7;

SH

IE III Fro. 9. Mechanism for lipoamide dehydrogenase.

128

CHARLES € WILLIAMS, I. JR.

in catalysis, then lipoamide dehydrogenase and glutathione reductase would share a common intermediate in catalysis with thioredoxin reductase (Sections IV,B and V,D) . The scheme in Fig. 9 suggests that lipoamide dehydrogenase functions by a simple binary complex mechanism, and this conforms to a vast body of kinetic evidence (Section, II1,D) , Since the oxidized enzyme can form a complex (or complexes) with NAD' and since EH, may form stable complexes with both NAD+ and NADH, a classic binary complex mechanism is too simple, Indeed, the substrate inhibition patterns are very complex and do not conform to any classic pattern. Until spectral properties and rates of formation and breakdown can be measured for each of these complexes the simple binary complex mechanism must serve. The reversibility of the reaction is also indicated by the scheme in Fig. 9. At pH 8.0, optimal for the reaction in the direction of NAD' reduction, the extrapolated maximal rate of NADH oxidation is about onethird that of NAD+ reduction, 7,500 and 20,000 min-' per enzyme-bound FAD, respectively. Much higher rates of NADH oxidation are observed a t lower pH provided NAD+ is present and the NADH concentration is low; a t pH 6.5 and at infinite lipoamide concentration, rates in excess of 80,000 min-l per FAD have been estimated. Many factors undoubtedly contribute to the difference in pH optimum for the reaction in opposite directions. The oxidation-reduction potentials of both substrate couples change considerably between pH 8.0 and pH 6.5; a t the latter pH, NADH is a much stronger reductant relative to dihydrolipoamide than is the case a t pH 8.0. The oxidation-reduction potentials of the enzyme must also change, and the combined changes result in the extreme sensitivity to NADH and the protection against this inhibition by NAD'. I n addition the predominant charge state of some groups on the enzyme would be shifted over this pH range; a group serving as an acid-base catalyst, so essential for thiol-disulfide interchange a t neutral pH (Fig. 81, would be effective in this role only a t pH values near its pK. Thus, both enzyme and substrates are quite different at the pH optima of the reaction in its respective directions. The means by which NAD' affects the oxidation of NADH is still uncertain. The evidence for two pyridine nucleotide binding sites is not compelling. The alternative explanation that NAD+ functions by reversing the equilibrium between EH, and 4-electron-reduced enzyme (EH,) is shown in Eq. (9). There is some kinetic evidence for a dead end' complex EH1+ NADH

EHrNADH

EHd-NAD+

EHI

+ NAD+

(9)

such as EH,-NADH. The kinetic evidence neither requires nor suggests two sites. Binding studies suggesting two sites have been carried out with

3.

FLAVIN-CONTAINING DEHYDROGENASES

129

the oxidized enzyme, whereas it is binding to EH, that is crucial t o the problem. The chemical modification, resulting in the abolition of the ability to form the charge transfer complex between FADH, and NAD‘, while not completely eliminating the ability of NADH t o reduce the enzyme, may be explicable in terms of very specific and subtle changes a t a single pyridine nucleotide site. Finally, it is of interest to consider the structural consequences of two sites for pyridine nucleotide and one for FAD on a monomer of 50,000 molecular weight. Between 90 and 140 amino acid residues are required to build up such a site (188). This would require a large number of the 475 residues in the molecule. This alone should make lipoamide dehydrogenase a prime candidate for crystallography and sequencing.

IV. Glutathione Reductase

Oxidized glutathione will be referred to as GSSG, and reduced glutathione will be referred to as GSH. Many of the properties of glutathione reductase have been discussed in Section 11. The ubiquity of glutathione reductase activity has been reviewed and some newer data given (39). The enzyme from yeast and from human erythrocytes has been most extensively studied. It has also been purified from r a t liver (189, IN), germinated peas (191), E . coli (5, 192), Penicillium chrysogenum (193), and sea urchin eggs (194). I n all cases the enzyme has been shown to be a flavoprotein.

FUNCTIONS A. METABOLIC Glutathione reductase catalyzes the virtually irreversible reduction of GSSG by NADPH. Its metabolic function is therefore synonymous with that of the product GSH. Glutathione is the most abundant thiol-disulfide pair in the cell by more than an order of magnitude; under most conditions the GSH:GSSG ratio is about 20:l (195). Since the ratio of 188. M. G. R o m a n n , D. Moras, and K. W. Olsen, Nature (London) 250, 194 (1974). 189. C. E. Mize and R. G. Langdon, JBC 237, 1589 (1962). 190. J. A. Ruzard and F. Kopko, JBC 238, 464 (1963). 191. L. W. Mapson and F. A. Isherwood, BJ 86,173 (1963). 192. C . H. Williams, Jr. and L. D. Arscott, “Methods in Enzymology,” Vol. 17, 503, 1971. 193. T. S. Woodin and I. H. Segel, BBA 167,64 and 78 (1968). 194. I. Ii and H. Sakai, BBA 350, 141 and 151 (1974). 195. P. C. Jocelyn, ed., “Biochemistry of the SH Group,” p. 10. Academic Press, New York, 1972.

130

CHARLES

H.

WILLIAMS, JR.

GSH :GSSG a t equilibrium of the reductase reaction is very high (calculated from data in 191, 196), it can be concluded that the reduction of GSSG is limited either by the enzyme level or by the availability of NADPH. GSH is utilized in two types of reactions: First, it is the substrate for enzymes such as GSH peroxidase (EC 1.11.1.9) and for a group of transhydrogenases, GSH-homocystine oxidoreductase (EC 1.8.4.1), GSHprotein disulfide oxidoreductase (EC 1.8.4.2), GSH-CoASSG oxidoreductase (EC 1.8.4.3), and GSH-cystine oxidoreductase (EC 1.8.4.4). The most thoroughly studied of these transhydrogenases is the GSH-protein disulfide oxidoreductase, better known as glutathione-insulin transhydrogenase (197,198).It catalyzes the first step in the sequential degradation of insulin (199). One of the products of the reaction, the phenylalanyl chain, is an inhibitor of glutathione reductase (200). It is not known if this constitutes a physiologically important feedback mechanism. The transhydrogenase contains either a reactive disulfide or a masked thiol since it is sensitive to thiol inhibitors only in the presence of its substrate, GSH (201). A nonspecific, GSH-protein disulfide transhydrogenase has also been purified (202). Second, GSH functions, presumably nonenzymically, in the reduction of protein thiols which have become oxidized to mixed disulfides (%XI). I n this latter function GSH in some cases converts inactive enzymes to active ones, or vice versa, and may thus serve as a means of metabolic control. Examples of this important possibility are glycogen synthetase D (EC 2.4.1.11) and fructose-1,6-diphosphatase(EC 3.1.3.11). The D form of glycogen synthetase is dependent for activity upon the presence of glucose 6-phosphate. The enzyme is inactivated by GSSG and reactivated by GSH (204). Mixed disulfide formation between thiols of the enzyme and GSSG leads to a decrease in affinity of the enzyme for its activator (205). A recent symposium on glutathione has covered many emerging functions of this peptide (206). Several potentially important effects have 196. W. M. Clark, “Oxidation-Reduction Potentials of Organic Systems,” p. 486. Williams & Wilkins, Baltimore, Maryland, 1960. 197. H.H.Tomizawa and Y. D. Halsey, JBC 234,307 (1959). 198. H.H.Tomizawa, JBC 237,428 and 3393 (1962). 199. P.T.Varandani, BBA 320,249 (1973). 200. R. G.Langdon, JBC 235, PC15 (1960). 201. P.T.Varandani and H.Plumley, BBA 151,273 (1968). 202. F. Tietze, BBA 220, 449 (1970). 203. R. Nesbakken and L. Eldjarn, BJ 87,526 (1963). 204. M.J. Ernest and X. H. Kim, JBC 248,1550 (1973). 206. M. J. Ernest and K. H.Kim, JBC 249,5011 (1974). 206. L. Flohe, H.C. Benohr, H. Sies, H. D. Waller, and A. Wendel, eds., “Proceed-

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131

been inferred by observation of the consequences of perturbing the GSHGSSG poise with the oxidizing agent, diamide [ (CH,) ,NCON= NCON (CH,) ,I. These include nerve transmission (20?’), membrane integrity (208), and protein synthesis (209). The relative specificity of this reagent for GSH has been demonstrated (210). A possible connection between a number of disease states, chiefly hemolytic and glutathione reductase deficiency, has been proposed (211). The conditions are characterized by a low GSH :GSSG ratio or by an inability to restore normal GSH-GSSG poise following a challenge, e.g., by a drug. In almost all cases the GSH deficiency has been traceable either to a lack of NADPH secondary to low glucose-6-phosphate dehydrogenase (211-215) or to riboflavin deficiency (212, 216-222). I n the latter case, glutathione reductase activity of erythrocytes was restored by administration of riboflavin (217-220). The enzyme has been purified and shown to have a diminished FAD affinity ( 2 2 1 , 2 2 2 ~ ) . GSH acts catalytically in the rearrangement of incorrectly formed ings of the 16th Conference of the German Society of Biological Chemistry.’’ Thieme, Stuttgart, 1974. 207. E. M. Kosower and N. S. Kosower, in “Proceedings of the 16th Conference of the German Society of Biological Chemistry” (L. Flohe et al., eds.), pp. 287-302. Thieme, Stuttgart, 1974. 208. N. S. Kosower and E. M. Kosower, in “Proceedings of the 16th Conference of the German Society of Biological Chemistry” (L. Flohe et al., eds.), pp. 216227. Thieme, Stuttgart, 1974. 209. N. S. Kosower and E. M. Kosower, in “Proceedings of the 16th Conference of the German Society of Biological Chemistry’’ (L. Flohe et al., eds.), pp. 276-287. Thieme, Stuttgart, 1974. 210. E. M. Kosower, W. Correa, B. J. Kinon, and N. S. Kosower, BBA 264, 39 (1972). 211. H. C. Benohr and H. D. Waller, in “Proceedings of the 16th Conference of the German Society of Biological Chemistry’’ (L. Flohe et al., eds.), p. 184. Thieme, Stuttgart, 1974. 212. E. Beutler, in “Proceedings of the 16th Conference of the German Society of Biological Chemistry” (L. Flohe et at?, eds.), pp. 109-114. Thieme, Stuttgart, 1974. 213. E. Beutler and S. K. Srivastava, Noture (London) 226, 759 (1970). 214. E. Beutler, Pharmacol. Rev. 21, 73 (1969). 215. S. K. Srivastava and E. Beutler, RJ 114, 833 (1969). 216. D. Glatzle, F. Weber, and 0. Wiss, Ezperientia 24, 1122 (1968). 217. E. Beutler. Science 165, 613 (1969). 218. B. Mandula and E. Beutler, Blood 36,491 (1970). 219. E. Beutler, J . Clin. Invest. 48, 1957 (1969). 220. S. K. Srivastava and E. Beutler, Ezperientia 26,250 (1970). 221. G. E. J. Staal, P. W. Helleman, J. DeWael, and C. Veeger, BBA 185, 63, (1969). 222. N. V. Paniker, S. K. Srivastava, and E. Beutler, BBA 215, 456 (1970). 222a. D. J. Worthington and M. A. Rosemeyer, Eur. J . Biochem. 48, 167 (1974).

132

CHARLES H. WILLIAMS, JR.

TABLE IV POSSIBLEALTERNATIVE SUBSTRATES OF GLUTATHIONE REDUCTASE Rate relative to GSSG ( % ) a

Substrate

Ref.

Yeast GR

A G-SS-CO GS--SOsG-SS-Cy G-SS-Pantothineb GSS-~-93-Hemoglobin GS-SeS-G bis-N,N( 7-Glutamyl) cystine

10 0.04 0.2

0.1

ca. 100 21

886,887 887 8.97 887 688 889

830

Erythrocyte GR DL-Lipoate GCystine DkHomocystine DTNB Cgstamine D-Pantothine 0

3.1 0.1 0.1 0.4 0.2 0.5

39 39 39 39 39 39

As discussed in Section II,A, rates of less than 0.5% are of doubtful significance. Progressive inhibitor in the presence of NADPH (831).

disulfides during protein folding. This reaction is catalyzed by a microsoma1 enzyme, protein disulfide isomerase (EC 5.3.4.1) (2.23,224). GSH unmasks a thiol group in the enzyme; this thiol group catalyzes the isomerization (22%). The specificity of glutathione reductase toward its disulfide substrate was emphasized in Section II,A, since there is virtually no reactivity with the substrates of the other pyridine nucleotide-disulfide oxidoreductases. Other authors have emphasized the lack of specificity of this enzyme since it can catalyze the reduction of a variety of mixed disulfides provided that glutathione or y-glutamylcysteine comprises one-half (193, 212) ; Table IV summarizes these ($9, 226-231). It is important to distin223. S. Fuchs, F. DeLorenzo, and C. B. Anfinsen, JBC 242,398 (1967). 224. E. D. Corte and R. M. E. Parkhouse, BJ 136,697 (1973). 225. F. DeLorenzo, S. Fuchs, and C. €3. Anfinsen, Biochemistry 5, 3961 (1966). 226. R. N. Ondarza and J. Martinez, BBA 113,409 (1966). 227. B. Mannervik and 8.A. Eriksson, “Glutathione,” pp. 120-132. Thieme, Stuttgart, 1974. 228. S. K. Srivastava and E. Beutler, BJ 119,353 (1970). 229. H. E. Ganther, Biochemistry 10,8089 (1971). 230. J. E. Smith, BBA 242, 36 (1971). 231. B. Mannervik and G. Nise, ARB 134,90 (1969).

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133

guish between this activity and that of the transhydrogenases (EC 1.8.4 group) (232-836). In rat liver supernatant, it is clear that glutathione reductase catalyzes the NADHP-dependent reduction of G-S-S-CoA (837) ; however, there is some evidence for a distinct enzyme catalyzing this reaction in yeast (238, 239) though this has been questioned (8.40). B. PROPERTIES OF THE 8-ELECTRON-REDUCED ENZYME, EH, The catalytic center of glutathione reductase can be represented as in Fig. 10, Structure I. I n catalysis the enzyme accepts two electrons from NADPH and donates two electrons to GSSG. The catalytic intermediate, Structure 111, will, as before, be referred to as EH,. The mechanism is formally identical to that given in Fig. 9 for lipoamide dehydrogenase. The spectrum of oxidized glutathione reductase and of EH, are shown in Fig. 1. The evidence for the similarity between glutathione reductase and lipoamide dehydrogenase in mechanism and structure has been discussed in Sections II,B and I1,C. Early recognitions of these similarities should be cited (241-243) as well as the suggestions that the flavoprotein (now referred t o as thioredoxin reductase) involved in the NADPH-linked reduction of methionine sulfoxide and “active sulfate” was remarkably like lipoamide dehydrogenase and glutathione reductase (7, 841, 24s). The principal features, in addition to EH,, common to lipoamide dehydrogenase and glutathione reductase deserve emphasis : the formation of complexes between the oxidized enzymes and their respective oxidized pyridine nucleotides; the formation of complexes between EH, and both oxidized and reduced pyridine nucleotides ; the formation of charge transfer complexes between 4-electron-reduced enzymes and oxidized pyridine 232. S. H. Chang and D. R. Wilken, JBC 241,4251 (1966). 233. A. Eriksson and B. Mannervik, FEBS (Fed. Eur. Bwchem. Sac.) Lett. 7 , 26 (1970). 234. M. Winell and B. Mannervik, BBA 184,374 (1969). 235. S. A. Eriksson and B. Mannervik, BBA 212,518 (1970). 236. P. L. Wendell, BBA 159, 179 (1968). 237. R. N. Ondarza, E. Escamilla, J. Gutierrez, and G . De La Chica, BBA 341, 162 (1974). 238. R. N. Ondarea, R. Abney, and M. Lopez-Colome, BBA 191, 239 (1969). 239. R. N. Ondarza and R. Abney, FEBS (Fed. Eur. Biochem. Soc.) Lett. 7 , 227 (1970). 240. S. Eriksson, C. Guthenberg, and B. Mannervik, FEBS (Fed. Eur. Bwchem. Soc.) Lett. 39, 296 (1974). 241. S. Black and B. Hudson, BBRC 5, 135 (1961). 242. V. Massey and C. Veeger, Annu. Rev. Biochem. 32,579 (1963). 243. S. Black, Annu. Rev. Biochem. 32,399 (1963).

134

CHARLES H. WILLIAMS, JR.

I

v Ip

/1

NADP+

m

F I ~ 10. . Mechanism for glutathione reductase.

nucleotide; and the high degree of homology in the sequences of amino acid residues around the active site disulfide. In both enzymes, EH, is formed and reoxidized a t rates commensurate with its function as an intermediate in catalysis. The principal differences between the two enzymes should also be mentioned. Lipoamide dehydrogenase catalyzes a reaction which is freely reversible over the neutral pH range while the glutathione reductase reaction is essentially irreversible, except a t high pH together with efficient removal of the product (GSH). Lipoamide dehydrogenase functions within a three-enzyme complex, accepting electrons from an enzyme-bound intramolecular dithiol, while glutathione reductase accepts electrons from NADPH and is reoxidized by GSSG yielding two molecules of GSH. Each enzyme is quite specific for its pyridine nucleotide, both in catalysis and in the formation of various characteristic complexes. Complex formation between EH, and pyridine nucleotide (NAD') was first noted in lipoamide dehydrogenase (87,118). It was clear that the spectra of EH, were different when the reductant was NADH or when it was dihydrolipoamide; and in light of the NAD+ requirement in the oxidation of NADH, a complex of EH, with NAD+ was hypothesized. Thus, when reductant-dependent differences were observed in the spectra

3.

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FLAVIN-CONTAINING DEHYDROGENASES

of 2-electron-reduced glutathione reductase EH,, an EH,-NADP' complex was proposed (23). However, i t has subsequently been shown quite conclusively that the major complex is EH,-NADPH in glutathione reductase (52, 5 3 ) . In order to demonstrate EH,-NADPH or EH,-NADP+ complexes, it was necessary to produce the uncomplexed EH,. On the assumption that the very slow turnover of yeast glutathione reductase with NADH was a reflection of the poor binding of this pyridine nucleotide, the rates of reduction of the enzyme to EH, by various concentrations of NADH were measured. A second-order rate constant was determined as 2 X lo4 M-' sec-*, and there was no indication of a Michaelis complex ( 5 2 ) . Evidence was then obtained for both EH,-NADPH and EH,-NADP+ ( 6 3 ) . The dissociation constant of the EH,-NADPH complex was found to be 2 p M EH,-NADP+ proved to be unstable, and the product (formed too slowly to be of catalytic importance) was not identified ( 5 3 ) . These results with the yeast enzyme (53) have been confirmed with the E. coli enzyme where the dissociation constant for the EH,-NADPH complex was found to be 12 pM (60).The spectrum of EH,-NADPH is shown in Fig. 11. Compared to uncomplexed EH, I

I

I

I

I

I

I

400

I

I

I

I

I

500

600

700

Wavelength (nm)

FIQ.11. Complex formation between the 2-electron-reduced ( E K ) form of E. coli glutathione reductase and NADPH. EHz was produced by anaerobic reduction with borohydride; time wm allowed for the slight excew of borohydride to react with water before beginning the titration with NADPH. 1, Oxidized; 2, EHz; 3, EK+0.45 equivalent NADPH; 4, EH,+0.90 equivalent NADPH; and 5, EHz+ 2.75 equivalents NADPH.

136 12

CHARLES H. WILLIAMS, JR. 1

1

I

I

I

I

I

i

Wavelength (nm)

FIO.12. Complex formation between the 2electron-reduced form of E. coli glutathione reductase and NADP+.l, Oxidized ; 2, EH,; 3, EHn 1.72 equivalents NADP’ ; and 4, EH2 1036 equivalents NADP’. The procedure was as in Fig. 11.

+

+

it has higher extinction a t long wavelengths and lower extinction a t 450 nm. The spectrum of EH,-NADP’ is shown in Fig. 12; it is similar in all respects to the analogous spectrum of lipoamide dehydrogenase (118) in that, compared with the free EH,, it has lower extinction a t 540 nm and higher extinction a t 700 nm with the isosbestic point a t 590 nm. I n both cases EH, was produced with borohydride and the stable spectrum was recorded after the small excess of borohydride had reacted with water (60). EH, from yeast produced in this way is relatively stable in air (though these experiments were done anaerobically) and can be dialyzed aerobically overnight with only minor reoxidation (W44). Only the halfreduced NADPH-cytochrome P-450 reductase is comparably stable to oxygen (946, 946). The findings with glutathione reductase from yeast and E . coli are con244. V. Massey and C. H. Williams, Jr., in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 370. Springer-Verlag, Berlin and New York, 1970. 245. B. S. S. Masters, H. Kamin, Q. H. Gibson, and C. H. Williams, Jr., JBC 240, 921 (1965). 246. H. Kamin, B. S. S. Masters, Q. H. Gibson, and C. H. Williams, Jr., Fed. Proc., Fed. Amer. SOC.E x p . Biol. 24, 1164 (1965).

3.

137

FLAVIN-CONTAINING DEHYDROGENASES

sistent with the following equations based in large measure on the schemes previously proposed ( 5 3 ) . NADP+ E-NADP+-L-

NADPH

NADP' (E-NADPH)-EH,-NADP+-L--

E

EH,

(10)

NADPH

EH, EH,-NADP+

-

EH,-NADPH

EH,-NADP

(12)

(anion) NADP'

EH,-NADPH - E H , - N A D P + - ~ -

EH,

(13)

The reactions in Eq. (lo), proceeding to the right from E, are part of normal catalysis as shown in Fig. 10 (51, 6 3 ) . The association of E with NADP+ leads to a dead end complex. I n the reaction of yeast glutathione reductase with NADPH, EH,-NADPH appears to be formed in the dead time of the rapid reaction spectrophotometer (ca. 3 msec) when observation is a t 540 nm (244) ; however, if 3.4 pM EH,(free) is mixed with 20 pkf NADPH, Eq. (11), a minimum rate of complex formation of 10,000 min-' can be estimated when observation is a t 590 nm (by assuming a maximum half-time of 4 msec) ( 6 3 ) .Thus, EH,-NADPH is formed a t a fixed concentration of NADPH (in the absence of GSSG, 2 5 O ) a t a rate comparable with the overall maximal rate of catalysis, i.e., 15,000 min-' (89). Rates of reoxidation of EH, and EH,-NADPH by GSSG have not been compared. Equation (12) represents the slow conversion of EH,-NADP' to another species (53) which has been very tentatively identified as the anion semiquinone-NADP' complex (60) on the basis of its spectral properties which are shown in Fig. 13. Its formation has never been observed to go to completion but it is favored by high NADP' concentrations, high pH, and is inhibited by excess NADPH (53, 60). An E P R signal forms as the conversion proceeds; the free radical concentration was estimated to be 50% of the FAD concentration (60). The extinction maxima a t 360, 410, and 480 nm are characteristic of the semiquinone anion ( l ) ,but the identification is made difficult by the presence of large amounts of EH,-NADP+. Slow changes associated with the production of an E P R signal (10% of the FAD) are observed with lipoamide dehydrogenase in the presence of excess NAD', but the spectral changes are insufficient to be interpreted (59).Equation (13) represents the fourelectron reduction of the enzyme and does not proceed unless NADP' is removed from the equilibrium as with NADase. The rates of the combined reactions [Eq. (13)] are a t least 10-fold slower in yeast glutathione reductase than they are in lipoamide dehydrogenase in parallel experi-

138

CHARLES H. WILLIAMS, JR.

Wavelength Inm)

FIG.13. Yeast glutathione reductase, semiquinone anion production from the 2electron-reduced form. Curve 1, oxidized enzyme, anaerobic conditions, pH 7.6 ; curve 2, 1 min after the addition of 1 equivalent of NADPH; curve 3, 22 hr later; curve 4, 1 hr after the addition of 10 equivalents of NADP'; curve 5, 23.5 hr later; curve 6, 18.5 hr after the addition of 5 equivalents of NADPH; and curve 7, 35 min after opening to air.

ments with identical concentrations of NADase ( d 9 ) . Indeed, since EH2 forms a stable complex with NADPH in glutathione reductase, the EH,/EH, couple must have an oxidation-reduction potential considerably more negative than that of the NADPH-NADP+ couple.

C. KINETICSTUDIES Equations (10) and (11) indicate three intermediates, EH,, EH,NADP', and EH,-NADPH, which are formed a t rates sufficient to require their consideration as reactants with GSSG. Figure 10 hypothesizes a simple binary complex mechanism based on the early kinetic studies with enzyme from erythrocytes (39, do), peas (191), yeast ( d 9 ) , and P. chrysogenum (193). If this hypothesis is correct then NADP+ dissociates from EH2 prior to reaction with GSSG and only EH2 and EH,-

3. FLAVIN-CONTAINING

DEHYDROGENASES

139

NADPH need be considered as possible reactants with GSSG. As was mentioned, the rate of reoxidation of EH, by GSSG has been measured (944), but it has not been compared with the rate of reoxidation of EH,-NADPH. However, the tight binding of EH, to NADPH and the rapid formation of this complex (see above) would seem to indicate that it is a significant intermediate in catalysis; if it were not, NADPH should be a more potent substrate inhibitor. Thus, the small deviations sometimes observed from the kinetics expected from a simple binary complex mechanism might have their origin in different rates of reaction of GSSG with forms such as EH, and EH,-NADPH. The early kinetic studies on glutathione reductase did not include investigation of product inhibition, so vital to a proper interpretation of kinetic data in the elucidation of the mechanism (947, 948). In the one case where product inhibition patterns were observed, they were not interpreted by more recent kinetic theory (40). Subsequent kinetic analyses (see below) , in which product inhibition patterns have been obtained, were either completed prior to the discovery of the EH,-NADPH complex (53) or have not considered it. Furthermore, the product inhibition patterns have been carried out a t only a single level of the fixed substrate; it is essential that the patterns be obtained a t more than one level of fixed substrate, especially where dead end complexes are involved (949) as has been so amply demonstrated with lipoamide dehydrogenase (95, 167). In spite of these deficiencies, the more recent kinetic studies have yielded much useful information. Reexamination of the kinetics of erythrocyte glutathione reductase using a preparation of higher specific activity (51) confirmed the substrate and product inhibition patterns and the salt effects of the earlier study (40). Product inhibition by NADP+ was found to be competitive with NADPH and noncompetitive toward GSSG. Since this was in conflict with the predicted inhibition patterns for a binary complex mechanism, the authors proposed that the mechanism was mixed, ordered sequential and binary complex (51); these results have been confirmed (950).Furthermore, these studies (@, 51) showed that there was substrate inhibition: a t high levels of GSSG; a t high levels of NADPH if the GSSG concentration was low; and that the inhibition was more pronounced a t low ionic strength. Both the K , for GSSG and the V,,, were affected by ionic strength: At 0.3 M phosphate, V,,, was 14,300 moles NADPH per min 247. K. Dalziel, BJ 84, 244 (1962). 248. W. W. Cleland, BBA 67, 104 (1969). 249. K. Dalziel, in “Pyridine Nucleoticl,e-Dependent Dehydrogenases” (H. Sund, ed.), p. 373. Springer-Verlag, Berlin and New York, 1970. 250. B. Mannervik, Acta Chem. Scand:W,2912 (1969).

140

CHARLES H. WILLIAMS, JR.

a,

per FAD, K,(NADPH) was 13 and K,(GSSG) was 125 p M ; at 0.03 M phosphate, V,,, was 7,700, K,(NADPH) was 9.5 VJM, and K,(GSSG) was 19 f l . Thus, high ionic strength stimulated only a t GSSG concentrations higher than 100 pM (51).The kinetics with NADH (40, 61) and with lipoate (40) were also investigated; they demonstrated the less marked specificity of the erythrocyte enzyme as compared to the highly specific yeast enzyme. Effects of a wide variety of cations and anions have been reported ($9).The pH optimum is broad and centered a t 6.8 (.!40). I$, The binary complex mechanism can be formalized as follows: kr

ki

E

+ NADPH + E-NADPH S EHp + NADP+ kz k4

kh

EHe

ki

+ GSSG Ske EHrGSSG ke E + 2 GSH

+

and V,,, is then k, * k,/(k3 k7). The rate of formation of EH,, k,, is too rapid to be measured accurately at 25”, but it can be measured a t 5” and is found to be 5250 min-I. If k, could be measured, then V,,, could be calculated and compared with that measured by steady-state kinetics which a t 5O is 1960 min-I; correlation between the calculated and measured rate would be evidence for the binary complex hypothesis. The unusual stability of the 2-electron-reduced form, EH,, of yeast glutathione reductase in the presence of oxygen makes possible the preparation of EH, free of any reductant by reduction with borohydride and dialysis. The rate of reoxidation of EH, by GSSG is found to be 2900 min-I. The calculated V,,, is then 1880 min-I, in excellent agreement with that found by steady-state kinetics (S44, 251). These data demonstrate that the presence of NADP’ or NADPH does not alter the rate of reoxidation of EH, by GSSG. Neither do these data preclude the reoxidation of EH,-NADP’ or EH,-NADPH complexes a t rates similar to the rate of reoxidation of free EH,. The yeast enzyme, like the erythrocyte enzyme, is inhibited by NADP’ and the inhibition is competitive with NADPH (&EL-254).The inhibition is rather weak, the inhibition constant being a t least 10-fold higher than the K,,,for NADPH. On the basis of these studies a mixed mechanism 251. C. H. Williams, Jr. and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 289. Univ. Park Press, Baltimore, Maryland, 1971. 252. B. Mannervik, in “Structure and Function of Oxidation Reductase Enzymes (A. Akeson and A. Ehrenberg, eds.), p. 425. Pergamon, Oxford, 1972. 253. B. Mannervik, BBRC 53, 1151 (1973). 254. €3. Mannervik, in “Glutathione” (L. Flohe et al., eds.), p. 114. Thieme, Stuttgart, 1974.

3.

-

FLAVI N CON TAI N I NG DEHY DROGEN ASES

141

has been proposed. It is claimed that the sequential mechanism applies at high concentrations of GSSG and that the binary complex mechanism applies a t low GSSG concentrations (253, 254). In this connection it should be recalled that the physiological ratio, GSH:GSSG is always high. The kinetics of the reverse reaction have been investigated and have been taken as evidence for a sequential mechanism (255).The p H optimum for the reverse reaction is about 8 (255) while that of the forward reaction is about 7 (29).These studies have not considered the EH2NADPH complex or the possibility that this complex as well as the EH2NADP' complex could be reoxidized by GSSG. Since NADP' is not as tightly bound to the oxidized enzyme, its rapid dissociation can be inferred. A reasonable proposal has been made which potentially reconciles all of the present data ( 5 3 ) . A hybrid binary complex mechanism has been proposed for transcarboxylase from Propionibacterium shermanni in which distinct sites exist for the two substrate pairs, analogous to NADPH-NADP+ and GSH-GSSG with glutathione reductase (256). From what is now known of pyridine nucleotide binding sites (188) and of the disulfide loop of glutathione reductase (see Section I1,C) , independent sites are an attractive hypothesis. Kinetic constants for the yeast enzyme are remarkably like those for the erythrocyte enzyme: V,, = 15,000 moles NADPH per min per mole of FAD, K,(NADPH) = 3.8 p M and, K,(GSSG) = 55 pLM (89). Specific anion effects on the yeast enzyme have been interpreted to indicate two anion binding sites near the active site (257').

D. THIOL GROUPS Glutathione reductase from yeast and from E . coli contains 4 and 5 thiol groups per FAD, respectively (Table I), in addition to the active center disulfide. Peptides containing 3 of the 4 thiols have been isolated and sequenced : Cys-Asn Asp ; Lys-Ile-Ala-Cys-Pro-Gly-Asn-Val-GlnLys ; Asp-Thr-Ile-Tyr- (His,Glx) -Val-Cys-Lys- (Thr,Gly,Ala,Leu,) ( 3 5 ) . The thiols in the yeast enzyme, like those in lipoamide dehydrogenase, are relatively unreactive. Reactivity with phenyl mercuric acetate (29) and with N-ethylmaleimide (258) have been reported; it is of interest that the 2-electron-reduced enzyme is stable in the presence of N-ethylmaleimide and excess NADPH (258), but four-electron reduction results in the presence of p-chloromercuriphenyl sulfonate ( 2 9 ) . 255. 256. 257. 258.

A. L. Icen, FEBS (Fed. Eur. Biochem.Soc.) Lett. 16,29 (1971). D. B. Northrop, JBC 244, 5808 (1969). G. Moroff and K. G. Brandt, ABB 159,468 (1973). R. F. Colman and S. Black, JBC 240, 1796 (1965).

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Erythrocyte glutathione reductase is not sensitive to cupric ion (969). An early study of the enzyme from rat liver showed that GSSG was bound to the enzyme (1 GS-/44,000 g of enzyme) and that GSH was released upon the addition of NADPH (960).This finding, made prior to the knowledge that the enzyme was a flavoprotein, implies that a thiol on the protein reacts with GSSG yielding 1 GSH and that the half of the GSSG bound to the protein in mixed disulfide linkage was released by reduction of that linkage by NADPH. A possible explanation of these data is that the preparation was a mixture of glutathione reductase and a glutathione transhydrogenase ; however, the very high specific activity of the preparation strongly indicates the need for the reinvestigation of this finding. V. Thioredoxin Reductare

Some of the catalytic and structural properties of thioredoxin reductase as they relate to analogous properties of lipoamide dehydrogenase and glutathione reductase have been covered in Section 11. The flavoprotein, thioredoxin reductase, catalyzes the electron transfer from NADPH to thioredoxin, a protein of 12,000 molecular weight containing a single disulfide. The reductase has a reactive disulfide in addition to FAD. Thus, electron flow is from NADPH to the FAD-disulfide system of thioredoxin reductase, to the disulfide of thioredoxin, and finally to a variety of acceptor systems. A. METABOLIC FUNCTIONS The metabolic function of the thioredoxin reductase-thioredoxin system is to supply reducing equivalents to a wide variety of acceptors. B y far the best characterized of these is the E . coli ribonucleotide reductase system (93, 961); the reductase consists of two subunits, proteins B1 and B2 (269, 963). The B1 protein contains three reactive dithiol-disulfide pairs and appears to be the immediate acceptor of reducing equivalents from thioredoxin. As isolated, the three pairs are in the reduced state and, in the presence of the B2 protein, three molecules of ribonucleotide can be reduced prior to any input of reducing equivalents from thiore259. N. S. Agar and J. E. Smith, Proc. Soc. Exp. Biol.' Med. 142, 562 U973). 260. C. E. Miae, T. E. Thompson, and R. G. Langdon, JBC 237, 1596 (1962). 261. P. Reichard, Eur. J . Bbchem. 3, 259 (1968). 262. N. C. Brown, Z. N. Canellakis, P. Reichard, and L. Thelander, Eur. J . Biochem. €I 561 , (1969). 263. L. Thelander, JBC 248, 4591 (1973).

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TABLE V SYSTEMS FOR WHICHTHIOREDOXIN Is THE POSSIBLE ELECTRON DONOR Ribonucleotide reductase, Escherichia coli (83,861-866) Yeast (46, 267) Lactobacillus leichmannii (868-870) Mammalian liver supernatant (808) Rat hepatoma (871) T4-induced E. coli (47, 878) Euglena gracilis (873) Nonspecific protein disulfide reductase, pea seeds (874) Methionine sulfoxide reductase, yeast (8, 876, 876) “Active” sulfate reductase, yeast (7, 876-877) &Proline reductase, Clostridium sticklandii (878, 979)

doxin (264).The B2 protein is an iron-sulfur protein (265). The ribonucleotide reductase system of Lactobacillus leichmannii, in contrast to the E. coli enzyme, is vitamin B,,-dependent (266). Table V (6, 7 , 23, 46, 47, 902, 261-265, 267-279) indicates the wide variety of systems for which thioredoxin is the possible electron donor. While the term “thioredoxin” was coined in connection with the ribonucleotide reductase system (23) this protein had been demonstrated in two systems previously (6, 7 ) . Thioredoxin reductase (EC 1.6.4.5) has been purified from E . coli (8, 3 6 ) , yeast (6, 7, 66, 275), and rat liver supernatant (202, 280). Only the enzyme from E . coli has been extensively characterized, and it will be discussed exclusively below. The similarity of the transhydrogenase from Azotobacter vinelandii to thioredoxin reductase has been noted )

(281) .

264. L. Thelander, JBC 249, 4858 (1974). 265. C. L. Atkin, L. Thelander, P. Reichard, and G. Land, JBC 248, 7464 (1973). 266. R. L. Blakley and H. A. Barker, BBRC 16,391 (1964). 267. E. Vitols, V. A. Bauer, and E. C . Stanhrough, BBRC 41, 71 (1970). 268. E. Vitols and R. L. Blakley, BBRC 21,466 (1965). 269. M. D. Orr and E. Vitols, BBRC 25, 109 (1966). 270. W. S. Beck, M. Goulian, A. Larsson, and P. Reichard, JBC 241, 2177 (1966). 271. E. C. Moore and P. Reichard, JBC 239,3453 (1964). 272. 0. Berglund, JBC 244, 6306 (1969). 273. F. K. Gleason and H. P. C. Hogenkamp, JBC 245,4891 (1970). 274. M. D. Hatch and J. F. Turner, BJ 76,556 (1960). 275. P. G. Porque, A. Baldesten, and P. Reichard, JBC 245, 2363 (1970). 276. P. G. Porque, A. Baldesten, and P. Reichard, JBC 245, 2371 (1970). 277. L. G. Wilson, T. Asahi, and R. S. Bandurski, JBC 236, 1822 (1961). 278. T. C. Stadtman and P. Elliott, JBC 228, 983 (1957). 279. D. S. Hodgins and R. H. Abeles, ABB 130,274 (1969). 280. A. Lawon, Eur. J . Biochem. 35, 346 (1973). 281. H. W. J. van den Broek and C. Veeger, Eur. J . Biochem. 24, 63 (197111

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CHARLES H. WILLIAMS, J R .

B. SPECIFICITY OF THIOREDOXIN REDUCTASE Thioredoxin reductase is specific for NADPH and moreover for the hydrogen from the B side of the nicotinamide ring (282). The specificity of the flavoprotein for various disulfides was discussed in Section I1,A. The specificity of the E . coli reductase for thioredoxin from several sources has been tested; thioredoxin from yeast is not reduced by NADPH in the presence of the E . coli reductase (275).The sequence of amino acids around the redox-active disulfide of yeast and E. coli thioredoxin shows a high degree of homology (46). Thus, it would seem unlikely that major determinants of substrate recognition lie in this region as had been suggested (62).When E. coli are infected by T 4 phage, a new thioredoxin is made simultaneously with the normal thioredoxin. Both thioredoxins are reduced by NADPH plus thioredoxin reductase ; K , values (apparent, measured a t 120 pM NADPH) for the normal and T4-induced thioredoxins are 10 and 6 p M , respectively. The sequence of amino acid residues around the reactive disulfide in the T4-induced thioredoxin is totally different from that of the normal thioredoxin; even the 2 residues between the half-cystines are different: -Cys-Val-TyrCys- (47). It is possible that the determinants of substrate recognition by the reductase will emerge from X-ray crystallography studies on the thioredoxins now in progress (47).

c. GENERALPROPERTIES OF THE E.C O l i ENZYME The reduction of thioredoxin by NADPH is virtually complete, an equilibrium constant of 48 having been estimated a t pH 7 (8).The equilibrium constant [T(SH)2][NADP+]/[T(S)2][NADPH], falls by a factor of 10 for each unit rise in pH (8).The activity is stimulated about 2-fold in phosphate buffer as compared to Tris and the pH optimum is about 7.7 (8).The molecular weight determined by sedimentation equilibrium is 66,000 (38) and by amino acid analysis is 73,OOO-75,000 based on 2 moles of FAD per mole of enzyme (68).The partial specific volume is 0.724 ml/g (8). The spectrum of the oxidized enzyme is given in Figs. 3 and 4. The extinction coefficient of the FAD a t 456 nm is 11.3mM-' cm-l. Spectral ratios are A(271 nm) :A(456 nm) = 5.8; A(380 nm) :A(456 nm) = 1.03 ( 3 6 ) . Kinetic parameters are given in Table VI. Plots of l / v against 1/(S) give parallel lines both a t 4O and 25O; on this basis the assumption has been made that a binary complex mechanism is operative. As with lipoamide dehydrogenase and glutathione reductase, this assumption is com282. A. Larsson and L. Thelander, JBC 240,2691 (1966).

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DEHYDROGENASES

TABLE VI THIOREDOXIN REDUCTASE-KINETIC CONSTANTS" 4

L(NADPH)(rM) K,(thioredoxin) ( p M ) Ki(NADP/) (&') Turnover number/FAD (min-I) a

0.8 1.7 520

25OC 1.2 2.8 15

2 ,000

Assays were carried out in 0.05 M phosphate buffer, pH 7.6, 1.5 mM EDTA.

* At 4", the disappearance of NADPH was followed at 340 nm in 5 cm pathlength cells. At 25", the assays contained 0.2 mM DTNB and the appearance of TNB anion was followed at 412 nm; the assays also contained 10 mM glucose &phosphate and glucose-&phosphate dehydrogenase to overcome inhibition by NADP/ except in the experiment estimating NADP+ inhibition.

plicated by the product inhibition pattern in which inhibition by NADP' is competitive with NADPH. The enthalpy of activation calculated from the turnover numbers a t the two temperatures is 10,000 cal/mole FAD (689).

D. REDUCED STATESOF THE ENZYME-MECHANISM The titration of thioredoxin reductase with NADPH is shown in Fig. 4. Four electrons per FAD seem to be required for reduction; this is con-

firmed in titrations with dithionite or NADPH in the presence of NADase (to hydrolyze the product NADP'). It can be seen that the first increments of NADPH cause a relatively larger change than do later increments. It seems likely that this is indicative of an unequal sharing of electrons between the FAD and the oxidation-reduction-active disulfide which favors the FAD. The long wavelength absorption extending to 900 nm has been ascribed to charge transfer interaction between FADH, and NADP'. It is absent when the titration is carried out in the presence of NADase or in reduction by dithionite. I n these latter titrations, a distinct long wavelength absorption is observed which does not extend beyond 700 nm (see Section V,E) (SO). These results have been confirmed, and in addition it has been shown that the enzyme can be partially reduced by excess reduced thioredoxin (31). Early studies indicated that the enzyme contained a second electron acceptor and that this acceptor was a disulfide (Section I1,B) (8, 30). This conclusion is greatly strengthened by the observation that enzyme 283. S. Ronchi, G. Zanetti, and C. H. Williams, Jr., unpublished results.

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CHARLES H . WILLIAMS, JB.

FIQ.14. Mechanism for thioredoxin reductase.

reduced by 1.5 equivalents of NADPH reoxidized partially upon the addition of an excess of p-chloromercuriphenyl sulfonate and that in a similar experiment using only 1.0 mole of NADPH reoxidation was complete. Thus electrons can be trapped on the nascent thiols by mercurial. The enzyme-bound FAD following such experiments is fully reduced by 1.0 mole of NADPH (30). The reduction of thioredoxin reductase by NADPH observed in the rapid reaction spectrophotometer a t 2O proceeds in three phases. The first phase is complete in the dead time of the instrument (ca. 3 msec) and results in a spectrum with enhanced absorption a t long wavelengths and very little change in absorption in the 450-nm region. This species decays to the 4-electron-reduced enzyme in two phases, the first having a rate constant of 2600 min-I and the second of 300 min-l. From the rate of the overall reaction a t low temperature (Table VI), it is concluded that the first and second phases are catalytically significant, while the third represents a side reaction ($84). The scheme shown in Fig. 14 represents a working hypothesis for the reaction mechanism of thioredoxin reductase. This constitutes a gross oversimplification. Thioredoxin reductase may be more complex than are 284. V. Massey, R. G . Matthews, G. P. Foust, L. G . Howell, C. H. Williams, Jr., G. Zanetti, and S. Ronchi, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 393. Springer-Verlag,Berlin and New York, 1970.

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lipoamide dehydrogenase and glutathione reductase, where only one spectrally distinct species, EH,, and its complexes with oxidized and reduced pyridine nucleotides is observed. I n thioredoxin reductase on the other hand, intramolecular electron transfer between the FAD and the disulfide does not involve a spectrally identifiable interaction between the two redox groups. I n the scheme, catalysis proceeds in a clockwise direction in the upper square when thioredoxin is in excess and I represents the oxidized enzyme; I1 is considered the rapidly formed species. The spectrum of the species formed with a rate constant of 2600 min-' has not been determined, but it is assumed to be 111, or more properly a mixture of I11 and VI. If NADPH is in excess, catalysis might be pictured as proceeding from I to I1 to I11 to IV to V to VI back to I11 and then counterclockwise around the lower square. The slow reaction with rate constant of 300 min-' may represent the dissociation of NADP' from VI. Distinction of the subtle differences between this scheme and that previously proposed (284) will require an extensive rapid reaction investigation. The electron distribution between FAD and the disulfide observed to favor the flavin indicates that the species IV will have a very transient existence.

E. LIGHT-ACTIVATED REDUCTION-NEUTRAL SEMIQUINONE The formation of a blue (neutral) semiquinone in high yield upon irradiation of thioredoxin reductase in the presence of a large excess of EDTA is shown in Fig. 3a. The semiquinone is further reduced to FADH, a t an even slower rate with maximal semiquinone formation a t 4 hr. I n contrast to this very slow semiquinone production, enzyme reduced by NADPH in the dark and subsequently exposed to light is rapidly converted to the semiquinone. The rate depends on the amount of NADPH used in the reduction; with 0.5 mole NADPH per FAD the half-time is less than 0.5 min, with 2.0 moles NADPH per FAD the half-time is about 2 min. The rate of free radical production (EPR) exactly parallels the rate of increase in absorbance a t 580 nm. The exact spectral characteristics of the semiquinone depend on the state of oxidation of the disulfidedithiol. In the dithiol form the maximum is a t 578 nm while in the disulfide form the maximum is at 588 nm. That the spectral properties are determined by the redox state of the disulfide is indicated by three findings. If semiquinone is produced by irradiation following reduction of the enzyme by 0.5 mole/FAD, the maximum is a t 588 nm, while if the semiquinone is formed following reduction by 2.0 moles/FAD, the maximum is a t 578 nm. Oxidation of enzyme irradiated in the presence of excess EDTA for various lengths of time requires ferricyanide stoichiometric with the ob-

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CHARLES H. WILLIAMS, J R .

served semiquinone for irradiation times less than 4 hr; but for longer times the ferricyanide required is greater up to a maximum of 4 equivalents. The semiquinone is not reduced by NADPH or reoxidized by thioredoxin; however, semiquinone exhibiting a maximum a t 578 nm, upon addition of thioredoxin, shifts its maximum to 588 nm and NADPH causes the opposite shift (58). The lack of reactivity of the semiquinone per se with either thioredoxin or NADPH shows that it cannot be involved in catalysis. The rapid production of semiquinone by irradiation of partially reduced enzyme is a light-activated disproportionation since it is totally dependent upon the presence of some oxidized enzyme. Enzyme fully reduced by dithionite forms no semiquinone, while enzyme partially reduced by dithionite rapidly forms semiquinone upon irradiation. Furthermore, the light-activated disproportionation of enzyme first reduced with NADPH results in the reduction of NADP'. Thus, FAD catalyzes the disproportionation in keeping with the known photosensitizing nature of free flavins. This reaction is reversed slowly (half-time a.150 min 25') in the dark. The semiquinone is rapidly reoxidized by oxygen to yield an enzyme with unaltered spectral and catalytic properties (58).Similar reactions have been very briefly reported for lipoamide dehydrogenase ; the dark reverse reaction is comparatively rapid, being complete in 30 min (163).

VI. Microsomal Electron Transport

It is generally accepted that liver microsomes contain two electron transport systems each containing a flavoprotein reductase. The paths of electron flow can be outlined as follows: NADPH -+NADPH-cytochrome reductase NADH-NADH-cytochrome reductase

P-450 lipid- cytochrome P-450 -0,

b , lipid_cytochrorne b , -"CN-sensitive

factor"

t 0 2

The "CN-sensitive factor" is the only component which has not been characterized as a molecular entity (284a). I n Section VI,A, the makeup 284a. P. Strittmatter, L. Spats, D. Corcoran, M. J. Rogers, B. Setlow, and R. Redline, Proc. Nat. Acad. Sci. U. S. 71, 4566 (1974) ; added in proof: The desaturaae is a single polypeptide chain of 63,000 daltona containing 62% nonpolar amino acids and one atom of nonheme iron.

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and metabolic functions of the NADPH-dependent system will be outlined; in Section VI,B, the NADH-dependent system will be outlined; and in Section VI,C, the possibilities for interchain electron transfer will be briefly surveyed. A. THENADPH-CYTOCHROME P-450 REDUCTASE-CONTAINING SYSTEM The mixed function oxidations (reactions requiring both oxygen and reducing equivalents) of endogenous substrates such as steroids and fatty acids, and of drugs, carcinogens, and other foreign compounds, are carried out by the cytochrome P-450 system. Most of these reactions are either hydroxylations or oxidative N-dealkylations in which half the oxygen molecule is incorporated and half is reduced to water. The system is always present, but it can be induced to much higher levels by the chronic administration of a potential substrate. There is some evidence that distinct systems are induced by different substrates containing spectrally distinct cytochromes, cytochrome P-450 and cytochrome P-448. A very good bibliography indicating documentation for these introductory comments is contained in Lu et al. ( 2 8 5 ) . This system has been solubiliaed from the liver microsomal membrane, resolved into three components and reconstituted (285-29291). All three components, flavoprotein, cytochrome P-450, and lipid, are required for maximal substrate metabolism. The active component in the lipid fraction has been found to be phosphatidylcholine (292). The system from kidney microsomes has been partially resolved and reconstituted (293, 294). The system from yeast has been resolved into three similar fractions and reconstituted; the yeast reductase and lipid fractions can be replaced by analogous fractions from liver (295). Thus, the system is perhaps ubiquitous in eukaryotic microsomes. As was mentioned in Section I, the systems carrying out similar reactions in prokaryotes and adrenal mito285. A. Y. H. Lu, R. Kuntzman, S. West, M. Jacobson, and A. H. Conney, JBC 247, 1727 (1972). 286. A. Y. H. Lu, K. W. Junk, and M. J . Coon, JBC 244, 3714 (1969). 287. A. Y. H. Lu, H. W. Strobel, and M. J. Coon, BBRC 36, 545 (1969). 288. A. Y. H. Lu, H. W. Strobel, and M. J. Coon, J . M o l . Pharm. 6, 213 (1970). 289. K. Ichihara, E. Kusunose, and M. Kusunose, Eur. J . Bwchem. 38,463 (1973). 290. W. Levin, D. Ryan, S. West, and A . Y. H. Lu, JBC 249, 1747 (1974). 291. A. Y. H. Lu and W. Levin, BBA 334,205 (1974). 292. H. W. Strobel, A. Y. H. Lu, J. Heidema, and M. J. Coon, JBC 245, 4851 (1970). 293. K. Ichihara, E. Kusunose, and M. Kusunose, BBA 239, 178 (19’711.. 294. K. Ichihara, E. Kusunose, and M. Kusunose, FEBS (Fed. Eur. Biochem. SOC.)Lett. 20, 105 (1972). 295. W. Duppel, J. M. Lebeault, and M. J. Coon, Eur. J . Biochem. 36,583 (1973).

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CHARLES H. WILLIAMS, JR.

chondria contain an iron-sulfur protein and seem to have no lipid requirement. The yeast system shows optimal activity with lysophosphatidylethanolamine (896). Although resolution and reconstitution represent the strongest evidence for the makeup of the NADPH-cytochrome P-450system, other techniques have contributed important indirect evidence. Immunochemical techniques have been used to indicate functionality of individual components, particularly the flavoprotein reductase ; antibodies have been prepared against the purified reductase (296, 897) and shown to inhibit such processes as aniline hydroxylation (296),o-hydroxylation of fatty acids (898), ethylmorphine demethylase (8973, NADPH-cytochrome c reductase and NADPH-cytochrome P-450 reductase activities (897), 21-hydroxylation of progesterone and 17-hydroxyprogesterone (899), NADPH-peroxidase activity (300),heme oxygenase (301), and steroid 17,20-lyase (Sob).Other techniques indicating the integrity of the system have been the increased turnover of the reductase in the microsomes of animals induced with potential substrates for the system such as phenobarbital (303),competitive inhibition between potential substrates (Sod), the influence of potential substrates on the activation energy of the reductase (SO6),and stimulation of NADPH-cytochrome P-450reductase activity (actually of the rate of anaerobic formation of the CO difference spectrum upon reduction with NADPH in microsomes) by potential substrates such as N-ethylmorphine (S06).

B. THENADH-CYTOCHROME b, REDUCTASE SYSTEM The history of the NADH-dependent microsomal electron transfer system has been quite different from that of the NADPH-dependent sys296. T. Omura, in “Microsomes and Drug Oxidations” (J. R. Gillette et al., eds.), p. 160. Academic Press, New York, 1969. 297. B. S. S. Masters, J. Baron, W. E. Taylor, E. L. Isaacson, and J. LoSpalluto, JBC 246, 4143 (1971). 298. F. Wadam, H. Shibata, M. Goto, and Y. Sakamoto, BBA 162, 518 (1968). 299. B. S. Masters, E. B. Nelson, B. A. Schacter, J. Baron, and E. L. Isaacson, Drug Metab. Disposition 1, 121 (1973). 300. E. G. Hrycay and P. J. O’Brien,ABB 157,7 (1973). 301. B. A. Schacter, E. B. Nelson, H. S. Marver, and B. S. S. Masters, JBC 247, 3601 (1972). 302. G. Betz, M. Roper, and P. Tsai, ABB 163,318 (1974). 303. H. Jick and L. Shuster, JBC 241,5366 (1966). 304. S. Orrenius and H. Thor, Eur. J . Biochem. 9, 415 (1969). 305. J. L. Holtzman and M. L. Carr, ABB 150,227 (1972). 306. J. L. Holtzman and B. H. Rumack, Biochemistry 12, 2309 (1973).

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tem in that the flavoprotein, NADH-cytochrome b, reductase (9, 307) and the cytochrome b, were purified and thoroughly characterized (17, 308, 309) long before data suggesting a function appeared. It was not until 1967 that it was suggested that cytochrome b, was the electron donor to a “cyanide sensitive factor” responsible for the desaturation of long chain fatty acids (310, 311, 311a) though it had been shown earlier that the CN-sensitive factor, and not cytochrome P-450,was involved in this process (318,313).The CN-sensitive factor has still not been characterized (284a). The system has been resolved and reconstituted, but the CN-sensitive factor used in the reconstitution was very crude (314). A requirement for lipid in the system had also been demonstrated (315, 316).An antibody to cytochrome b, has been shown to inhibit the desaturase of plasmalogen biosynthesis (317). C. POSSIBLE SYNERGISM BETWEEN

THE

Two MICROSOMAL SYSTEMS

Interaction between components of the NADH-dependent system and those of the NADPH-dependent system has been demonstrated, e.g., between NADPH-cytochrome P-450 reductase and cytochrome b, (318). The physiological significance of these interactions has been a matter of much controversy. A series of studies have demonstrated that the addition of NADH stimulates microsomal hydroxylation or oxidative N-demethylation utilizing the normal donor NADPH (295, 318-321). This has led to the proposal 307. P. Strittmatter and S. F. Velick, JBC 228, 785 (1957). 308. P. Strittmatter, in “Flavins and Flavoproteins” (E. C. Slater, ed.), Vol. 1, p. 325. Elsevier, Amsterdam, 1966. 309. P. Strittmatter, Fed. Proc. Fed. Amer. SOC.Exp. Biol. 24, 1156 (1965). 310. N. Oshino, Y. Imai, and R. Sato, Proc. Znt. Congr. Biochem., Yth, 1967 Abstracts, p. 725 (1968). 311. N. Oshino, Y. Imai, and R. Sato, J . Biochem. (Tokyo) 69, 155 (1971). 311a. N. Oshino and R. Sato, J . Biochem. (Toleyo) 69,169 (1971). 312. N. Oshino and Y.Imai, and R. Sato, BBA 128, 13 (1966). 313. J. L. Gaylor, N. J. Moir, H. E. Seifried, and C. R. E. Jefcoate, JBC 245, 5511 (1970). 314. T. Shimakata, K. Mihara, and R. Sato, J . Biochem. (Tokyo) 72, 1163 (1972). 315. P. Jones and S. Wakil, JBC 242,5267 (1967). 316. P. W. Holloway and S. J. Wakil, JBC 245, 1862 (1970). 317. F. Paltauf, R. A. Prough, B. S. S. Masters, and J. M. Johnston, JBC 249, 2661 (1974). 318. B. S. Cohen and R. W. Estabrook, ABB 143,37 (1971). 319. B. S. Cohen and R. W. Estabrook, ABB 143,46 (1971). 320. B. S. Cohen and R. W. Estabrook, ABB 143,54 (1971). 321. A. Hildebrandt and R. W. Estabrook, ABB 143,66 (1971).

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CHARLES H. WILLIAMS, JR.

that one of the two electrons required by cytochrome P-450 for hydroxylation or oxidative N-demethylation is supplied by NADH via NADHcytochrome b, reductase and cytochrome b, (320, 321). The role of NADH has been shown not to be obligatory since the terminal reactions proceed rapidly in reconstituted systems free of any cytochrome b, (322) and in microsomes in the absence of NADH (318).The stimulatory role of NADH seems to be twofold. The K,,, for NADPH in the reduction of cytochrome b, is of the same order of magnitude as its K,. in oxidative demethylation, 0.5 and 1 respectively (318). As will be seen in Sections VII and VIII, the specificity of the two flavoproteins for their respective reduced pyridine nucleotides is virtually absolute. Thus, reduction of cytochrome b, by NADPH in microsomes must be via NADPHcytochrome P-450 reductase. The presence of NADH then serves to prevent the interchain transfer of electrons from NADPH, i.e., competition between cytochrome b, and cytochrome P-450 for NADPH-cytochrome P-450 reductase is prevented by maintaining cytochrome b, reduced (321,322).The second function of NADH would then be that of electron donor to cytochrome P-450 by way of cytochrome b, (320422).Indeed, NADH can function as the sole electron donor in the hydroxylation of 3,4-benzpyrene1 and in this reaction cytochrome P-448 is more effective than is cytochrome P-450 (323).NADH has also been shown to stimulate NADPH-peroxidase (324) though there is some uncertainty whether the interchain electron transfer involves cytochrome b, or if the transfer is directly from NADH-cytochrome b, reductase to cytochrome P-450 (323,

a,

324)

The predicted stoichiometry in microsomal mixed function oxidations for NADPH :substrate: oxygen is 1:1:1 (326).In the absence of substrate NADPH oxidase is measured and in the presence of substrate this background oxidase activity is present though oxygen consumption increases (326,326).As the level of substrate increases the expected stoichiometry The addition of potential substrates which canis approached (326,326’). not be hydroxylated, such as perfluoro-n-hexane, leads to increased oxygen consumption and this has been termed “uncoupling” (327). It has been demonstrated that some of the electrons lost from the system are 322. A. Y. H. Lu, S. B. West, M. Vore, D. Ryan, and W. Levin, JBC 249, 6701 (1974). 323. E. G. Hrycay and P. J. O’Brien,ABB 160,230 (1974). 324. S. B. West, W. Levin, D. Ryan, M. Vore, and A. Y. H. Lu, BBRC 58, 516 (1974). 325. D. Y. Cooper, R. W. Estabrook, and 0.Rosenthal, JBC 238, 1320 (1963). 328. S.Orrenius, J . Cell Biol. 26,712 (1965). 327. V. Ullrich and H. Diehl, Eur. J . Biochem. 20,509 (1971).

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DEHYDROGENASES

lost as superoxide anion (328). It has been suggested that a further sparing effect of NADH in NADPH-dependent mixed function oxidations is in reducing superoxide that has been lost from the system ; it was shown that the sparing action depended upon cytochrome b, (329). The physiological significance of interchain electron transfer is still unknown. Efforts a t reconstitution of the two systems into a single lipid matrix are just beginning (330). The possible interactions are indicated below (331, 322) : NADPH +NADPH-cytochrome

P - 450

lipid:

cytochrome P - 450 -+02

reductase

/

NADH+NADH-cytochrome reductase

b,

cytochrome b, +CN-sensitive factor

+

0 2

I n one respect such schemes are quite misleading since they indicate a one-to-one relationship between components. It has been shown however that the ratio of NADPH-cytochrome P-450 reductase to cytochrome P-450is 1 to 50 (332).

D. NADPH-DEPENDENT MIXEDFUNCTION AMINEOXIDASE Microsomes contain, in addition to the two cytochrome reductases just discussed, a flavoprotein which catalyzes the mixed function oxidation of secondary and tertiary amines to the hydroxylamines and amine oxides, respectively (333, 334). This flavoprotein, which contains about 2 moles of phospholipid and 1 mole of FAD per 70,000 g of protein, is specific for NADPH (333, 334). The enzyme is also able t o catalyze the further oxidation of the hydroxylamines to nitrones (336).The reactions 328. M. J. Coon, T. A. Van der Hoeven, R. M. Kaschnita, and H. W. Strobel, Ann. N . Y . Acad. Sci. 212, 449 (1973). 329. H. Staudt, F. Lichtenberger, and V. Ullrich, Eur. J. Biochem. 46, 99 (1974). 330. A. I. Archakov, G. I. Bachmanova, V. M. Devichensky, I. Karuaina, N. S. Zherebkova, G. A. Alimov, G. P. Kuznetsova, and A. V. Karyakin, BJ 144, 1 (1974). 331. H. A. Sasame, S. S. Thorgeirsson, J. R. Mitchell, and J. R. Gillette, Pharmacologist 15, 170 (1973). 332. R. W. Estabrook and B. Cohen, in “Microsomes and Drug Oxidat[ons” (J. R. Gillette et al., eds.), p. 95. Academic Press, New York, 1969. 333. D. M. Ziegler, D. Jollow, and D. E. Cook, in “Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 507. Univ. Park Press, Baltimore, Maryland, 1971. 334. D. M. Ziegler and C. H. Mitchell, ABB 150, 116 (1972). 335. L. L. Poulsen, F. F. Kadlubar, and D. M. Ziegler, ABB 164, 774 (1974).

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CHARLES € WILLIAMS, I. JR.

are insensitive to carbon monoxide, indicating that cytochrome P-450 is not involved (336). It has been shown by immunochemical means that this flavoprotein is distinct from microsomal NADPH-cytochrome P-450 reductase (337). N-Hydroxylamines can also be reduced to the amine by a system consisting of NADH, NADH-cytochrome b, reductase, and a third protein fraction (336,338).

VII. NADH-Cytochrome b, Reductare

Microsomal NADH-cytochrome b, reductase and its acceptor substrate cytochrome bs are amphipathic proteins, that is, they are each composed of a hydrophobic domain and a soluble domain (74, 339, 340). The hydrophobic domains serve to anchor the proteins by strong noncovalent interaction with the lipid bilayer of the microsome. The soluble domains, containing the active sites, FAD or heme, project into the surrounding cytosol. The two domains, in each case, are connected by what is presumed (because of their proteolytic lability) to be rather flexible sections of polypeptide, imparting considerable mobility to the projecting catalytic domains. The proteins have been shown also to have translational mobility. Interaction between the reductase and the cytochrome is controlled by both types of mobility (34l-!&4). A. MOLECULAR PROPERTIES OF THE AMPHIPATHIC AND SOLUBLE FORMS OF THE REDUCTASE NADH-cytochrome b, reductase was originally solubilized from the microsomes by incubation with cobra venom, and in this form it was thoroughly characterized and its mechanism worked out (17, 307, 308) ; it has also been solubilized by the action of lysosomes normally contaminating the microsomal fraction (346-347). The two soluble forms were 336. F. F. Kadlubar, E. M. McKee, and D. M. Ziegler, ABB 158, 46 (1973). 337. B. S. S. Masters and D. M. Ziegler, ABB 145, 358 (1971). 338. F. F. Kadlubar and D. M. Ziegler, ABB 162,83 (1974). 339. A. Ito and R. Sato, JBC 243,4922 (1968). 340. L. Spate and P. Strittmatter, Proc. Nat. Acad. Sci. U.S. 68, 1042 (1971). 341. M. J. Rogers and P. Strittmatter, JBC 249, 895 (1974). 342. M. J. Rogers and P. Strittmatter, JBC 248, 800 (1973). 343. P. Strittmatter, M. J. Rogers, and L. Spate, JBC 247, 7188 (1972). 344. M. J. Rogers and P. Strittmatter, JBC 249, 5565 (1974). 345. S. Takesue and T. Omura, J . Biochem. (Tokyo) 87,259 (1970). 346. S. Takesue and T. Omura, J. Biochem. ( T o k y o ) 67,267 (1970). 347. P. Strittmatter, JBC 246, 1017 (1971).

3.

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FLAVIN-CONTAINING DEHYDROGENASES

TABLE VII AMINOACIDANALYSISOF VARIOUS FORMSOF NADHbs REDUCTASP CYTOCHROME

Amino acid Cysteic acid Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan Lysine Histidine Arginine Total residues Molecular weight

Detergentextracted A

Lysosomalextracted B

7 34 16 19 38 33 27 21 28 10 24 40 13 18 6 25 12 20

6 28 14 13 29 29 20 15 16 8 19 27 8 13 2 20 9 16

39 1 44,185

292 32 ,840

A-B

Chymotryptic core

6 12 2 5 13 5

6 28 11 12 26 28 19 14 19 8 19 29 9

1 6 2 6 9 4

7

5

13

4 5 3 4

2 20 9 16

99 11,340

288 32,526

Data from Spatz and Strittmatter ( 7 4 ) .

found to be essentially identical (347). The amphipathic form of the flavoprotein was solubilized with detergents and tends to aggregate in aqueous media (74, 348). Amino acid analyses of the two forms are shown in Table VII. They differ from one another by 100 residues, or about 11,000 in molecular weight. The amino acid content of the hydrophobic domain has been calculated by difference, and the composition is dominated by apolar amino acids (74). Treatment of the detergent-extracted enzyme with chymotrypsin results in a soluble form of the protein; its amino acid content is very similar to that of the lysosomal form (74). The spectra of the lysosomal-extracted and the detergent-extracted reductase are identical in the visible and near-ultraviolet, the extinction coefficient a t 461 nm being 10.6 mM-' cm-'. Differences between these spectra in the 260280-nm region are accounted for by the additional tryptophan and tyrosine residues ; the approximate extinction coefficients at the ultra348. E. s. Pnaiili and B. DeBernard, BBA 253,323 (1971).

156

CHARLES H . WILLIAMS, JR.

violet maxima are 62 and 90 mM-* cm-' for the lysosomal-extracted and detergent-extracted enaymes, respectively ( 7 4 ) .

B. REVIEW OF THE MECHANISM OF STRITTMATTER Studies leading to the proposal of a mechanism for NADH-cytochrome b, reductase were carried out with the snake venom-extracted enzyme, i.e., the catalytic domain. In Section VII,C, work will be described showing that this mechanism obtains for the enzyme bound to microsomes except that the rate limiting step changes because of a presumed restriction in the diffusion rate in the lipid milieu as compared to an aqueous medium (341, 342). Table VIII (9, 74, 307, 349-362) compares the various catalytic activities of the enzyme. Turnover with ferricyanide as acceptor is as rapid as with the natural acceptor. Oxygen and cytochrome c are very poor acceptors. The influence of substitutions on the pyridine ring is large, TABLE VIII SUBSTRATE SPECIFICITY OF NADH-CYTOCHROME bs REDUCTASE Electron donor

Electron acceptor

Turnover numbera

NADH* NADH" NADHd NADHa AcPyADHf PYA1ADHf NADPHe

Ferricyanide Cytochrome bs Cytochrome c Oxygen Ferricyanide Ferricyanide Ferricyanide

29,000 29,000 0 1.4 3,500 400 2.8

Moles substrate oxidized per mole enzyme per minute at pH 8.1 and 25". NADH, 125 p M ; ferricyanide, infinite (307).The apparent K , for NADH is 2.7 p M (ferricyanide, 5.5 p M ) (307). "NADH, 500 p M ; cytochrome b6, infinite, i.e., varied from 12 to 50 p M ; cytochrome c (25p M ) reduction monitored at 550 nm. The indicated K , for cytochrome br is 20 p M (74, 349). Electron transfer between cytochrome b6 and cytochrome c was demonstrated not to be limiting (9, 5007). d Strittmatter and Velick (9). * Strittmatter (360). f Donor, 125 p M ; ferricyanide, 250 p M (561,368). The apparent K , for ferricyanide is 2.2 p M (NADH, 120 p M ) (307).AcPyADH, 3-acetylpyridine adenine dinucleotide; PyAlADH, 3-pyridinealdehyde adenine dinucleotide. a

349. 350. 351. 352.

A. Loverde and P. Strittmatter, JBC 243,5779 (1968). P. Strittmatter, JBC 234,2665 (1959). P. Strittmatter, JBC 237,3260 (1962). P. Strittmatter, JBC 234,2661 (1959).

3.

157

FLAVIN-CONTAINING DEHYDROGENASES

I

0.100

I

I

I

I

I

300

3%

400

450

I

Wavelength (nm)

FIG. 15. Difference spectra of NADH-cytochrome b, reductase. Curve 1, enzyme reduced by NADH in the sample compartment and enzyme in the reference compartment; curve 2, the same but with NADPH. Curve 3 was generated by subtracting curve 2 from curve 1 (360).

a t least a t the concentrations of donor tested. NADPH is a very poor donor. These differences have been exploited to great advantage in the mechanistic studies. Protection of the reductase by NADH from inhibition by thiol group reagents suggested that the enzyme formed a stable complex with pyridine nucleotide (353). Such a complex was readily demonstrated by difference spectroscopy. When the enzyme was reduced by NADH or AcPyADH a prominent positive band was observed a t 317 nm (Fig. 15) ; this band was very small (and blue-shifted) when NADPH, a very poor substrate, was the reductant. Furthermore, addition of NAD' following NADPH resulted in a difference spectrum identical with that produced by NADH. The dashed line in Fig. 15 represents the absorption resulting from NAD' binding. Thus, this band was attributed to a reduced enzymeNAD' complex (350). 353. P. Strittmatter, JBC 233, 748 (1958).

158

CHARLIB H . WILLIAMS, JR.

F

5

s

9

Wavelength Lnm)

FIG 10. NADH-cytochrome bs reductase. Curve 1. oxidized enzyme. anaerobic conditions; curve 2, -after the addition of 1 equivalent'of PyAlADH ;o the oxidized enzyme; curve 3, after the addition of 1 equivalent of NADH to the oxidized enzyme; and curve 4, after the addition of 1 equivalent of NADH to the complex shown in curve 2 (564).

Reduction of the enzyme with NADH or PyAlADH give quite different and informative results as shown in Fig. 16 (364). Anaerobic reduction with 1 equivalent of NADH yields enzyme with spectral properties typical of FADH, but with very flat absorption extending into the near infrared. Reduction of 3-pyridinealdehyde adenine dinucleotide (PyAlADH) on the other hand is only partial, and the resulting spectrum can be duplicated by assuming a mixture composed of 0.35 equivalent of oxidized enzyme and PyAlADH, and 0.65 equivalent of reduced enzyme-PyAlAD+ complex; again the long wavelength absorption is present and a t a level higher than was the case with NADH. The fluorescence of the PyAlADH is fully quenched, indicating that it is bound. Addition of NADH following PyAlADH displaces this bound PyAlADH; curve 4 can be duplicated by assuming a mixture of 1 equivalent each of PyAlADH and reduced enzyme-NAD+ complex. Thus, the spectral evidence indicates complexes both between oxidized enzyme and reduced pyridine nucleotide and between reduced enzyme and oxidized pyridine nucleotide as shown:

3.

159

FLAVIN-CONTAINING DEHYDROGENASES

TABLE IX COMPARISON OF RATESFOR OVERALL TURNOVER WITH RATES OF ENZYME FLAVIN REDUCTION BY VARIOUS PYRIDINE NUCLEOTIDES (361, 366)"

kc for flavin Nucleotide NADH (a-D)NADH (8-D)NAI)H AcPyADH (a-D)AcPyADH (8-D)AcPyADH PyAlADH (a-D)PyA1ADH 0

Turnover number 68.6 18.7 68.6 8.33 0.8 8.33 0.96 0.11

reduction (sec-l) 69 19

-

8.5 0.97 8.5 1.02 0.12

H rate/D rate Turnover

k

3.66 1.00

3.6

10.4 1 .oo

8.8 1.0 8.5

-

8.7

-

Data from Strittmatter (361, 3666).

* Moles substrate oxidized per mole enzyme

FAD per second at pH 8.1 and 0'. Rate of absorbance change measured anaerobically in a rapid reaction spectrophotometer at 480 nm, pH 8.1, and 0'.

Furthermore, the displacement of PyAlADH by NADH demonstrates that the formation of these complexes is reversible (354). The rate of reduction of the FAD by the enzyme-bound NADH is the same as the rate of turnover with ferricyanide, indicating that the electron transfer from reduced pyridine nucleotide to enzyme-bound flavin is the rate limiting step. This was true with each pyridine nucleotide tested (Table IX). Large deuterium isotope effects are observed but only when the heavy atom is on the CY side of the pyridinium ring. Therefore, direct hydrogen transfer takes place stereospecifically (351, 355). As would be expected for complexes of this type, the stereospecificity is retained in reversing the reaction which can be effected in 0.5 M borate, pH 9.5. With (a-D)AcPyADH the deuterium is recovered totally on the CY side of the ring following reduction of the enzyme and displacement with borate (356). If flavin reduction is rate limiting, then during turnover the enzyme must be largely in the oxidized form. However, since the enzyme is not affected by thiol group reagents during turnover, the protection afforded by NADH must result from the reduced pyridine nucleotide-oxidized enzyme complex (308). Regardless of the rates of hydrogen transfer, which vary with different pyridine nucleotides, the 354. P. Strittmatter, JBC 238, 2213 (1963). 355. P. Strittmatter, JBC 240, 4481 (1965). 356. P . Strittmatter, JBC 239, 3043 (1964).

160

CHARLES H. WILLIAMS, JR.

rates of complex formation are very fast and were complete in the dead time of the rapid reaction spectrophotometer (354). Reoxidation of the enzyme-pyridine nucleotide complex by ferricyanide takes place in two one-electron steps. The rate of the first step is too rapid to measure and the rate of the second step can be measured only if NADPH is used to prereduce the enzyme. I n this case the complex with NADP' is virtually nonexistent and the changes measured are those of uncomplexed enzyme. These observations demonstrate that a species is formed within the mixing time and that it decays rapidly (78 sec-', Oo) . Repetition a t 10 nm intervals generates the spectrum of the intermediate which is that of the neutral semiquinone. The fact that both steps in the reoxidation of the enzyme-pyridine nucleotide complexes (i.e., when NADH is the reductant) are complete in less than 2 msec demonstrates that they meet the kinetic requirements of an intermediate, rapid formation and reoxidation (356). Reoxidation of the enzyme-pyridine nucleotide complexes by cytochrome b, also takes place in two steps. The rate of the first step is again too fast to measure. The rate of the second step is markedly dependent upon the pyridine nucleotide involved being 190, 68, and 18 sec-l, Oo for NADH, AcPyADH, and NADPH, respectively (356). A mechanism has been proposed for NADH-cytochrome b, reductase based on these elegant studies (Fig. 17) (308,366). The hydrogen which is stereospecifically and directly transferred has been printed in boldface. Catalysis proceeds in a clockwise direction, and after the first catalytic cycle the dissociation of oxidized pyridine nucleotide and the association of reduced pyridine nucleotide are shown as a single step since both are very rapid processes and this emphasizes that the thiol is not (kinetically speaking) exposed during catalysis. The interaction of the thiol and the

FIO.17. Mechanism for NADH-cytochrome bs reductase (308).

3.

161

FLAVIN-CONTAINING DEHYDROGENASES

pyridine nucleotide is hypothetical, but it should be pointed out that interactions between nucleophiles such as the thiol and pyridine nucleotides typically exhibit an absorption near 320 nm as is the case with the complexes formed by this enzyme (350). The interaction pictured between the flavin and a tyrosine residue is also hypothetical and will be discussed further in Section VI1,D.

C. MECHANISM OF THE REDUCTASE BOUNDTO

THE

MICROSOME

The mechanism for NADH-cytochrome b, reductase described in Section VI1,B was worked out with the soluble enzyme, and the question of its applicability to the interaction of the amphipathic proteins can now be considered. The turnover numbers of the detergent-extracted reductase in the reduction of ferricyanide or detergent-extracted cytochrome b, by NADH are 21 and 77% lower than those for the soluble proteins, respectively. The apparent K , values for NADH and cytochrome b, are raised about 2-fold. The addition of detergent has no effect on the turnover with ferricyanide but doubles the rate with cytochrome b,. These results indicate that reduction of ferricyanide catalyzed by the amphipathic reductase is nearly normal but that the reduction of cytochrome b, is inhibited by polymerization ( 7 4 ) . The detergent-extracted proteins can be rebound to microsomes ; indeed, vast excesses over endogenous levels can be bound (341444).All of the excess bound protein is active (343, 344) and electron transfer between different molecules of cytochrome b, does not occur a t an appreciable rate (341).These facts have two important consequences. First, it seems highly unlikely that a fixed array exists in which a reductase molecule interacts with a dozen or so cytochromes (the endogenous ratio) but rather that translational movement occurs in which the interacting partners are constantly changing. Second, if this picture is correct, the kinetic characteristics of the system bound to the microsome can be investigated by relatively conventional means, i.e., substrate concentrations can be varied and rate limiting steps identified. The essential difference is that diffusion may be restricted. The active sites on the other hand should have the same conformations as in the soluble proteins since in this picture they project into the aqueous phase. Rebinding of the reductase and the cytochrome b, to liposomes in a ratio of 1:13 completely restores the activity, inhibited in solution by polymerization. Thus, the phospholipid is an essential component in the interaction of the amphipathic proteins (342). The rate of reduction of cytochrome b, is dependent on its concentration in the microsome and on

162

CHARLES H. WILLIAMS, JR.

the enzyme concentration. The data indicate that no kinetically significant complex forms between the reduced reductase and cytochrome b, (3441).The relative rate effects on ferricyanide reduction of various analogs of NADH are the same for the soluble and bound reductase. However, the overall rate in the microsomes is limited by the rate of electron transfer to cytochrome b, rather than by the rate of intramolecular electron transfer between complexes as is the case in solution (S42). The data for the reduction of cytochrome b, could be fitted to the steady-state rate equation generated from the following sequence of reactions which also conform to the other characteristics of the bound system just outlined: kr

kl

+

EFAD NADH $ E:iEA

---t

E:i&

t 2

ENAD+

ENAD+ FADH'

+ Cyt bs"" + Cyt bs""

+ Cyt b P d E:kg+ + Cyt bsred k$ Efi:,D++ + NAD+ Rr

+ E:fEH. k7

+

EFAD

It can be seen that this sequence of reactions is simply an expression in kinetic terms of the mechanism shown in Fig. 17 which was derived for the soluble enzyme. Whereas in the soluble system k, was the rate limiting step, k, (and perhaps k,) is rate limiting in the bound system. The bound system is therefore diffusion-limited. The data indicate that in the intact microsome, NADH-cytochrome b, reductase is turning over a t about 60% of the maximum velocity (3.41). D. STRUCTURALSTUDIES The apoenzyme of NADH-cytochrome b, reductase is readily prepared by precipitation at low pH in the presence of high concentrations of potassium bromide (171). [Such inhibition of flavin rebinding by anions was first observed with the Old Yellow Enzyme (567).]The apoenzyme is stable a t low temperature and neutral pH. FAD recombination is followed with great sensitivity by the quenching of fluorescence which is total upon rebinding. When rebound, FAD is fully functional ; F M N and riboflavin also bind and give activities of 66 and 2076,respectively. The binding constants are FAD, less than 1 nM; FMN, 8 n M ; and riboflavin approximately 20 tJM indicating that all portions of the FAD molecule are important for tight binding. Reaction of the free thiols on the apo367. H. Theorell and A. Nygaard, Acta Chem. Scand. 8, 1649 (1954).

3.

FLAVIN-CONTAINING DEHYDROGENASES

163

enzyme has no effect on F M N rebinding, but rebinding is totally inhibited by reaction with 2 equivalents of iodine, provided the thiols have been protected from the iodine by prior reaction with N-ethylmaleimide. Apoenzyme thus modified has ultraviolet spectral properties consistent with the formation with a single diiodotyrosine residue. This residue may be important in its interaction with enzyme-bound FAD (171). Relative dissociation constants have been determined for the tight binding of NADH and AcPyADH to the apoenzyme; binding results in the quenching of pyridine nucleotide fluorescence. Their binding is strongly inhibited by ADP-ribose. NAD+ is bound 25 times less tightly than is NADH, but ADP-ribose is bound only 24 times less tightly, indicating that oxidation of the pyridinium ring weakens binding as was noted in the consideration of the mechanism (Section VI1,B). Binding of NADH to apoenzyme is inhibited by the reaction of a single thiol (358). Reversible dissociation of FAD is effected by lowering the p H ; FAD rebinds upon neutralization. The rebinding has been studied by optical and fluorescence spectroscopy and by the return of the ability of NADH to reduce the FAD. The kinetics are identical by the three criteria and indicate a half-time of 5 min for the refolding of the FAD and NADH binding sites ; simple first-order kinetics are approximated for only the first half of the reactions. FAD is not essential for the refolding but does accelerate it. The half-time for the reformation of the pyridine nucleotide binding site in the absence of flavin is about 8 min. The acceleration by FAD can also be demonstrated by observation of the changes in protein fluorescence during refolding (359, 360). The reactivity of the thiols in the native and denatured reductase has been investigated (352, 559, 360). In the native enzyme 3 thiols react with p-mercuribenzoate or mersalyl. Addition of an anionic detergent make an additional thiol available. At p H 2 where the apoenzyme is in equilibrium with FAD, 5 thiols react with p-mercuribenzoate or mersalyl. If the pH is raised, 2 equivalents of reagent are displaced as the protein refolds even if the FAD has been removed. This strongly indicates that the reactivity of a particular thiol is not merely a matter of its “availability” as is so often claimed, but is also a matter of the absolute reactivity of that thiol in its unique environment; this can only be tested with reversibly reacting reagents such as mercurials. Of the 2 thiols which become unreactive upon raising the pH, the reactivity of 1 is altered very quickly and the other with a half-time of about 5 min. ,As would 358. P. Strittmatter, JBC 236, 2336 (1961). 359. P. Strittmatter, JBC 242, 4630 (1967). 360. P. Strittmatter, in “Flavins and Flavoproteins” (K. Yagi, ed.), Vol. 2, p. 85. Univ. Park Press, Baltimore, Maryland, 1968.

164

CHARLES H. WILLIAMS, JR.

be expected with equilibria, the half-time for FAD rebinding is dependent on the mercurial concentration and upon its absolute reactivity; 1 mersalyl is displaced spontaneously in the rebinding and the second only after addition of excess thiol (569,360). N-Ethylmaleimide reacts with 3 of the 6 (Table VII) thiols in the soluble reductase, with 2 rapidly and with another slowly. In the presence of NADH, N-ethylmaleimide reacts with 1 thiol rapidly and with another slowly. It is concluded therefore that one of the rapidly reacting thiols is in the pyridine nucleotide binding site (36R). The reactivity of the lysyl residues in NADH-cytochrome b, reductase has been investigated and found to fall into three groups. Modification of the first most reactive group (about half the total) results in loss of the ability to interact with cytochrome bg. In the presence of NADH all but one of the remaining lysyl residues react, resulting in destabilization of the holoenzyme structure (349). The soluble reductase exists in two conformational states in the pH range 10.7 to 11.5, an active holoenzyme and an inactive flavoprotein (347). The inactive species is less compact, the FAD while still bound is in a more polar environment as are the 2 tryptophan residues, an additional thiol becomes reactive, and the protein is much more readily attacked by trypsin. Prolonged digestion leads to a compact FAD-binding peptide of 10,000 molecular weight (347).Tryptic digestion of the native soluble reductase leads to the removal of 47 amino acid residues; the remainder is composed of two polypeptide chains held together by noncovalent forces and has 65% of the normal activity. The two polypeptides can be separated and recombined into an active flavoprotein (361). Elucidation of the total primary sequence of NADH-cytochrome b, reductase is now well advanced (347).

E. THEFUNCTIONAL METHEMOGLOBIN REDUCTASE SYSTEM OF THE MATURE ERYTHROCYTE Methemoglobinemia has long been associated with the absence of an NADH-linked diaphorase (362, 363). However, flavoproteins isolated from the red cell were never sufficiently active to account for the methemoglobin reductase activity calculated to be necessary. It has now been shown that a methemoglobin reductase system of high activity is composed of soluble forms of cytochrome b, reductase and cytochrome b, 361. P. Strittmatter, R. E. Barry, and D. Corcoran, JBC 247, 2768 (1972). 362. E. M. Scott and D. D. Hoskins, Blood 13, 795 (1958). 363. E. M. Scott and J. C. McGraw, JBC 237,249 (1962).

3.

FLAVIN-CONTAINING DEHYDROGENASES

165

(364-366). The reductase has been purified by two groups of investigators (366, 3673, and its properties are very similar to those of NADH-cyto-

chrome b, reductase solubilized from liver microsomes by lysosomal digestion (347). FAD is the prosthetic group (366, 367), and the spectral properties are very similar to those of the microsomal enzyme (367). Its molecular weight is 34,000 (367). The turnover number with cytochrome b, coupled to cytochrome c is 1280 moles cytochrome b, reduced per minute per FAD which is quite low compared to the microsomal enzyme (366). Other activities are also low; turnover with ferricyanide is about 10,000 per minute. In the absence of cytochrome b,, it is virtually inactive with cytochrome c, oxygen, and methemoglobin, which is a pattern like that of the microsomal enzyme. The effects of ionic strength, pH, and EDTA are all similar to those of the microsomal enzyme (366). This reductase has been established as the missing component in methemoglobinemic erythrocytes (368). The mechanism of the presumed transition from a bound enzyme in the blast cell to a soluble enzyme in the mature erythrocyte will be of great interest.

VIII. NADPH-Cytochrome P-450 Reductase

The flavoprotein responsible for the reduction of cytochrome P-450 in microsomes has only recently been solubilized in a form active in the reconstitution of the many cytochrome P-450-linked systems (286, 369-371) (Section V1,A). In this form the reductase is not pure and has been characterized only in very preliminary experiments (369, 371). Studies on its mechanism have not been reported. Several modified forms of this enzyme have been isolated which turn over a t high rates with cytochrome c but are either inactive or only partially active with cytochrome P-450 (337, 372-374). They have been referred to as NADPH364. D. E. Hultquist and P. G. Passon, Nature (London), New Biol. 229, 252 (1971). 365. P. G. Passon, D. W. Reed, and D. E. Hultquist, BBA 275,51 (1972). 366. P. G. Passon and D. E. Hultquist, BBA 275,62 (1972). 367. F. Kuma and H. Inomata, JBC 247,556 (1972). 368. F. Kuma, S. Ishizawa, K. Hirayama, and H. Nakajima, JBC 247,550 (1972). 369. A. F. Welton, T. C. Pederson, J. A. Buege, and S. D. Aust, BBRC 54, 161 (1973). 370. T. A. Van der Hoeven and M. J. Coon, JBC 249,6302 (1974). 371. J. L. Vermilion and M. J. Coon, BBRC! MI, 1315 (1974). 372. T. Omura and S. Takesue, J . Biochem. (Tokyo) 67,249 (1970). 373. T. Iyanagi and H. S. Mason, Biochemistry 12,2297 (1973). 374. T. C. Pederson, J. A. Buege, and S. D. Aust, JBC 248, 7134 (1973).

166

CHARLES H. WILLIAMS, JB.

cytochrome c reductase (10, 11, 375). The connection between this enzyme and the microsomal hydroxylation system was made (376, 377) soon after the microsomal origin of the NADPH-cytochrome c reductase was established (10,11).

A. GENERALPROPERTIES NADPH-cytochrome P-450 reductase is composed of a single polypeptide chain of 70,000-80,000 molecular weight (369, 371, 373, 374) associated with one molecule of FAD and one molecule of F M N (370, 371, 373, 378, 379). These results apply to the enzyme whether solubilized by proteolytic digestion or by detergent extraction. The minimum molecular weight based on the flavin content is somewhat higher, 87,000 (3729, possibly indicative of the flavin lability observed upon irradiation in ammonium sulfate (380). The detergent-solubilized reductase has a lower flavin content, 0.64 and 0.79 moles of FMN and FAD per 79,000 g of enzyme (371). The absorbance ratio, 275 nm:455 nm of 6.5, indicates a relatively low content of aromatic amino acids (372, 374). The extinction of the flavins a t 455 nm is 10.7 mM-l cm-l (373). The demonstration of the amphipathic nature of NADH-cytochrome b, reductase (74) suggests the question of whether NADPH-cytochrome P-450 reductase, another microsomal enzyme, is also amphipathic. The molecular weight of the detergent-solubilized reductase has been determined in two laboratories by sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis, both finding values of 79,000 ($6’9,371). The molecular weight of the bromelain-solubilized enzyme is 71,000 (374) and that of the trypsin-solubilized enzyme is 68,000 (373), both determined by SDS gel electrophoresis. This is suggestive of a molecular weight difference of 10,000. However, if this is correct, the minimum molecular weight derived from flavin analysis would indicate that the trypsin-solubilized reductase is approximately 20% flavin depleted (873). Until enzyme with full flavin complement can be prepared, comparative amino acid analysis such as that used to demonstrate the difference between 375. B. L. Horecker, JBC 183,593 (1950). 376. S.Orrenius, G. Dallner, and L. Ernster, BBRC 14, 329 (1964). 377. D. Y. Cooper, S. Levin, S. Narashimhulu, and S. Rosenthal, Science 147, 400 (1965). 378. H. Kamin, i n “Reactivity of Flavins-The Proceedings of a Symposium

Dedicated to the Late Professor Leonos Michaelis under the Auspices of. the Japanese Biochemical Society” (K. Yagi, ed.), p. 137. Univ. Park Press, Baltimore, 1975. 379. B. S. S.Masters, R. A. Prough, and H. Kamin, personal communication. 380. J. P. Baggott and R. G. Langdon, JBC 245, 5888 (1970).

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the forms of NADH-cytochrome b, reductase (74) will not settle this question. The NADPH-cytochrome c reductase of yeast is an FMN-containing enzyme (381,382). It is also of microsomal origin (383).

B. CATALYTIC ACTIVITIE~ OF

THE

REDUCTASE

The turnover of NADPH-cytochrome P-450reductase with its natural acceptor can only be studied as a coupled reaction in which the cytochrome P-450 is acting as a hydroxylase or N-demethylase. In the absence of a hydroxylatable substrate, cytochrome P-450acts as an oxidase (370).The oxidase activity of the reductase is very low (10). The various modified forms of NADPH-cytochrome P-450 reductase have been routinely assayed as cytochrome c reductases. The kinetics of this reaction have been studied both by steady-state and by rapid reaction methods. The pH optimum is 7.6 to 8.2 (10, 11, 375)) and the activity increases with ionic strength being optimal a t 0.2 M phosphate, pH 7.6 (11, 384).Varying the NADPH concentration at a series of cytochrome c concentrations results in a family of parallel lines in reciprocal plots. V,,, is 1200 moles cytochrome c reduced per minute per mole of flavin a t infinite concentrations of both substrates (245); K , for NADPH is 4 p M and for cytochrome c is 5.5 pM (10, 245). The dissociation constant for the reductase-cytochrome c complex is 4.6 pM (380).NADP' and AMP are competitive inhibitors of the reaction (10, 385). The K , for both rises with ionic strength, an indication that one of the effects of high salt concentrations is to displace NADP'. At 0.1 M phosphate, the K i is 2 pM for both inhibitors (11). Specific activities are usually reported on a protein basis and range in recent studies from 40 pmole cytochrome c per minute per milligram for the lipase- and trypsin-solubilized enzymes (337, 372, 373) to 56 units per milligram for the bromelain-solubiliaed reductase (374). Comparable figures for the detergentsolubilized enzyme have not been measured a t the same temperature (371).The turnover number at infinite concentration of both substrates with dichlorophenolindophenol as acceptor is virtually identical with that with cytochrome c a t the same pH, though the pH optimum for dichlorophenolindophenol may not be the same (386).The reductase is also very 381. E. Haas, B. L. Horecker, and T. R. Hogness, JBC 136, 757 (1940). 382. E. Haas, C. J. Harrer, and T. R. Hogness, JBC 143, 341 (1942). 383. G. Schatz and J. Klima, BBA 81, 448 (1964). 384. M. H. Bilimoria and H. Kamin, Ann. N . Y . Acud. Sci. 212, 428 (1973). 385. E. F. Neufeld, N. 0. Kaplan, and S. P. Colowick, BBA 17, 526 (1955). 386. B. S. S. Masters, M. H. Bilimora, H. Kamin, and Q. H. Gibson, JBC 240, 4081 (1965).

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CHARLES H. WILLIAMS, JR.

active with menadione (386), ferricyanide, and neotetrazolium (10) as acceptors. A very important conclusion was reached based on the effect of p-mercuribenzoate on the NADPH oxidase and the NADPH-cytochrome c reductase activities of microsomes, namely, that the natural acceptor might be a component reactive with oxygen and involved in hydroxylations or demethylations (11). It was found that in the absence of cytochrome c, the oxidase activity was largely inhibited by p-mercuribenzoate. In the presence of cytochrome c, NADPH oxidation exceeded cytochrome c reduction in the absence of p-mercuribenzoate and the two rates equaled each other in the presence of p-mercuribenzoate. Thus, a mercurial sensitive oxidase distinct from the reductase was indicated, and this component was hypothesized to be connected with hydroxylation and/or demethylation (11). The influence of mercurials on the NADPH-cytochrome c reductase activity is complex. The activity in microsomes is stimulated about 50% by p-mercuribenzoate (11) . Mersalyl inhibits the NADPH-cytochrome c reductase activity (587). The lipase-solubilized reductase is inhibited by p-mercuribenzoate, is protected from this inhibition by NADPH, and the inhibition is relieved by thiols (10). Careful titration of this enzyme with p-mercuribenzoate a t pH 6.5 results in an almost 3-fold stimulation upon addition of 2 moles of mercurial per flavin; the control activity is again observed when 7 equivalents have been added. At pH 7.7, a stimulation of 70% is seen with 1 equivalent and loss of activity is complete (extrapolated) with 6 equivalents (245).The protection of the enzyme by NADPH against mercurial inhibition is reminiscent of the effects with NADH cytochrome b, reductase (360). NADPH-cytochrome P-450 reductase catalyzes the reduction of ferric ion to ferrous ion in the presence of chelators (584,388). This reaction is similar to the peroxidation of microsomal lipids which has been shown to be dependent on the NADPH oxidase system of microsomes (389,390). Lipid peroxidation is thought to be involved in prostaglandin biosynthesis (590392).The microsomal system can be mimicked in the peroxidation of extracted microsomal lipid by a combination of NADPH, NADPHcytochrome P-450 reductase, and ferric ion chelated with EDTA (374, 387. M. R. Franklin and R. W. Estabrook, ABB 14'3,318 (1971). 388. M. M. Weber, H. M. Lenhoff, and N. 0. Kaplan, JBC 220, 93 (1966). 389. P. Hockstein and L. Ernster, BBRC 12,388 (1963). 390. H. E. May and P. B.McCay, JBC 243, B98 (1968). 391. W. Lands, R. Lee, and W. Smith, Ann. N . Y . Acad. Sci. 180, 107 (1971). 392. B. Samuelsaon, Progr. Biochem. Pharmacol. 5,109 (1969).

3.

FLAVIN-CONTAINING DEHYDROGENASES

169

393). I n this model system, as contrasted with the simple ferric ion reductase activity of the flavoprotein (388),the metal is not the ultimate electron acceptor but presumably serves the dual role of oxygen activation and electron carrier. The reaction may involve superoxide anion since it is inhibited by superoxide dismutase (erthrocuprein) (394). Xanthine plus xanthine oxidase can also serve as electron donor, and this latter model system is also inhibited by superoxide dismutase (396).Superoxide dismutase also inhibits the menadione-mediated NADPH oxidase activity of NADPH-cytochrome P-450 (396) as well as the reconstituted benzphetamine hydroxylation system (397). The involvement of NADPHcytochrome P-450 reductase in microsomal lipid peroxidation has been confirmed by the demonstration that the reaction in microsomes is totally inhibited by antibody to the purified reductase (374). It has been suggested that lipid peroxidation by microsomes requires another component, in addition to the reductase, which takes the place of the ferric ion chelate in the model system (374).

C. MECHANISM Studies on the mechanism of NADPH-cytochrome P-450 reductase have been carried out thus far only with the trypsin- or lipase-solubilized forms. Assuming that this enzyme is composed of several semi-autonomous domains, and assuming further that modification during solubilization is restricted to the domain involved in the interaction with cytochrome P-450,then, as was the case with NADH-cytochrome b, reductase, mechanism studies on the soluble enzyme will contribute to the ultimate understanding of the operation of the reconstituted system. The fact that the soluble reductase is composed of a single polypeptide chain gives hope that the modification is a subtle one. Reduction of the lipase-solubilized enzyme by NADPH is more rapid than either turnover with cytochrome c or the rates of reconstituted systems (646).In rapid reaction spectrophotometric studies, changes a t 550 nm are taken is indicative of flavin radical (FlH) ; the oxidized (Fl) and reduced (FlH,) forms of the enzyme have negligible absorbance a t this wavelength. Changes a t 500 nm indicate formation of FlH, (negative) or reoxidation of FlH, (positive) ; F1 and F l H are isosbestic a t 500 nm. Both FlH and FlH, are formed a t rates consistent with their 393. 394. 395. 396. 397.

T. C.Pederson and S.D. Aust, BBRC 48,789 (1972). R. A. Prough and B. S.S.Masters, Ann. N . Y . Acad. Sci. 212,89 (1973). T. C. Pederson and S.D. Aust, BBRC 52,1071 (1973). T. Lyanagi and I. Yamazaki, BB.4 172,370 (1969). H. W. Strobe1 and M. J. Coon, JBC 246, 7826 (1971).

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CHARLES H. WILLIAMS, JR.

participation in catalysis. Although F1H appears to be formed more rapidly than FlH,, there is no lag in FlH, production and it is suggested that FlH2 is formed both directly (in the dead time of the apparatus, ca. 3 msec) and from F1H. The rapid appearance of FIH2 is observed even a t very low ratios of NADPH to flavin (0.2) (246, 898). Anaerobic reduction with ratios of NADPH to flavin of less than 0.5 or aerobic reduction with excess NADPH lead to the production of a spectrally characteristic species of the enzyme which is not oxidized by oxygen or by cytochrome c (246). This species has long wavelength absorbance, characteristic of the neutral flavin semiquinone, but the absorbance in the 455-nm region is much higher than that of a typical neutral semiquinone ( 1 ) . Masters et al. (,%?46) interpreted this species to be the 2-electron-reduced enzyme. Since the enzyme contains two flavins per mole they maintained that each flavin was half-reduced (246). However, it has been pointed out in several discussions that the spectral characteristics of this species strongly suggested that it is a mixture of oxidized flavin and semiquinone flavin (399). Anaerobic reaction with excess NADPH leads to further reduction but only about half the long wavelength absorbance disappears. The authors interpreted this species as being fully reduced (four electron) enzyme with a small amount of semiquinone (246). The ratio of NADPH oxidized to cytochrome c reduced is 1 : l in experiments in which stoichiometric enzyme is used, but this ratio approaches the expected value of 2 in the presence of catalytic quantities of enzyme. This, taken together with the fact that cytochrome c does not fully reoxidize the enzyme, indicated to the authors that the enzyme was cycling in catalysis between the 4-electron-reduced and the 2-electron-reduced species (246, 398). Furthermore, the kinetics of enzyme reduction suggested that the flavins were not independent but rather interacted with one another (386,398). A reexamination of the mechanism of NADPH-cytochrome P-450 reductase has followed the crucial finding that all forms of this enzyme (detergent-, lipase-, and trypsin-solubilized) contain equimolar amounts of FAD and F M N suggesting that the flavins might have distinct roles (373). Distinct roles have been found for the FAD and FMN in sulfite oxidase (400, 401). The static spectral results with the lipase-solubilized 398. H. Kamin, B. S.S. Masters, and Q. H. Gibson, in “Flavins and Flavoproteins” (E. C. Slater, ed.), Vol. 1, p. 306. Elsevier, Amsterdam, 1966. 399. P. Hemmerich, in “Flavins and Flavoproteins” (E. C. Slater, ed.), Vol. 1, p. 319. Elsevier, Amsterdam, 19613. 400. L. M. Siegel, H. Kamin, D. C. Rueger, R. P. Presswood, and Q. H. Gibson, in “Flavins and Flavoproteins” (H. Kamin, ed.), Vol. 3, p. 523. Univ. Park Press, Baltimore, Maryland, 1971. 401. E. J. Faeder, P. 8. Davis, and L. M. Siegel, JBC 249, 1599 (1974).

3.

FLAVIN-CONTAINING DEHYDROGENASES

171

Wavelength (nm)

FIG.18. Anerobic titration of NADPH-cytochrome P-450 reductase with NADPH. Curve 1, oxidized enzyme; curves 2-6 after the addition of 0.16, 0.24, 0.49, 0.98, and 1.4 moles of NADPH per mole of total enzyme-bound flavin. The inset, B, shows the changes at 455 and 585 nm as a function of the NADPH added (409).

enzyme cited above have been confirmed with the trypsin-solubilized enzyme but reinterpreted. On the basis of EPR quantitation, NADPH titrations, and ferricyanide titrations, the air stable species has been shown to be the 1-electron-reduced enzyme. The species formed by excess NADPH has been shown to be a roughly equimolar mixture of 3-electronand 4-electron-reduced enzyme (373, 402). The anaerobic titration of the enzyme with NADPH is shown in Fig. 18. It can be seen that curves 1, 2, and 3 are isosbestic a t 500 nm; in a separate experiment curve 3 is shown to be virtually identical with the spectrum of the air stable species. Furthermore, the redox state of the enzyme in curve 3 is produced by the addition of one electron per two flavins. Addition of increasing amounts of NADPH results in further reduction of the enzyme-to the final mixture of 3-electron-reduced and 4-electron-reduced enzyme. Titration of the enzyme with dithionite gives a clear end point upon the addition of 2 moles of reductant per mole of enzyme (two flavins) ; thus, 402. T. Iyanagi, N. Makino, and H. S. Mason, Biochemistry 13, 1701 (1974).

172

CHARLES H. WILLIAMS, JR.

no electron accepting groups other than the two flavins are present (402). It is reemphasized that these experiments were carried out with trypsinsolubilized enzyme (373). Recalling the apparent molecular weight difference, it is possible that this modified form is catalytically different from the lipase-solubilized form (337, 409). Titrations with NADPH and ferricyanide have been repeated with the lipase-solubilized enzyme (404). The results indicate that 2 moles of ferricyanide are required to reoxidase the air stable form of the enzyme. Redox potentials have been determined for each of the steps of reduction of the trypsin-solubilized reductase (402): step 1, one electron consumed, Eo' = -109 mV; step 2, two electrons consumed, EO' = -276 mV; and step 3, one electron consumed, Eo' = -371 mV at p H 7.0, 25O. As expected, the redox potential of step 3 is more negative than the potential of the NADPH-NADP+ couple and was determined from the dithionite titration. The overall potentiometric-spectrophotometric titration curves could be very closely fitted with a computer-generated curve based on the assumptions of four one-electron reduction steps and extinction coefficients of 4.9 and 4.5 mM-l cm-' for the semiquinones, F1,H and F1,H; the 23', values assumed for steps 2 and 3 were -270 and -290 mV. The precise fit was very sensitive to all of the assumptions ( 4 M ) . Three alternative mechanisms were proposed based only on the thermodynamic data (402). All of these assumed distinct functions for each flavin and interaction between the flavins. They also assumed that electrons would be transferred to cytochrome P-450one a t a time; this has been shown to be the case with cytochromes P-450that receive electrons from iron-sulfur proteins rather than from the flavoprotein directly (or through the indirect mediation of lipid) (406, 406). One of these mechanisms ( 4 2 ) is shown below. It seems to fit best with the kinetic data determined for the lipase-solubilized reductase (246,398).In this scheme, SH is a hydroxylatable substrate and SOH its hydroxylated product, and F1, and F1, are the high potential and low potential flavins, respectively.

+ H+ + e NADP+ + FliHn FliHz + Flz FliH. + FlzH. Flz + P-45O'+SH + H+ FlrH. + P-450"SH P-45OS+--SH + 02 P-450*+--SH-01 NADPH + FIIH-/FII+ H+ 2 NADP+ + FliHn/FlzH* FllHz + P-4501+--SH-Oz S FI1H. + P-450a+ + SOH + OHNADPH

Fl1

$

F!

403. B. S. S. Masters, C. H. Williams, Jr., and H. Kamin, "Methods in Enzymology" Vol. 10, p. 535, 1967. 404. B. S. S. Masters, R. A. Prough, and H. Kamin, Biochemistry 14,607 (1975). 405. J. J. Huang and T. Kimura, BBRC 44, 1065 (1971). 406. C. A. Tyson, J. D. Lipscomb, and I. C. Gunsalus, JBC 247, 5777 (1972).

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FLAVIN-CONTAINING DEHYDROGENASES

173

The mechanism hypothesizes that electrons are donated to the cytochrome P-450 by the two couples with potentials near -276 mV. This mechanism together with the others that are consistent with the thermodynamic data (408) now form the working hypotheses for kinetic experiments which hopefully will distinguish between them and further define the mechanism of this important system.

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Metal-Containing Flavoprotein Dehydrogenases YOUSSEF HATEFI DIANA L. STIGGALL I . Introduction . . . . . . . . . . . . . . . . I1. NADH Dehydrogenases . . . . . . . . . . . . . A. NADH Dehydrogenase of Mammalian Mitochondria . . B. NADH Dehydrogenases of Yeast . . . . . . . . C . NADH Dehydrogenase of Azotobader uinelandii . . . I11. Succinate Dehydrogenases . . . . . . . . . . . . A . Mammalian Succinate Dehydrogenase (EC 1.3.99.1) . . B. Succinate Dehydrogenase in Microorganisms . . . . I V . LGlycerol-%phosphate Dehydrogenase (EC 1.1.99.5) . . . . V. Choline Dehydrogenase (EC 1.1.99.1) . . . . . . . . . V I . Lactate Dehydrogenases. . . . . . . . . . . . . A . L( +)-Lactate: Cytochrome c Oxidoreductase (Cytochrome b z ) (EC 1.1.2.3) . . . . . . . . B. D( -)-Lactate:Cytochrome c Oxidoreductase (EC 1.1.2.4). C . DZHydroxyacid Dehydrogenase (EC 1.1.99.6). . . . V I I . Nitrite Reductases (EC 1.6.6.4) . . . . . . . . . . VIII . Adenylyl Sulfate Reductases (EC 1.8.99.2) . . . . . . . I X . Sulfite Reductases ( H a : NADPH Oxidoreductases) (EC 1.8.1.2) . A. NADPH-Sulfite Reductases . . . . . . . . . B . Reduced Methyl Viologen-Sulfite Reductases . . . . X Addendum . . . . . . . . . . . . . . . . .

.

175 177 177 216 22 1 222 222 254 256 260 263 263 269 272 273 279 286 287 295 295

.

1 Introduction

Flavoproteins are involved in a large variety of key metabolic reactions in all forms of life. They catalvze over a potential span of several hundred millivolts oxidation-reduction reactions involving alkanes. alkenes. alco175

176

YOUSSEF HATEFI AND DIANA L. STIGGALL

hols, aldehydes, ketones, inorganic and organic acids, amines,. thiols, disulfides, quinones, nicotinamide-adenine dinucleotides, purines, pyrimidines, pteridines, and transition metal complexes. They can also catalyze one- and two-electron reduction of molecular oxygen. Many flavoproteins contain metal such as iron, molybdenum, and zinc. The combination of flavin and metal often serves to adjust electron transfer between singleelectron and double-electron donors and acceptors. Multiple-electron reduction of an acceptor without detectable loss of intermediates is achieved, as will be seen below, by the device of having multiple flavins and metals in the same enzyme molecule. Excellent sources of recent publication on the chemistry of flavins and flavoproteins are available (1-6).

The present chapter is concerned with metal-containing flavoprotein dehydrogenases. The enzymes discussed are respiratory chain-linked NADH dehydrogenases, succinate dehydrogenases, ~-glycerol-S-phosphate dehydrogenase, choline dehydrogenase, L (+)-lactate: cytochrome c oxidoreductase, D (-) -lactate :cytochrome c oxidoreductase, D-2-hydroxyacid dehydrogenase, nitrite reductases, adenylyl sulfate reductases, and sulfite reductases. Among these, NADH and succinate dehydrogenases have been the subject of intensive study for the past 20 years, in part because of their importance as the principal electron entry points into the mitochondria1 respiratory chain. Progress of research on these two enzymes has been reviewed by various workers. One laboratory alone has produced more than twenty reviews of various sorts during as many years (see, for example, 6-96).These are excellent reference sources, and 1. E. C. Slater, ed., “Flavins and Flavoproteins,” Symp. Adn Flavoproteins. Elsevier, Amsterdam, 1986. 2. K. Yagi, ed., “Flavins and Flavoproteins,” 2nd Int. Symp. Univ. Park Press, Baltimore, Maryland, 1968. 3. H. Kamin, ed., “Flavins and Flavoproteins,” 3rd Int. Symp. Univ. Park Press, Baltimore, Maryland, 1971. 4. T. E. King, H. S. Mason, and M. Morrison, eds., “Oxidases and Related Redox Systems.” Univ. Park Press, Baltimore, Maryland, 1973. 6. W. Lovenberg, ed., “Iron-Sulfur Proteins,” Vols. 1 and 2. Academic Press, New York, 1973. 6. T. P. Singer and E. B. Kearney, in “The Proteins” (H. Neurath and K. Bailey, eds.), 1st ed., Vol. 2, Part A, p. 123. Academic Press, New York, 1964. 7. T. P. Singer and E. B. Kearney, Proc. Znt. Congr. Biochem., 4th, 1968 Symposium 11,Vol. 11,209 (1960). 8. T. P. Singer, in “Biological Structure and Function” (T. W. Goodwin and 0. Lindberg, eds.), Vol. 2, p. 103. Academic Press, New York, 1961. 9. T. P. Singer, S. Minakami, and R. L. Ringler, Proc. Znt. Congr. Biochem., bth, 1961 Symposium V, p. 174 (1963). 10. T. P. Singer, “The Enzymes,” 2nd ed., Vol. 7, p. 345, 1963. 11. T. P. Singer, “The Enzymes,” 2nd ed., Vol. 7, p. 383, 1963.

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177

skillfully propound the position of that laboratory regarding NADH and succinate dehydrogenases. I n recent years, new information has appeared, however, which has clarified many of the basic controversial issues and erroneous assumptions. In the following account of these two enzymes, it will be attempted, therefore, to offer a critical analysis of the major aspects rather than to present an exhaustive review of the chronological development of the field. II. NADH Dehydrogenarer

A. NADH DEHYDROGENASE OF MAMMALIAN MITOCHONDRIA Preparations of NADH dehydrogenase from mammalian mitochondria may be divided into three types: (1) NADH-ubiquinone reductase or complex I of the electron transport system, (2) the high molecular weight NADH dehydrogenases, and (3) the low molecular weight NADH dehy12. T. P. Singer and T. Cremona, in “Oxygen in the Animal Organism” (F. Dickens and E. Neil, eds.), p. 179. Pergamon, Oxford, 1964. 13. T. P. Singer, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 448. Wiley, New York, 1965. 14. T. P. Singer, in “Non-Heme Iron Proteins” (A. San Pietro, ed.), p. 349. Antioch Press, Yellow Springs, Ohio, 1965. 15. T. P. Singer, Compr. Biochem. 14, 127 (1966). 16. T. P. Singer, E. Rocca, and E. B. Kearney, in “Flavins and Flavoproteins,” Symp. Adn Flavoproteins (E. C. Slater ed.), p. 391. Elsevier, Amsterdam, 1966. 17. T. P. Singer, in “Biological Oxidations” (T. P. Singer, ed.), p. 339. Wiley (Interscience), New York, 1968. 18. T. P. Singer and M. Gutman, in “Pyridine Nucleotide-Dependent Dehydrogenases” (H. Sund, ed.), p. 375. Springer-Verlag, Berlin and New York, 1970. 19. T. P. Singer and M. Gutman, Advan. Enzymol. 34,79 (1971). 20. T. P. Singer, M. Gutman, and E. B. Kearney, in “Biochemistry and Biophysics of Mitochondria1 Membranes” (G. F. Azzone et al., eds.), p. 41. Academic Press, New York, 1972. 21. T. P. Singer, in “Biochemical Evolution and the Origin of Life” (E. Schoffeniels, ed.), p. 203. North-Holland Publ., Amsterdam, 1971. 22. T. P. Singer, D. J. Horgan, and J. E. Casida, in “Flavins and Flavoproteins,” 2nd Int. Symp. (K. Yagi, ed.), p. 192. Univ. Park Press, Baltimore, Maryland, 1968. 23. T. P. Singer, M. Gutman, and V. Massey, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 1, p. 225. Academic Press, New York, 1973. 24. T. P. Singer, E. B. Kearney, and M. Gutman, in “Biochemical Regulatory Mechanisms of Eukaryotic Cells” (E. Kun and S. Grisolia, eds.), p. 271. Wiley (Inter: science), New York, 1972. 25. T. P. Singer, E. B. Kearney, and W. C. Kenney, Advan. Enzymol. 37, 189 ( 1973).

26. T. P. Singer, E. B. Kearney, and B. A. C. Ackrell, in “Mechanisms in Bioenergetics” (G. F. Azzone et at., eds.), p. 485. Academic Press, New York, 1973.

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YOUSSEF HATEFI AND DIANA L. STIGGALL

Mitochondria

FIa. 1. Scheme showing the fractionation of beef heart mitochondria into enzyme complexes I, 11, 111, IV, and V with the use of deoxycholate (DOCA),cholate, ammonium sulfate, and ammonium acetate. From Hatefi et al. (31).

drogenases. The latter two have also been referred to as type I and type I1 NADH dehydrogenases, respectively. As will be seen, the two preparations of NADH dehydrogenase are related to complex I, except that one appears to have irreversibly lost ubiquinone reductase activity and the other has grossly modified enzymic properties.

1. NADH-Ubiquinone Reductase (Complex I ) NADH-ubiquinone reductase was isolated by Hatefi et al. in 1961 (27-29). A procedure was developed for the resolution of the mitochondrial electron transport system into four enzyme complexes. Recently, a fifth fraction, which is capable of energy conservation and ATP-PI exchange, was also isolated (30,31). The overall scheme for the isolation of the five component enzyme complexes of the mitochondria1 electron transporhxidative phosphorylation system is given in Fig. 1. It is seen 27. Y. Hatefi, A. G. Haavik, and D. E. Griffiths, BBRC 4, 441 and 447 (1961). 28. Y. Hatefi, A. G. Haavik, and D. E. Griffiths, JBC 237, 1676 (1962). 29. Y. Hatefi, Compr. Biochem. 14, 199 (1966). 30. Y. Hatefi, D. L. Stiggall, Y. Galante, and W. G. Hanstein, BBRC 61, 313 ( 1974). 31. Y. Hatefi, W. G. Hanstein, Y. Galante, and D. L. Stiggall, Fed. Proc., Fed. Amer. Ism. Em. Biol. 34,1699 (1975).

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179

TABLE I COMPOSITION OF COMPLEX 10

a

Component

Concentration (per mg protein)

FMN (acid-extractable) Nonheme iron Acid-labile sulfide Ubiquinone- 10 Cytochromes Lipids

1.4-1.5 nmoles 23-26 ng-atoms 23-26 nmoles 4.2-4.5 nmoles
From Hatefi et al. (88, 38).

that the procedure, which yields complexes I to V in high yield, is relatively simple. Deoxycholate and cholate are used in combination with KC1, a neutral salt of low ionic strength, for differential solubilization of mitochondria1 components ; and ammonium acetate and ammonium sulfate are employed for fractionation and isolation of the fragments. The details of composition and enzymic properties of the enzyme complexes shown in Fig. 1 have been described (99, 39). It was also shown in 1961 that complexes I, 11, 111, and IV interact stoichiometrically to reconstitute a particulate unit with full activity and inhibitor-response properties for electron transfer from NADH and succinate to molecular oxygen (97,29, 33). These and the subsequent studies of other laboratories (3436) have established that complex I represents in a highly purified form the segment of the electron transport system from NADH to ubiquinone. The complex carries site 1 of energy conservation and, under appropriate conditions, it is capable of interaction with the isolated coupling factors of mitochondria to couple the energy derived from oxidation of NADH by ubiquinone to the synthesis of ATP (3 5 ,5 6 ). a. Composition. The composition of complex I is shown in Table I. Essentially all the flavin of complex I is acid-extractable, and according to Rao et al. (37), more than 96% of the flavin is FMN. The ratio of 32. Y. Hatefi, W. G. Hanstein, K. A. Davis, and K. S. You, Ann. N. Y . Amd. Sci. 227,504 (1974). 33. Y . Hatefi, A. G. Haavik, L. R. Fowler, and D. E. Griffiths, JBC 237, 2661 (1962). 34. N. R. Orme-Johnson, W. H. Orme-Johnson, R. E. Hansen, H. Beinert, and Y. Hatefi, BBRC 44,446 (1971). 35. C. I. Ragan and E. Racker, JBC 248,2563 (1973). 36. C. I. Ragan and E. Racker, JBC 248,6876 (1073). 37. N. A. Rao, S. P. Felton, F. M. Huennekens, and B. Mackler, JBC 238, 449 (1963).

180

YOUSSEF HATEFI AND DIANA L. STIGQALL I

I

I

1

'

I

mM-' 0,

FIQ.2. Lineweaver-Burk plot of ubiquinone-6 reduction by complex ( I. Y. Hatefi, unpublished).

flavin:iron:labile sulfide is 1:16-18: 16-18. Preparations of complex I usually contain a total of about 0.1 nmole of cytochromes b c1 per mg protein, which represents a contamination of complex I by about 0.8% complex 111. As in beef heart mitochondria, the ubiquinone of complex I is ubiquinone-10 and the phospholipids, which comprise more than 90% of the total lipid of complex I, are phosphatidylcholine, phosphatidylethanolamine, and cardiolipin (38).The latter appears to be more firmly bound to complex I proteins than phosphatidylcholine and phosphatidylethanolamine (36).The minimum molecular weight of complex I is 6.5-7 X lo5 g protein (or 8-8.5 x lo5 g protein plus lipid) per mole of flavin. However, acrylamide gel electrophoresis of complex I treated with sodium dodecyl sulfate and mercaptoethanol reveals the presence of at least ten polypeptide bands ranging in molecular weight from 10,000 to 70,000 (89, 4.0). b. Activities. The physiological electron acceptor for complex I is ubiquinone-10. However, because of its water insolubility, the lower homologs, particularly ubiquinone-1 which is slightly water soluble, are more efficiently reduced by preparations of complex I. Indeed, special assay conditions are needed to demonstrate the in vitro reduction of added ubiquinone-10 or ubiquinone-6. Figure 2 shows a double reciprocal plot of data for the reduction of ubiquinone-6 by complex I and NADH. The VmaXis 21.5 pmoles NADH oxidized by ubiquinone-6 per min per mg protein, which is very close to the V,,, = 25 obtained with ubiquinone-1 (41). However, the apparent K , for QB (41a) is 0.63 mM,

+

38. S. Fleischer, H. Klouwen, and G . Brierley, JBC 238, 2936 (1961). 39. R. A. Capaldi, ABB 183,99 (1974). 40. Y. Hatefi and K. E. Stempel, JBC 244, 2350 (1969). 41. A. J. Merola, R. Coleman, and R. Hansen, BBA 73,638 (1963). 41a. Abbreviations: Ql and Qd, uniquinone-1 and ubiquinoneb, respectively ; APAD, acetylpyridine adenine dinucleotide; EPR, electron paramagnetic resonance;

4. METAL-CONTAINING

ACTIVITIESOF COMPLEX 1

Donor NADH NADH NADH NADH NADH NADH NADH NADH NADH NADH NADH NADPH NADPH NADH NADPH Succinate

181

FLAVOPROTEIN DEHYDROGENASES

WITH

TABLE I1 VARIOUS ELECTRON DONORSAND ACCEPTORS5

Acceptor Ferricy anide Ubiquinone-1 Ubiquinone-6 2,3-Dimethoxy-5,f5-dimethylbenzoquinone 2,3-Dimethoxy-5-methylbenzoquinone 2-Methylnaphthoquinone (menadione) 2,6-Dichloroindophenol Vitamin K I a-Tocopherylquinone Lipoic acid Cytochrome c Ubiquinone-1 Ferricy anide APAD APAD Ubiquinone-1, cytochrome c

Inhibition Specific by Amytal, activity* rotenone 685" 25e 21. 5c <1.5 <1.3 1.9 1.5 0.0 0.0 0.0 3-4 +d

0.9e

+ +

+ +

3.9' 0.3" 0.0

From Hatefi el al. (88, 40, 80). bSpecific activity is expressed as micromoles substrate oxidized per min per mg protein a t 38". 5 Vmax with respect to acceptor concentration. d The rate has not been reported. c Measured at p H 6.5.

whereas the K , for Q1is of the order of 40-50 p M (36,40).This difference probably reflects the water insolubility of Q6,thus resulting in an erroneously high K,. As in submitochondrial particles, the NADH-Q reductase activity of complex I is inhibited by Amytal, rotenone, piericidin A, or mercurials (28, 29, 41, 42). While Q1 is rapidly reduced by complex I, the replacement of its isoprenyl side chain with a methyl group or a proton results in substantial loss of reactivity (28).The small degree of reduction achieved with 2,3-dimethoxy-5,6-dimethylbenzoquinone (aurantiogliocladin) or 2,3-dimethoxy-5-methyl-benzoquinoneis not inhibited by Amytal and rotenone (Table 11). a-Tocopherylquinone is not NMR, nuclear magnetic resonance ; mV, millivolt ; pCMB, pchloromercuribenzoate ; pCMS, p-chloromercuriphenyl sulfonate ; PMS, phenazine methosulfate ; EDTA, ethylenediaminetetraacetate; SDS, sodium dodecyl sulfate ; TTFA, 2-thenoyltrifluoroacetone ; ETP, submitochondrial (electron transfer) particles; MVH, reduced methyl viologen ; DEAE-cellulose, diethylaminoethyl cellulose ; succ, succinate ; APS, adenosine 5'-phosphosulfate ; PAPS, 3'-phosphoadenosine 5'-phosphosulfate. Other abbreviations are standard [see JBC 244,2 (1969)1. 42. Y. Hatefi, K. E. Stempel, and W. G. Hanstein, JBC 244, 2358 (1969).

182

YOUSSEF HATEFI AND DIANA L. STIGGALL

ACTIVITIEBOF

THE

TABLE I11 NADH DEHYDROQENABE OF BAUQH AND KIN@

Acceptor Ferricyanide Juglone 2,3-Dicyano-5,6-dichloro-l14-benzoquinone Cytoohrome c Duroquinone Menadione 2,6-Dichloroindophenol Q6

K,(mM)

Specific activityb

5.0 0.3 0.7

1,700 64 58 0.16 3.8 2.9 2.4 16 37 59

-

1.5 0.07 0.2

-

-

Q1

4.3

Q1

From Baugh and King (43). Micromoles NADH oxidized x min-1 X mg-1 at 30' except for the Q homologs which were assayed at 22'. a

reduced by complex I, nor is the naphthoquinone vitamin K,.However, 2-methylnaphthoquinone (menadione, vitamin K,) is slowly reduced in an Amytal-rotenone insensitive manner. As will be seen below, the low molecular weight NADH dehydrogenase, which is derived from complex I after destabilization of complex I structure, has a very high rotenoneinsensitive menadione reductase activity. Therefore, the small activity seen in preparations of complex I might be due to the presence of small amounts (<2%,) of partially destabilized complex I. Among artificial electron acceptors only ferricyanide is rapidly reduced by complex I ; dichloroindophenol and methylene blue are poorly effective. As seen in Table 11, preparations of complex I are also capable of reducing cytochrome c a t a slow rate. This reaction is completely sensitive to inhibition by rotenone or antimycin A and probably results from the presence of traces of complex I11 in complex I preparations. Baugh and King (4.3,4%) have recently reported the isolation of a preparation from mitochondria with high NADH-ferricyanide and NADH-Q reductase activities (Table 111). As compared to complex I, the preparation contains 20-25% less FMN, but more iron and labile sulfide, the ratio of FMN :nonheme iron :labile sulfide being 1 :28 :28. The enzyme is isolated from Keilin-Hartree particles (prepared from beef heart mitochondria) after treatment with Triton X-100 and subsequently with cholate. It is claimed to be water soluble and free of phospholipids. HOWever, satisfactory analytical data for the absence of lipid (e.g., phos43. R. F. Baugh and T. E. King, BBRC,49, 1165 (1972). 43a. C. I. Ragan, W. R. Widger, and T. E. King, BBRC 80, 894 (1974).

4.

183

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES 1

I

1

I

I

1

5 -0.1

4 a -0.2 400 450 500 nm

550

FIG.3. Difference spectrum of NADH-reduced minus oxidized complex I at 5 mg protein per ml. Dotted line is the base line before addition of NADH to the sample cuvette. From Hatefi e t al. (88). phorus in ashed samples) and detergents (especially cholate which is less easily removable than Triton X-100) have not been provided. As seen in Table 111, the Km of the preparation of Baugh and King for ubiquinone-1 is 4.3 mM. This is two orders of magnitude greater than the K m of complex I. According to Ragan and Racker ( 3 6 ) ,it seems unlikely that the reactions catalyzed by the preparation of Baugh and King represent physiological events, because a t high ubiquinone-1 concentrations there is no phosphorylation a t site 1 linked to NADH oxidation (44). It has been shown by Ragan and Racker (36) that phospholipids are necessary for the ubiquinone, but not the ferricyanide, reductase activity of complex I. Thus, removal of about 50% of complex I lipids by extraction with cholate under special conditions resulted in a reversible loss of ubiquinone reductase activity without affecting the ferricyanide reductase activity. More phosphatidylcholine and phosphatidylethanolamine were removed by this procedure than cardiolipin. Readdition of either phosphatidylcholine or phosphatidylethanolamine restored considerable rotenone-sensitive ubiquinone-1 reductase activity, which was further augmented when small amounts of cardiolipin were also added. Using preparations depleted of ubiquinone-10 by pentane extraction, these authors have also shown that enzyme-bound ubiquinone-10 is not necessary for the reduction of added ubliquinone-1 by complex I or the inhibition of this reaction by rotenone. c. Spectral Properties. The absorption spectrum of NADH-reduced minus oxidized complex I is shown in Fig. 3. The bleaching afforded by NADH is the result of the reduction of flavin and the iron-sulfur chromophores, and the peak a t about 430 nm, which is superimposed on the flavin plus iron-sulfur bleaching, is the result of the Soret absorption of the reduced cytochromes of contaminating complex 111. Less than 50% 44. G . Schatz and E. Racker, JBC 241,1429 (1966).

184

YOUSSEF HATEFI AND DIANA L. STIGGALL

of the bleaching a t 450 nm could be attributed to flavin reduction (assuming full reduction by NADH). This discrepancy between the flavin content of complex I and the degree of bleaching afforded by NADH (or by succinate in the case of complex 11, see below) a t 450 nm had suggested that the reduction of nonheme iron is probably responsible for the additional bleaching (29, 46). Subsequent isolation and spectral studies of ferredoxins and demonstration of the presence of iron-sulfur species in complexes I, 11, and 111 substantiated these early predictions regarding the absorbancies of complexes I and I1 at about 450 nm not accounted for by flavin. The presence in complex I of NADH-induced, low-spin EPR signals was shown in early studies on this enzyme complex (28). These measurements were made a t near liquid nitrogen temperature, and the possible relevance of these signals to the mechanism of electron transfer from NADH to ubiquinone was recognized. Subsequently, the resolution of complex I by chaotropic agents into three distinct fractions, each containing iron-sulfur components (see below), suggested that complex I might contain more than one species of iron-sulfur moiety. Thus, E P R studies of NADH-treated complex I a t near liquid helium temperature indicated the presence of four iron-sulfur centers, which were designated centers 1, 2, 3, and 4 (34, 4 6 ) . These centers are identified in Fig. 4 with letters q, r, s (center 1); 0, p (center 2) ; and 1, m, n (overlapping centers 3 and 4). The field positions of the prominent peaks on the g scale are given in Table IV. As seen in A of Fig. 4, partial reduction of complex I with NADH resulted only in the appearance of centers 2 and 3. Further addition of NADH then produced centers 1 and 4 (B and C of Fig. 4). This is an expression of the reduction potential of these iron-sulfur tenters, which appears to be in the order 3 2 2 > > 4> 1 ( 4 6 ) . As seen in C of Fig. 4, the overlapping signals resulting from centers 3 and 4 (1, m, n) are emphasized a t high power and low temperature (7.7OK), whereas under these conditions center 2 (0,p) appears to be considerably saturated. Kinetic experiments with NADH or reduced acetylpyridine adenine dinucleotide (APAD), which reacts much more slowly than NADH, showed that the sequence of appearance of signals resulting from reduction of the iron-sulfur centers was 2, 3 4, 1 (34, 3 6 ) . All centers were reduced with NADH at 4 O within 6 msec ( 4 6 ) .Earlier studies of Beinert and his colleagues (47) had shown that the half-time of the appearance of

+

45. Y. Hatefi, “The Enzymes,”2nd ed., Vol. 7, p. 495, 1963. 46. N. R. OrmeJohnson, R. E. Hansen, and H. Beinert, JEC 249,19!2!2 (1974). 47. H. Beinert, G. Palmer, T. Cremona, and T. P. Singer, EERC 12, 432 (1983) ; JBC 240, 475 (1965).

4.

185

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

I

m n P

0

q

A

FIG. 4. E P R spectra of NADH-reduced complex I showing iron-surfur centers 1 (4, r, s), 2 ( 0 , p), and 3 + 4 (1, m, n). A, reduced with 10.6 electron neq of NADH; B and C, with 127 neq. Microwave frequency was approximately 9.2 GHz, power 03 mW, modulation amplitude, 7.5 G ; temperature 13°K for A and B, 7.7"K for C. From OrmeJohnson et al. (84). TABLE I V

FIELDP O S I T I O N S

A N D ABSIGNMENTS OF

OBSERVED IN COMPLEX I

AT

RESONANCES

13'Ka

Iron-sulfur center

Field positions*

g averagec

1 2 3 4

2.022, 1.938, 1.923 2.054, 1.922 2.100, 1.886, 1.862 2 . 103,d 1 .864d

1.96 1.97 1.95

From Orme-Johnson et al. (46).

* The numbers are the measured field positions of prominent peaks given on the g value scale. The average g values were calculated by assuming tha t, the values measured at the low and high field peaks correspond to g. and g., respectively, and by interpqlating or extrapolating a probable value of gv from the position of the center line or peak. d Since center 4 is only seen in the presence of center 3, i.e., the field position of the combined resonances is measured, these values may only be approximate. It is likely that the values differ somewhat more from those of center 3.

186

YOUSSEF HATEFI AND DIANA L. STIGGALL

TABLE V INTEGRATED INTENSITIES OF EPR RESONANCES FROM IRONSULFUR CENTERS OF COMPLEX I I N RELATION TO THE SPECTROPHOTOMETRICALLY DETERMINED FLAVIN CONTENT'

Iron-sulfur center

Ratio of concentration of iron-sulfur centers to flavin concentration

1+2+3+4 1 2 1+2 3 4 by difference

4.0 0.81 0.89 2.2 1.8

+

, From Orme-Johnson et al. (46)

the g = 1.94 signal (at 77OK, the temperature used for the EPR experiments, the signal mainly results from center 1) of NADH dehydrogenase preparations treated with slowly reacting NADH analogs corresponded to one catalytic cycle of the enzyme as measured by the reduction of ferricyanide. It is important to note that double integration of the signals of complex I have indicated that on the basis of electron consumption the molarity of each of the four iron-sulfur signals appears to be comparable to that of the flavin (Table V). Since the iron-sulfur centers appear to be of the ferredoxin type, this means that each center might involve 2 or 4 iron atoms (depending on whether they are plant type or clostridial type) and take up one electron. This is rather interesting in view of the fact that in complex I the ratio of flavin:iron:labile sulfide is very close to 1: 16: 16, which would agree with the possibility of four clusters of 4 irons and 4 labile sulfides each. Indeed, as will be seen below, the low molecular weight NADH dehydrogenase isolated from complex I has a composition of 1 flavin:4 iron:4 labile sulfide. According to Beinert and his colleagues (467, complex I takes up approximately 20 electron neq/mg protein, or 13-14 electrons per mole of flavin. The four iron-sulfur centers (assuming clostridial ferredoxin-type clusters of 4 iron and 4 labile sulfide per center) plus FMN would account for 6 electrons, and ubiquinone could account for another 5-6 (Table I).However, complex I and the high molecular weight NADH dehydrogenase also contain a multiplicity of thiol groups (see below) a t least one of which appears to become susceptible to inhibition by mercurials after treatment of the preparation with NADH (23,48). Assum48. D. D. Tyler, R. A. Butow, J. Gonze, and R. W. Estabrook, BBRC 10, 551 (1965).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

187

ing utilization of a pair of electrons in this activation process, then the titration data of Beinert and co-workers becomes remarkably accurate for complex I. DerVartanian et al. (49) have examined the E P R properties of the dehydrogenase of Baugh and King ( 4 3 ) ,which has a flavin:iron:labile sulfide ratio of 1:28:28. They have concluded that quantitation of the four reduced iron-sulfur centers by double integration accounts for only 36% of the iron content of the preparation. They felt that in this preparation the behavior of the EPR resonances suggests the presence of unidentified iron complexes in addition to iron-sulfur centers. Ohnishi et al. (50) have examined a preparation of complex I made by Ragan and Racker (36). According to the latter authors, their preparation has a ratio of flavin :iron: labile sulfide of 1 :23 :22 (the flavin content is 1.26 nmoles/mg protein). Ohnishi et a2. (50) found seven iron-sulfur centers in this preparation when examined a t different temperatures, high microwave powers, and in the presence of redox mediators ranging from -445 to -189 mV. Data regarding quantitation of these signals and their kinetic competence have not yet been presented. Such data are necessary in order to weigh the significance of the additional signals reported by Ohnishi et al. As stated above, iron-sulfur center 1 of complex I is relatively temperature-insensitive and observable a t liquid nitrogen temperature. This signal degenerates upon prolonged exposure of various preparations to NADH and leads to the appearance of new signals (46,51).The interpretation that these latter signals resulted from the presence of molybdenum in complex I (56) has been abandoned ( 5 3 ) . Preparations of complex I or mitochondria1 inner membrane fragments do not contain more than trace amounts of molybdenum (0.03 atom/mole of acid-extractable flavin) (34, 5 3 ) . These and other complications involved in the interpretation of E P R data have been discussed by Beinert and his colleagues (46,54) as well as by Albracht ( 5 5 ) . 2. High Molecular Weight N A D H Dehydrogenases In 1962, Ringler et al. (see 56) reported the isolation of a NADH dehydrogenase from bovine heart mitochondria with the use of Naja naja venom phospholipase A and Triton X-100. The preparation contained 0.9 nmole 49. D. DerVartanian, R. F. Baugh, and T. E. King, BBRC 50, 629 (1973). 50. T. Ohnishi, J. S. Leigh, C. I. Ragan, and E. Racker, BBRC 56, 775 (1974). 51. M. Kawakita and Y. Ogura, J . Biochem. (Tokyo) 66,203 (1969). 52. S. P. J. Albracht and E. C. Slater, BBA 223,457 (1970). 53. S. P. J. Albracht, H. VanHeerikhuiren, and E. C. Slater, BBA 2 5 6 , l (1972). 54. N. R. OrmeJohnson, R. E. Hansen, and H. Beinert, JBC 249, 1928 (1974). 55. S. P. J. Albracht, BBA 347, 183 (1974). 56. R. L. Ringler, S. Minakami, and T. P. Singer, JBC 238, 801 (1963).

188

YOUSSEF HATEFI AND DIANA L. STIGGALL

of flavin per mg protein, 16 g-atoms of iron per mole of flavin, and 6.2% lipid by dry weight. It catalyzed the reduction of ferricyanide by NADH with a calculated turnover number of 6.6 x lo6 moles of NADH oxidized per minute per mole of “flavoprotein” a t 38O. The flavin was a mixture of 25-30% FAD and the rest F M N or FMN plus riboflavin. The enzyme also contained 5’-AMP in amounts roughly equivalent to the sum of FMN plus riboflavin. These results were somewhat reminiscent of the split products of FAD in the NADH-cytochrome c reductase preparation of Mahler et al. (57). Thus, the authors considered that the ffavin moiety of the enzyme might be FAD or both FAD and FMN (56).The identity of the flavin of mitochondria1 NADH dehydrogenase was settled by Rao et al. (37)in 1963 by a careful analysis of the total and acidextractable flavin of several preparations from mitochondria, especially complex I and its parent particle complex 1-111. They found that in these preparations more than 96% of the flavin was acid-extractable FMN. Subsequently, Cremona and Kearney (68) modified the preparation of the dehydrogenase of Ringler et al. (66),deleted the use of Triton X-100, and obtained a more purified preparation of the enzyme containing 1.232 0.02 nmoles of flavin per mg protein. The flavin was identified as , ~the ) preparation a t pH 10 FMN. The sedimentation coefficient ( s ~ ~ of (to prevent aggregation) was estimated to be 14 2 0.5 in the concentration range of 6-10 mg/ml. However, a skewing of the sedimentation boundary was observed, which was stated to result from 30-3576 colorless impurity (actual results not shown). On the basis of its flavin content, the preparation would have a minimum molecular weight of 813,000per mole of flavin. However, the authors corrected for the presumed 3 M 5 % impurity calculated from their sedimentation patterns a t pH 10, and concluded that the molecular weight of the “pure flavoprotein” is of the order of 550,000.Subsequent pubhations from Singer’s laboratory have used this figure as the established molecular weight of the high molecular weight NADH dehydrogenase (19, 23). Using a similar correction as above, the turnover number of the dehydrogenase preparation has been calculated to be 800,000 per minute at 30°. This and the earlier preparation of Ringler et al. are claimed to be water-soluble, even though both were isolated after phospholipase treatment of particles, which results in the formation of detergent-like lysolipids, and Triton X-100 was added to the preparation of Ringler et al. to prevent aggregation. The NADH dehydrogenase preparation of Baugh and King (43) is also stated to be water-soluble. Both Triton X-100 and 57. H. R. Mahler, N. K. Sarkar, L. P. Vernon, and R. A. Alberty, JBC 199, 685 (1052). 58. T.Cremona and E. B. Kearney, JBC 239,2328 (1964).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

189

cholate were used during its isolation, but it is claimed that addition of cholate resulted in the removal of Triton X-100 which was added earlier, and that subsequent chromatography on agarose and sucrose gradient centrifugation removed the added cholate. Huang and Pharo (69) have isolated a NADH dehydrogenase with the use of Lubrol, which is very similar in enzymic properties, absorption spectrum, and flavin content (1.17 nmole/mg protein) to the preparation of Cremona and Kearney. They stated, however, that detergent appears to be essential for the solubility of the dehydrogenase since its removal resulted in an insoluble, but active, preparation. Thus, it seems prudent to reserve judgment on whether or not these preparations of high molecular weight NADH dehydrogenase are truly water-soluble. By comparison to complex I, they ought to contain a considerable amount of hydrophobic membrane “proteins” or polypeptides, and the complete absence of lysolipids in the preparation of Cremona and Kearney (58) and detergents in the preparations of Ringler et al. (56) and Baugh and King (43) has not been demonstrated. On the other hand, while water solubility of a complex enzyme is convenient for laboratory experiments, it may be of little physiological significance when in the native state the complex is tightly membrane bound. Table VI summarizes the composition and activities of various preparations of high molecular weight NADH dehydrogenase and provides a comparison with complex I. 3. Low Molecular Weight NADH Dehydrogenases

The low molecular weight form of mitochondrial NADH dehydrogenase was first isolated from pig heart muscle by Edelhoch et al. (60) and Mahler and his associates (57) in 1952. The mitochondrial origin of the enzyme was demonstrated by de Bernard (61).These and similar preparations reported subsequently by Mackler (62), Kumar et al. (651, and Pharo et al. (64) were isolated by extracting the source material (heart muscle or various submitochondrial preparations) with 9-1 1 % ethanol at pH 4.8-5.3 and 430450, a procedure originally devised for isolation of the Straub diaphorase (lipoyl dehydrogenase) ( 6 5 ) . TWO other preparations of basically similar composition and catalytic properP. C. Huang and R. L. Pharo, BBA 245, 240 (1971). H. Edelhoch, 0. Hayaishi, and L. J. Tepley, JBC 197, 97 (1952). B. de Bernard, BBA 23, 510 (1957). B. Mackler, BBA 50, 141 (1961). S. A. Kumar, N. A. Rao, S. P. Felton, F. M. Huennekens, and B. Mackler, ABB 125, 436 (1968). 64. R. L. Pharo, L. A. Sordahl, S. R. Vyas, and D. R. Sanadi, JBC 241, 4771 59. 60. 61. 62. 63.

(1966). 65. F. B. Straub, BJ 33, 787 (1939).

TABLE VI COMPOSITION AND PROPERTIES OF HIGHMOLECULAR WEIGHT NADH DEHYDROGENASES

Flavin (nmole/mg protein)

Flavin :iron : sulfide

1.4-1.5

1:16-18: 16-18

Ringler et al. (1962) Cremons-Kearney (1964)

0.9 1.23

1:16:? 1: 17-18:27-28

HUmg-Phmo (1971) Baugh-King (1972)

1.17 1.13

1:26:? 1:28: 28

0.5-0.6

1:28:28

Preparation Complex I (1961)

Tottmar-Ragan (1971) (from C. utilis)

Reactions catalyzed NADH + Q, K*Fe(CN),, APAD NADPH + Q, KIFe(CN)6JNAD NADH -+KIFe(CN), NADH, NADPH -+ K*Fe(CN)* NADH -+ APAD NADH + KIFe(CN), NADH + Q, &Fe(CN),, APAD NADPH + K,Fe(CN),, NAD NADH, NADPH + KIFe(CN)s

KZADH K$ (rM) (PM) 7

44

108

References 27J28

4 0

66

u,

68,83,84 69

83

4,300

43,43a 133

s

4 xgI3 M 9

Ei 3z P

r u,

Er

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

191

Temperature, *C

FIO.5. Kinetics of the resolution of complex I as a function of temperature of the incubation medium. The release of menadione reductase activity was measured as an index of resolution 2.5 min after incubation of complex I with 0.47 M NaClO, at the temperatures indicated. From Davis and Hatefi (69).

ties are those of King and Howard (66) and Hatefi and Stempel (40, 67, 68). The former was extracted from Keilin-Hartree particles from beef heart after incubation of the particles at 37O with boiled snake venom (as a source of phospholipase) in the presence of CaC1,. The latter was isolated after resolution of complex I with various chaotropic agents (67, 69). The above dehydrogenase preparations are qualitatively similar in composition and enzymic activity and have molecular weights (often calculated from flavin content) between 7 and 12 X lo4. They contain FMN, nonheme iron, and labile sulfide (wherever examined), and have a wide electron acceptor specificity with respect to quinoid structures and ferric compounds. They are all inhibited by mercurials. The quantitative differences in composition and activity appear to be related to their isolation and purification conditions. Among these dehydrogenases, the preparation of Hatefi and Stempel (40, 67, 68) has been more fully studied and appears to have been obtained with the least damage since it exhibits the highest enzymic activities and the highest content of flavin, iron, and labile sulfide. Therefore, the properties of this preparation as an example of the low molecular weight NADH dehydrogenases will be more thoroughly discussed. The enzyme is isolated from complex I after resolution of the complex with chaotropic agents. The resolution process is highly temperaturedependent (Fig. 5 ; activation enthalpy from data of Fig. 5 AH' = $37 66. T.E. King and R. L. Howard, JBC 237, 1686 (1962). 67. Y. Hatefi and K. E. Stempel, BBRC 26,301 (1967). 68. Y. Hatefi, K. E. Stempel, and W. G . Hanstein, JBC 244, 2358 (1969). 69. K. A. Davis and Y. Hatefi, Biochemistry 8,3355 (1969).

192

YOUSSEF HATEFI AND DIANA L. STIGGALL

0.8

0.6 k 0.4

0.2

Salt concentration (M)

FIQ.6. Effect of various chaotropes on the first-order rate constant (k) of the resolution of complex 1. TBA, tribromoacetate; TCA, trichloroacetate; DCA, dichloroacetate ; TFA, trifluoroacetate; MCA, monochloroacetate; and AcO; acetate. From Hanstein et at. (71).

kcal/mole), irreversible, and retarded in the presence of NADH. Its rate is finely controllable by the concentration and the potency of the chaotropic agent employed (Fig. 6), and strongly inhibited when the medium H,O is replaced with the more structured solvent, D,O (Fig. 7 ) . The sig-

h C Q . [MI

FIG.7. Solvent isotope effect on the resolution of complex I by NaClO, at 20" and 30"; k, first-order rate constant in min-'. From Hanstein et al. (73).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

193

Fxa. 8. Absorption spectrum of the soluble iron-sulfur protein (4.3 mg/ml) isolated from complex I. Dashed line, after treatment with dithionite; dotted line, after treatment with sodium mersalyl to destroy the iron-sulfur chromophore. From Hatefi et al. (3.9).

nificance of these results in relation to the effect of water structure on the structural stability of complex I in aqueous media has been discussed elsewhere (6‘9-73). The resolved complex is composed of two fractions, a soluble part, which comprises about 15% of complex I proteins, and a water-insoluble part consisting of the rest of the protein and the bulk of complex I lipids. The soluble fraction is easily separated from the inscluble material by centrifugation. Upon fractionation with ammonium sulfate, it yields a soluble flavoprotein containing iron and labile sulfide and a dark brown protein, which contains large amounts of iron and labile sulfide but no flavin. The latter appears t o be an iron-sulfur protein and exhibits an EPR signal which is characteristic of iron-sulfur center 2 of intact complex I (46‘). Its absorption spectrum is shown in Fig. 8. The insoluble fraction also contains equimolar amounts of iron and labile sulfide and little or 110 flavin. The flavoprotein fraction represents the low molecular weight NADH dehydrogenase and contains per mg protein 13.5-14.5 nmoles of F M N (acid extractable), 60-65 ng-atoms of iron, and 58-60 nmoles of acid 70. Y. Hatefi and W. G. Hanstein, Proc. N a t . Acad. Sci. US. 62, 1129 (1969). 71. W. G. Hanstein, K . A. Davis, and Y. Hatefi, ABB 147, 534 (1971). 72. Y. Hatefi and W. G. Hanstein, “Methods in Enzymology,” Vol. 31, Part A, p. 770, 1974. 73. W. G. Hanstein, K. A. Davis, and Y. Hatefi, ABB 163, 482 (1974).

194

YOUSSEF HATEFI AND DIANA L. STIGGALL

labile sulfide. The ratio of flavin to iron to labile sulfide is, therefore, close to 1:4:4, suggesting that iron and labile sulfide might be in a clostridial ferredoxin-type cluster. The visible spectrum of the oxidized dehydrogenase analyzed for contributions of flavin and iron-sulfur chromophore, plus the spectra of NADH- and dithionite-reduced enzyme are shown in Fig. 9. Its enzymic properties and kinetic constants with respect to various electron acceptors are given in Table VII. Comparative data for complex I are provided in Table VIII. I n addition to ubiquinone-1, the enzyme also reduces higher isoprenologs of ubiquinone a t appreciable rates (40,67). The dehydrogenase preparations obtained by acid-ethanol extraction of particles a t elevated temperatures vary considerably in their content of flavin, iron, and labile sulfide, and in their activities. These differences appear to be largely a consequence of destruction of the iron-sulfur chromophore a t acid pH. As seen in Fig. 10, incubation of the low molecular weight dehydrogenase preparation of Hatefi and Stempel a t pH 4.8 and 3 8 O resulted after 1 hr in nearly complete loss of labile sulfide (Fig. 1OC) and reductase activity with respect to menadione, cytochrome c,

I---

FIa. 9. Spectral characteristics of the soluble NADH dehydrogenaee (1.6 mg/ml) isolated from complex I. Traces 1, spectrum of oxidized enzyme; 2, NADH-reduced enzyme; 5, dithionite-reduced enzyme; 4, flavin contribution to 1 after destruction of iron-sulfur chromophore with sodium mersalyl ; 3, iron-sulfur contribution to.1 obtained by subtraction of 4 from 1; 6, 4 plus dithionite showing that after destruction of the iron-sulfur chromophore with mersalyl and reduction of flavin with dithionite the enzyme has no absorption in the visible region. From Hatefi and Stempel (40).

4.

196

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES k

100.

,

,

,

,

,

,

- -- ------_-

I

M 4020

-

,

pHs8.0 "

1

I

1

1

102030405060 min at 38'

102030405060 min at 38'

(b)

(a)

10 2 0 3 0 4 0 3 3 6 0 min at 38' (C)

FIQ. 10. Effect of incubation at pH 4 8 and 38" on the activities [(a) pH 8.0, (b) pH 4.81 and labile sulfide content (c) of NADH dehydrogenase.Ks, menadione. ' Key: (-) Ks, (---) cytochrome c, and (---) ferricyanide. From Hatefi (76).

and ferricyanide (Fig. 10B). By comparison, a sample of the enzyme incubated a t pH 8.0 lost less than 20% labile sulfide (Fig. lOC), 40% menadione and ferricyanide reductase activity, and very little cytochrome c reductase activity (Fig. 10A). Kumar et al. (63) have estimated the molecular weight of the acidethanol-extracted dehydrogenase by Sephadex gel exclusion to be approximately 70,000. Assuming one mole of FMN per mole of enzyme, the molecular weight of 70,000 agrees with the flavin content of the preparations of Kumar et al. (6S),Pharo et al. ( 6 4 ) , and Hatefi and Stempel (40).Kumar et al. (63) have also determined the amino acid composition of their preparation, except that the value for tryptophan is not

TABLE VII ENZYMIC PROPERTIES OF SOLUBLE NADH DEHYDROGENASE'

Reaction NADH + KIFe(CN)* NADH + cytochrome c NADH + &I NADH --+ menadione NADH + 2,6-dichloroindophenol Reduced APAD + &I

Specific activitp 215 43 150-160 160-170 100

VEDfeO 330 76 250 330 125 3.8

cxps=

400" 220 175 190

K:"" (rM) 65 64 133 133 133 715

Inhibition by KFPtor 1 pM rotenone (PM) (%I 1650' 600 167 160 62

None 10-20 10

Inhibition by >0.25 mM NADH

-

+ + +

From Hatefi and Stempel (40). b Expressed as micromoles of NADH oxidized per min per mg of protein at 38". All activities are completely inhibited with 0.1 mM pmercuriphenyl sulfonate. At 0.75 mM NADH.

3

srn

M I %

3:

2I 3

*

z E

9

r rn

Br

F

tp

E

9

t: 0 0

i5!

TABLE VIII NADH DEHYDROGENASE PROPERTIES OF COMPLEXES I A N D 1-111.

2 ci

Inhibition by 0.5 mM

Reaction

Enzyme complex

Specific activity

NADH -+ K3Fe(CN)6 NADH 4 cytochrome c NADH -+ cytochrome c NADH 4 &I NADH + menadione NADH -+ 2,6-dichloroindophenol

I I 1-111 1-111 I I

lo@ 2d 25-30 14 1.9 1.5

K:*”” (pM)

7 14 14 15-17

KrPto.

pCMS

1 pM

I

c 0 w

1PM

rotenone antimycin A

(pM)

(%I

(%I

(%)

NADH

400W

None

None

None

>O.lmM

12

100 100 100

100 100 100

100 100

None None

None None

12 44

None None None

>0.25mM

From Hatefi and Stempel (40). Per mole of flavin, this activity is considerably higher in complex I than in the soluble, low molecular weight dehydrogenase. c A t 0.15 mM NADH; V ~ ~ ~ ( C= N685. ’6 d This activity results from the presence in complex I of 0 . 5 1 % complex I11 contamination.

0

Y

9 z

tl

ii ;s

i% e3

198

YOUSSEF HATEFI AND DIANA L. STIGGALL

given. According to these investigators, a considerable amount of flavin can be removed from the enzyme by treatment with Florisil or Bio-Gel. The depleted enzyme retains its ferricyanide reductase activity, but loses considerable activity for reduction of dichloroindophenol and cytochrome c. The latter is partially restored by addition of large amounts of FMN. The authors concluded from these data that reducing equivalents from NADH first go to the iron-sulfur moiety of the enzyme, then to flavin. Ferricyanide accepts electrons from the iron-sulfur moiety, but indophenol, quinones, and cytochrome c are reduced a t the flavin site. I n agreement with this conclusion they have shown that chromatography of the enzyme on DEAE-cellulose at pH 6.8 results in nearly complete removal of flavin and labile sulfide, and about two-thirds of the iron. This preparation had no reductase activity with any of the acceptors, even when assayed in the presence of added FMN. The above mechanistic conclusions are not generally accepted, however, because (a) the preparation of Kumar et al. has very low reductase activities (probably related to its low content of iron and labile sulfide) , (b) the cytochrome c reductase activity restored by addition of FMN is only about 1% of the maximal cytochrome c reductase activity of the more active preparations of the enzyme (do),and ( c ) no attempt was made to reconstitute the ironsulfur moiety of the DEAE-cellulose-treated enzyme by treatment with sulfide and ferrous ions to see whether ferricyanide reductase activity can be restored. It has been stated by Yang (74)that the partial loss of ferricyanide reductase activity (and 450 nm absorption) of the preparation of Kumar et al. upon aging in air a t room temperature could be restored to a considerable extent by treatment of the enzyme with 2mercaptoethanol, FeCl,, and Na,S followed by filtration through a column of Sephadex G-25. However, the validity of this type of reconstitution experiment rests on several important controls which were not presented. 4. Relevance of the Low and High Molecular Weight Preparations

to Mitochondria1 NADH Dehydrogenase The ubiquinone reductase activity of their low molecular weight dehydrogenase led Pharo et al. (64, 75) to conclude that the enzyme represented the mitochondria1 NADH-ubiquinone reductase. However, it has been shown that the quinone reductase activity of the low molecular weight dehydrogenase is different from that of intact respiratory particles or complex I in many important respects, including kinetic constants, re74. C. S. Yang, in “Flavins and Flavoproteins,” 3rd Int. Symp. (H. Kamin, ed.), p. 664. Univ. Park Press, Baltimore, Maryland, 1971. 75. D. R. Sanadi, R. L. Pharo, and L. A. Sordahl, in “Non-Heme Iron Proteins” (A. San Pietro, ed.), p. 429. Antioch Press, Yellow Springs, Ohio, 1966.

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

199

mM DPNH

FIG. 11. Effect of NADH concentration on the ferricyanide reductase activities of complex I and the soluble, low molecular weight dehydrogenase. From Hatefi and Stempel (40).

sponse to inhibition by Amytal, rotenone, and piericidin A, and the apparent involvement of several EPR-active iron-sulfur centers. As compared to respiratory particles or complex I, the soluble enzyme exhibits low ferricyanide reductase activity per mole of flavin and very high reductase activities with respect to menadione, 2,6-dichloroindophenol, or cytochrome c as electron acceptor. The latter activity, in contrast to that found in submitochondrial particles or complex 1-111, is insensitive to inhibition by rotenone, piericidin A, or antimycin A, and 4 s marked by a very high K , for cytochrome c (600 p.M versus 12 p M in the case of complex 1-111) (Tables VII and VIII; also see 40). In addition, the ferricyanide reductase activity of complex I is sharply inhibited at NADH concentrations above 0.1 mM whereas the activity of the soluble dehydrogenase is not (Fig. 11). Conversely, the former activity is not inhibited by mercurials, but the latter is. It has also been shown that the ubiquinone reductase activity of particles is 70% inhibited by 50 m M guanidineHCl, whereas the same activity catalyzed by the low molecular weight dehydrogenase is 75% activated (68, 76). The cytochrome c reductase activity of the soluble enzyme was first discovered by Mahler and co-workers (5‘7); hence, the designation “Mahler’s DPNH-cytochrome c reductase.” However, these investigators had recognized the unphysiological nature of this activity, and Mahler and Glenn (7’7) pointed 76. Y. Hatefi, PTOC.Nat. Acad. Sci. U . S. 60,733 (1968). 77. H. R. Mahler and J. L. Glenn, in “Inorganic Nitrogen Metabolism” (W. D. McElroy and B. Glass, eds.), p. 575. Johns Hopkins Press, Baltimore, Maryland, 1956.

200

YOUSSEF HATEFI AND DIANA L. STIGGALL

out that the cytochrome c reductase activity of their dehydrogenase might be because cytochrome c behaves as a single electron acceptor similar to, but not identical with, the physiological electron acceptor in the respiratory chain. This early prediction is interesting since a single electron accepting iron-sulfur structure is very likely the natural acceptor for the membrane-bound dehydrogenase. It should also be mentioned that interaction with cytochrome c is not a peculiar property of the above enzyme. A number of flavoproteins containing FMN or FAD as prosthetic group and utilizing NADH or NADPH as electron donor are known which interact with cytochrome c (29, 46). Examples of this unphysiological phenomenon are in the case of Old Yellow Enzyme (78) and Straub’s diaphorase (lipoyl dehydrogenase) (79). It is clear from the above section that the differences between the enzymic properties of the soluble, low molecular weight NADH dehydrogenase and the particulate system represented by complex I are very large, even though the former is obviously a component of the latter enzyme system. Singer and his colleagues believe that the low molecular weight enzyme is a “peptide fragment” of the high molecular weight NADH dehydrogenase (16, 19). While in essence the parent-progeny relationship is obvious, the vigorously espoused views concerning peptide fragment and the equivalence of the high molecular weight preparation to mitochondrial NADH dehydrogenase are no longer acceptable. It has been pointed out that the small molecular weight dehydrogenase is not likely to be a peptide fragment because the procedures leading to its isolation are not likely to cleave peptide bonds (69, 80). Furthermore, as will be seen below, complex I and the corresponding section of respiratory particles catalyze the dehydrogenation of NADPH without the intermediation of NAD and the transhydrogenase reaction. Studies on complcx I with NADH and NADPH as substrates have shown that flavin and iron-sulfur center 1 are reduced by NADH, but apparently not by NADPH. Therefore, there appears to exist in complex I a segment, containing flavin and a portion of the total iron and labile sulfide, which is specific for dehydrogenation of NADH. It is highly probable that the small molecular weight NADH dehydrogenase represents this segment of complex I, except that conversion from membrane-bound to soluble state has modified its enzymic properties, some of which (e.g., wide acceptor specificity) might be simply the result of better access of acceptors to the soluble enzyme. Much has been written by Singer and his colleagucs in defense of thc 78. A. Akeaon, A. Ehrenberg, and H. Theorell, “The Enzymes,” 2nd ed., Voi. 7, p. 477,1963.

79. V. Massey, BBA 30, 205 (1958). 80. Y. Hate6 and W. G. Hanstein, Biochemistry 12,3515 (1973).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

201

thesis that their preparation is the true NADH dehydrogenase of the respiratory chain (14, 15, 19, 2 3 ) . They have done extensive studies on their preparation and have compared its composition, enzymic, and E P R properties to complex I and the low molecular weight dehydrogenase preparations. They have pointed out quite correctly that theirs is the first preparation of a NADH dehydrogenase which displays a high ferricyanide reductase activity comparable on the basis of NADH dehydrogenase flavin to the activity of complex I and intact respiratory chain preparations. By comparison, the low molecular weight NADH dehydrogenases have very low ferricyanide reductase activity. However, as compared to complex I, the dehydrogenase of Singer and co-workers appears to have several important differences. The comparison to complex I as a reliable point of reference is valid since (a) the above dehydrogenase resembles complex T in its size, ferricyanide reductase activity, and content of nonheme iron, and ( b ) complex I appears to be the smallest segment yet isolated which displays all the catalytic and inhibitor-response properties of the NADH-ubiquinone reductase portion of intact respiratory chains, including the important ability of conserving oxidative energy and reacting with mitochondria1 coupling factors to synthesize ATP ( 3 5 ) .The differences are as follows: a. The NADH dehydrogenase of Singer and co-workers is incapable of reducing ubiquinone, which is the physiological electron acceptor for complex I. This inability has been referred to the fact that the dehydrogenase is essentially devoid of lipids. That phospholipids are necessary for ubiquinone reduction by complex I has been demonstrated by Ragan and Racker (36 ) as discussed above. However, the latter investigators were able to restore ubiquinone reductase activity by adding phospholipids to deficient complex I preparations. I n the absence of such activity restoration experiments, the view that the preparation of Singer e t al. has not sustained irreversible damage during isolation would remain an assumption. According to Ragan and Racker ( 3 6 ) ,special reducing conditions are needed during the removal of lipids from complex I in order to preserve the ability of the complex to exhibit ubiquinone reductasc activity upon readdition of lipids. It remains to be seen whether the dehydrogenase of Singer et al. can be isolated under similar reducing conditions with preservation of ubiquinone reductase activity when assayed in the presence of added lipids. b. As stated above the ratio of flavin to iron to labile sulfide is 1:16-18:16-18 for complex I. This ratio is stated to be 1:17-18:27-28 for the NADH dehydrogenase. The molar excess of labile sulfide as compared to iron is surprising and contrary to literature data for all species of iron-sulfur proteins known. However, this high labile sulfide value

202

YOUSSEF HATEFI AND DIANA L. STIGGALL

might have resulted from the low extinction coefficient (21,000 liters mole-' cm-l) used by the authors (81) for labile sulfide determination by the method of Fog0 and Popowsky (82). A more correct molar extinction coefficient is between 27,500 and 30,000, which-when applied to the labile sulfide value published for NADH dehydrogenase-would lower it to about 20, a value in much better agreement with the iron content of the preparation. c. According to Watari et al. (83) and Singer and Cremona ( I d ) , the K , for NADH of their preparation is 108 p M . This value is more than 15-fold greater than the K , of complex I for NADH (7 p i l l ) determined similarly in the NADH-ferricyanide reductase assays (Table VIII) . This difference is rather serious because the high K,,, value is characteristic of the low molecular weight NADH dehydrogenase derived from complex I. The K , for NADH of the low molecular weight enzyme, also determined in the ferricyanide reductase assay, is about 65 (Table VII), and as detailed above it is generally agreed that the isolated low molecular weight dehydrogenase shows major differences in catalytic properties as compared to its membrane-bound counterpart. Thus, with respect to its K,,, for substrate, the NADH dehydrogenase of Singer and co-workers is similar to the modified, low molecular weight enzyme, and differs from complex I and other submitochondrial particles. This difference might be associated with structural modifications responsible for the inability of the high molecular weight NADH dehydrogenase to interact with ubiquinone under appropriate conditions. Although it was not recognized as a reaction involving a separate mechanism, the published data of Singer's laboratory show clearly that their preparation also has NADPH dehydrogenase activity (84). Consequently, the high molecular weight NADH dehydrogenase preparations appear to be segments of the respiratory chain related to complex I. These preparations appear to have preserved the ferricyanide reductase activity of the system but irreversibly lost the physiological ubiquinone reductase activity. The argument as to which preparation-the high or the low molecular weight enzyme-represents the respiratory chain NADH dehydrogenase is perhaps irrelevant in view of our present knowledge. Both are clearly derived from complex I. However, the low molecular weight preparation has grossly modified enzymic properties (see also Section II,B), and the high molecular weight preparation appears to have retained the NADH and NADPH dehydrogenase activities of com81. 82. 83. 84.

C. J. Lusty, J. M. Machinist, and T. P. Singer, JBC 240, 1804 (1965). J. K. Fog0 and M. Popowsky, Anal. Chem. 21,732 (1949). H. Watari, E. B. Kearney, and T. P. Singer, JBC 238, 4063 (1963). C. Rossi, T. Cremona, J. M. Machinist, and T. P. Singer, JBC 240, 2634 (1965).

4.

METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

203

plex I, but only with respect to ferricyanide as electron acceptor. These considerations lead, therefore, to the conclusion that proper purification of NADH dehydrogenase beyond the stage of complex I has yet to be achieved.

5. Inhibitors of N A D H Dehydrogenase Thiol reagents, barbiturates, rotenoids, and piericidin A inhibit NADH dehydrogenation and ubiquinone reduction in appropriate preparations. According to Singer and his colleagues (19,23, 85-87) there are five types of --SH groups in various preparations of NADH dehydrogenase. Type I is the -SH group found in small molecular weight NADH dehydrogenase, and is apparently involved in mercurial inhibition of its reductase activities. According to Kumar et al. ( 6 3 ) , NADH treatment increases the mercurial sensitivity of the enzyme with respect to ferricyanide and cytochrome c, but not dichloroindophenol, reduction. Hatefi et al. (68) have shown that (a) incubation of the enzymes with 1-2 pM pCMS results in activation by as much as 20-300/0, while higher concentrations inhibit, and (b) contrary to thc results of others (37) N-ethylmaleimide does not cause inhibition even after partial inactivation by heat a t pH 4.8 or in the presence of 3 M urea. In contrast to the low molecular weight enzyme, the ferricyanide reductase activity of the high molecular weight dehydrogenase or complex I is not inhibited by mercurials, suggesting that type I -SH groups are not accessible to mercurials in these preparations. The type I1 -SH group appears to be peculiar to the high molecular weight dehydrogenase of Singer et al. At Oo it reacts rapidly and reversibly with -SH reagents without inhibiting ferricyanide reductase activity, but incubation a t 15O-3Oo results in gradual and irreversible inactivation. The temperature effect suggests structural destabilization and recalls the temperature dependence of the resolution of complex I by chaotropic agents. Davis and Hatefi (69) have shown that in the presence of moderate concentrations of NaC104 the resolution of complex I does not occur at temperatures below 15O (Fig. 5 ) . Type I11 -SH groups are found in particles and high molecular weight preparations. This type of -SH group, which was discovered by Tyler et al. (48), reacts with mercurials and results in inhibition of electron transport only after the preparation is pretreated with NADH. The conditioning by NADH is reversible inasmuch as addition of ferricyanide to the NADH-treated enzyme reverts it to the mercurial insensitive state. This type of -SH 85. T. Cremona and E. B. Kearney, JBC 240,3645 (1965). 86. H. Mersmann, J. Luthy, and T. P. Singer, RBRC 25,43 (1966). 87. M. Gutman, H. Mersmnnn, J . Luthy, and T. P. Singer, Biochemistry 9, 2678 (1970).

204

YOUSSEF H A T E F I AND DIANA L. STIGGALL

group is considered to be located very near the substrate binding site, but possibly not directly involved in electron transport activity, since relative to the turnover number of the enzyme both the NADH conditioning and the ferricyanide reversal are slow processes (23).Type IV -SH groups are also found in particles and high molecular weight preparations. According to Singer and co-workers (19, 23), they react readily with low levels of mercurials, and mercaptide formation with this type of thiol group is considered to affect the ferricyanide reductase reaction by increasing both the K,,, and the V,,, for ferricyanide by severalfold. Type V -SH groups are detectable only in complex I and parent particles because they affect ubiquinone reduction and piericidin binding. Mercaptide formation with this type of -SH group requires relatively high concentrations (30-80 p M ) of mercurials and results in inhibition of electron transport from NADH to ubiquinone, but not to ferricyanide and other acceptors reacting on the substrate side of ubiquinone (Table VIII) . I n submitochondrial particles, mercaptide formation with type V -SH groups also results in the loss of one of two specific binding sites for rotenone and piericidin A, and a sigmoidal to hyperbolic change in the piericidin inhibition titration curves (23, 88). The ubiquinone reductase activity of complex I is inhibited by barbiturates (e.g., Amytal and Seconal), Demerol, rotenone, or piericidin A. These compounds appear to inhibit electron transfer from the iron-sulfur centers of complex I to ubiquinone. Absorption and fluorescence spectroscopic studies on submitochondrial particles had suggested to Chance et al. (89) the existence of two consecutive flavoproteins between NADH and ubiquinone. These authors placed the site of rotenone and Amytal inhibition between the two flavoproteins. Hatefi (76) showed that in complex I there is only one type of flavoprotein, and that the additional bleaching by substrate a t the wavelength pair 475 minus 510 nm used by Chance et al. results from reduction of the iron-sulfur components of complex I. Therefore, the site of Amytal and rotenone inhibition could be between the flavoprotein and an iron-sulfur moiety of complex I. While the conclusion regarding the absence of two consecutive flavoproteins in the complex I region of the respiratory chain was correct and confirmed (go), the interpretation regarding the site of inhibition of rotenone and Amytal was not. It is now generally agreed that the flavin and all the EPR-active iron-sulfur moieties of complex I are located on the substrate side of the inhibition site of Amytal, rotenone, and piericidin A. 88. M. Gutman, T. P. Singer, and J. E. Casida, JBC 245, 1992 (1970). 89. B. Chance, L. Ernster, P. B. Garland, C.-P. Lee, P. A. Light, T. Ohnishi, C. I. Ragan, and D. Wong, Proc. N u t . Acad. Sci. U.S. 57, 1498 (1967). 90. C. I. Ragan and P. B. Garland, BJ 10, 399 (1969).

4. METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

205

Palmer et al. (91) have suggested that in addition to the above site, rotenone and piericidin A also inhibit electron transport immediately on the substrate side of cytochrome cl. This view has not been accepted by others. Teeter et al. (9.2) have shown that secondary effects of rotenone and piericidin can be observed a t other regions of the respiratory chain when high concentrations of the inhibitors are used, as by necessity did Palmer et al. in their EPR experiments. The studies of Horgan, Singer, and their colleagues (19, 22, 23, 88, 93, 94) with radioactive piericidin A and rotenone have led these authors to the following conclusions :

(a) There are specific and unspecific binding sites for both rotenone and piericidin A, the latter being reversible by washing of the particles with bovine serum albumin. ( b ) Rotenone (and several other rotenoids), piericidin A, and Amytal bind noncovalently and inhibit a t the same specific binding site in phosphorylating and nonphosphorylating preparations. (c) Piericidin binds more tightly than rotenone, and titration data indicate that 2 moles of piericidin bind with comparable affinity per mole of NADH dehydrogenase in submitochondrial particles. (d) Titration curves relating the degree of NADH oxidase inhibition to inhibitor concentration are sigmoidal, thus indicating that the two binding sites are not equivalent in terms of their contribution to inhibition of electron transport. (e) Unlike submitochondrial particles, the number of binding sites per mole of NADH dehydrogenase in complex I and complex 1-111 is close to unity. Other aspects of rotenone and piericidin inhibition studied by Singer and co-workers are related more to submitochondrial particles than to complex I. These studies have been compiled in reviews (.22, 23) by these investigators and will not be detailed here. As stated above, the low molecular weight NADH dehydrogenase of Pharo et al. (64) was considered incorrectly to be the NADH-ubiquinone reductase of the respiratory chain. This was in part because the ubiquinone reductase activity of the preparation could be partially inhibited by Amytal and by very low concentations of rotenone. It was demonstrated by others that these effects were different from the inhibitions 91. G.Palmer, D.J. Horgan, H. Tisdale, T. P. Singer, and H. Beinert, JBC 243, 844 (1968). 92. M. E. Teeter, M. L. Baginsky, and Y. Hatefi, BBA 172, 331 (1969). 93. D. J. Horgan, T. P. Singer, and J. E. Casida, JBC 243, 834 (1968). 94. D.J. Horgan, H. Ohno, T. P. Singer, and J. E. Casida, JBC 243, 5967 (1968).

206

YOUSSEF HATEFI AND DIANA L. STIGGALL

obtained with complex I or submitochondrial particles (19, 22, $2). The results of Hatefi et al. (42) show the following differences: (a) Substantial inhibition of NADH-ubiquinone-1 reductase activity of the soluble, low molecular weight enzyme requires more than 100-fold as much rotenone as is necessary for a comparable degree of inhibition of complex I. (b) Barbiturates inhibit the ubiquinone reductase activity of the former enzyme only when higher ubiquinone isoprenologs are used as electron acceptor. Unlike the reaction catalyzed by complex I, barbiturates do not inhibit the ubiquinone-1 reductase activity .of the soluble dehydrogenase. (c) Piericidin A, which is the most potent inhibitor known for ubiquinone reduction by complex I and submitochondrial particles, is essentially ineffective on ubiquinone reduction by the soluble enzyme. Among several iron chelators used, only o-phenanthroline inhibited the soluble dehydrogenase (42). It was shown by Hatefi et al. (42) that incubation of the enzyme with o-phenanthroline results in the loss of labile sulfide, while pretreatment with bathophenanthroline sulfonate, Tiron (1,2-dihydroxybenzene 3,5-disulfonate) or ethylenediamine tetraacetate protects the enzyme against the loss of labile sulfide and inhibition of activity upon subsequent incubation with o-phenanthroline. The unique destructive ability of o-phenanthroline has been demonstrated by these investigators for several iron-sulfur proteins (95,96). While in contrast t o complex I the ferricyanide reductase activity of the low molecular weight dehydrogenase is not inhibited at high NADH concentrations (>0.2 mM) , its quinone reductase activities are. Indeed, there seems to be a correlation between NADH inhibition, the apparent Km of the enzyme for NADH, and whether the acceptor is a single-electron (ferricyanide, cytochrome c) or a two-electron (ubiquinones, menadione, and dichloroindophenol) recipient (40). With single-electron acceptors, the apparent K m for NADH is about 65 pM and the reaction is relatively insensitive to the concentration of NADH. However, with two electron acceptors, the apparent K , for NADH is twice as much (133 UJM) and the reaction is sharply inhibited a t NADH concentrations greater than 0.25 mM. Hatefi and Stempel (40) have suggested that these phenomena might be a consequence of the dissimilar affinity for NADH of the various oxidation-reduction states of the enzyme (i.e., oxidized, half-reduced, fully reduced) transiently produced during electron transfer to one-electron versus two-electron acceptors. Millimolar concentrations 95. Y. Hatefi and W. G. Hanstein, ABB 138,73 (1970). 96. R. M. Kaschnitz and Y. Hatefi, ABB 171,292 (1975).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

207

of NAD partially inhibit NADH dehydrogenase ( 4 0 ) . A similar inhibition is caused by AMP, ADP, and ATP, but not by adenosine ( 4 2 ) . Guanidinium hydrochloride and alkylguanidinium salts have been implicated as inhibitors of electron transport and oxidative phosphorylation a t site 1 (97, 98). Hatefi et al. (42) have shown that the ubiquinone reductase activity of complex I is inhibited by guanidinium hydrochloride (10-50 mM), but the ferricyanide reductase activity of complex I and all the reductase activities of the low molecular weight dehydrogenase are activated. Guanidinium ion was more potent than the alkylated derivatives] and the activation effect analyzed for menadione reduction indicated a decrease in K , for NADH and an increase in V,,,, both of which were dependent on the concentration (10-100 m M range) of guanidinium ion.

6. N A D P H Oxidation and N A D P H to N A D Tramhydrogenation b y Complex I The ability of submitochondrial particles to catalyze transhydrogenation from NADPH to NAD has been known for many years. The reverse reaction, i.e., from NADH to NADP, is slow but can be accelerated when energy is supplied to the system (e.g., ATP). The energy requirement of the reverse reaction and the different equilibria of the transhydrogenase reaction in the absence and presence of an energy supply are ~ U Z zling thermodynamic problems. The mitochondrial transhydrogenase reaction has been under vigorous investigation (99-102), a recent review is available (103), and the topic is covered in Chapter 2. Until 1973, it was generally agreed that the mitochondrial respiratory chain is incapable of oxidizing NADPH directly (103-105).NADPH oxidation was considered to occur only through the transhydrogenase reaction and with the obligatory intermediation of NAD (103-105). In 1973, 97. J. B. Chappell, JBC 238,410 (1963). 98. B. Chance and G. Hollunger, JBC 238,432 (1963). 99. J. Rydstrom, A. Teixeira da Cruz. and L. Ernster, Eur. J. Biochem. 17, 56 (1970). 100. A. Teixeira da Cruz, J. Rydstrom, and L. Ernster, Eur. J. Biochem. 23, 203 (1971). 101. J. Rydstrom, A. Teixeira da Cruz, and L. Ernster, Eur. J. Biochem. 23, 212 (1971). 102. R.R.Fisher and N. 0. Kaplan, Biochemistry 12, 1182 (1973). 103. N.0.Kaplsn, Harvey Lect. 66, 105 (1972). 104. L. Ernster, C.-P. Lee, and U. B. Torndal, in “The Energy Level and Metabolic Control in Mitochondria” (S. Papa et al., eds.), p. 439. Adriatrica Editrice, Bari, 1969. 105. F. A. Hommes, in “Energy-Linked Functions of Mitochondria” (B. Chance, ed.), p. 39. Academic Press, New York, 1963.

208

YOUSSEF HATEFI AND DIANA L. STIGGALL

(CI

FIQ.12. (A) First-derivative E P R spectra of complex I treated with NADH or NADPH. Conditions: complex I, 45 mg/ml; temperature 14°K; microwave frequency 9.225 GHz; power, 2 mW, modulation amplitude, 6.3 G ; gain, 50. g = 2 was at 3295 G. Where indicated 1.5 mM NADH or NADPH was added. Small letters of the alphabet in this, (B), (C), and Fig. 13 denote the same signals as in Fig. 4. (B) The EPR spectrum of NADH-treated complex I shown in (A) a t gain of 200 and 0.3 mW power. (C) The E P R spectrum of NADPH-treated complex I shown in (A) at gain of 200 and 0.3 mW power. From Hatefi and Hanstein (80).

Hatefi (106-108) and Hatefi and Hanstein (80) demonstrated, however, that NADPH is oxidized directly by the respiratory chain a t a site close to, but apparently not identical with, the site of NADH oxidation. It was found that NADPH oxidation by respiratory particles is very slow a t neutral pH (about 50 nmoles/min/mg protein) , but quite appreciable a t pH values between 5 and 6 ( 2 2 5 0 nmoles/min/mg protein). Indeed, the rate difference between pH 9 and pH 6 was found to be about 35-40-fold, 106. Y.Hatefi, BBRC 50, 978 (1973). 107. Y. Hatefi, i n “Dynamics of Energy Transducing Membranes” (L. Ernster R. W. Estabrook, and E. C. Slater, eds.), p. 125. Elsevier, Amsterdam, 1974. 108. Y.Hatefi, Fed. Proc., Fed. Amer. SOC. Exp. Biol. 32,595 (1973).

4.

m

I 0

r

t

-1

209

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

G=100

I-

n

s

\/

H-

FIQ. 13. Computer-derived difference of NADH-treated minus NADPH-treated complex I shown in Fig. 12. From Hatefi and Hanstein (80).

an indication of the fact that others working a t neutral or more alkaline pH values had missed (or dismissed as resulting from the presence of traces of NAD) the direct oxidation of NADPH by the respiratory chain. Both NADPH dehydrogenase (assayed with ferricyanide as acceptor) and NADPH-to-NAD transhydrogenase activities of respiratory particles were found to fractionate mainly into complex I. Electron paramagnetic resonance studies on complex I a t neutral pH showed that NADPH reduced iron-sulfur center 2 and partially the overlapping iron-sulfur centers 3 4. Iron-sulfur center 1 was not detectably reduced by NADPH, nor was the flavin of complex I as evidenced from the difference absorption spectra of NADPH-treated minus NADH-treated complex I. In agreement with previous findings described above, all iron-sulfur centers were reduced by NADH. These results are depicted in Figs. 12-14. Sub-

+

-0.0

400

500

600

Wavelength Inm)

FIG.14. Absorption spectrum of NADPH-treated minus NADH-treated complex I. Conditions: complex I, 6 mg protein/ml of 0.66 M sucrose containing 50 mM Tris-HC1 ( p H KO), 1 mM histidine, and 0.25% (v/v) Triton X-100. The sample cuvette was treated with 200 pM NADPH, and the reference cuvette with 100 p M NADH. Dashed line, untreated complex I in both cuvettes. From Hatefi and Hanstein (80).

210

YOUSSEF HATEFI AND DIANA L. STIGGALL

5

6

7

a

9

PH

FIQ.15. pH dependence of NADPH oxidase, NADPH to 3-acetylpyridine adenine dinucleotide (AP-DPN) transhydrogenase, and NADH oxidase activities of submitochondrial particles (ETP), Conditions: oxidase activities were measured in the presence of 2 mM NADH or NADPH, 0.25 121 sucrose, 100 mM sodium phosphate for pH values 6-9, and 100 mM sodium acetate for pH values 5.0 and 5.5. ETP concentration was 2.16 mg/ml for the NADPH oxidase, and 0.216 mg/ml for the NADH oxidase assays. The transhydrogenasc reaction was measured by the AmincoChance spectrophotometer at 400 minus 450 nm. The extinction coefficient used for reduced 3-acetylpyridine adenine dinucleotide a t 400 nm was 2300 liters mole-' cm-'. Media were the same as in the oxidase assays. Dotted lines indicate uncertainty about the pH 5 rates because of possible acidity damage to ETP. The ordinate refers to nanomoles of NADPH or NADH oxidized min-' x mg-' of ETP protein at 30". From Hatefi and Hanstein (80).

sequent studies of Hatefi and Bearden (109) indicated that as compared to NADH reduced complex I, the low field signal due to overlapping centers 3 4 was not only smaller but also a t a position approximately 3 G upfield (corresponding to Ag = 0.002) when it was generated with NADPH as reductant. These results indicated, therefore, that the partial center 3 4 reduction by NADPH might have resulted mainly or entirely from center 3 (see Table IV). Thus, i t appears that a t neutral pH NADPH can reduce those components of complex I (iron-sulfur centers 2 and 3) whose reduction potentials appear to be close to zero, but not those whose reduction potentials are between -300 and -200 mV (flavin and iron-sulfur centers 1 and 4, see Section II,A,7). As stated above, submitochondrial particles and complex I exhibit both NADPH dehydrogenase and NADPH-to-NAD transhydrogenase activities. These activities have similar pH dependencies (Fig. 15) and are

+

+

109. Y . Hatefi and A. J. Bearden, unpublished.

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211

both specific with respect to abstraction of the 4-B hydrogen of NADPH (10'7). The NADH dehydrogenase of the respiratory chain is also 4-B specific, whereas transhydrogenation from NADH to NADP is 4-A specific in agreement with the 4-B specificity of the reverse reaction with regard to NADPH. The similarities between NADPH dehydrogenase and NADPH-to-NAD transhydrogenase have suggested that both reactions might be catalyzed by the same enzyme. They have also created suspicion regarding the noninvolvement of NAD in NADPH dehydrogenation in spite of the differences detailed above in the reduction of respiratory components by NADH and NADPH and the demonstration of the absence of detectable NAD in the experiments involving complex I (submitochondrial particles contain about 0.2 nmole NAD per mg protein) (80). Two lines of evidence have furnished unambiguous evidence, however, that NADPH oxidation by submitochondrial particles and complex I can occur under conditions that the transhydrogenase reaction is completely inhibited. The differences in the reduction of respiratory components with NADH or NADPH as substrate is reflected in the degree of bleaching afforded by these substrates a t 475 minus 510 nm in complex I and rotenoneor piericidin-treated submitochondrial particles. Thus, as seen in Fig. 16 (left-hand trace), addition of NADH doubles the bleaching a t 475 minus 510 nm obtained by addition of NADPH to piericidin-treated particles. The middle trace shows a similar effect when NAD is added instead of NADH. This results from the presence of transhydrogenase activity, which yields NADH from NAD and excess NADPH. I n the right-hand

475.510 nm

#l

i 1 min+

0 CI

>

FIG.16. Effect of palmitoyl-coenzyme A on reduction of chromophores a t 475 minus 510 nm in ETP via NADPH to NAD transhydrogenation. Conditions: ETP, 2.2 mg protein/ml; NADPH, 60 p M ; NADH, 60 p n l ; NAD, 140 pM; sodium succinate, 1.75 m M ; piericidin A , 5.3 pM ; antimycin A, 1 p M ; 2-thenoyltrifluoroacetone (TTFA), 1 m M ; palmitoyl-CoA (P-CoA), 200 pM. From Hatefi and Hanstein (80).

212

YOUSSEF HATEFI AND DIANA L. STIGGALL

trace the preparation has been treated with appropriate amounts of palmitoyl coenzyme-A (110) to inhibit the transhydrogenase reaction. The bleaching by NADPH is seen, the NAD effect resulting from the transhydrogenase reaction is largely, abolished, but subsequent NADH addition is still effective. These results clearly show that NADPH reduction of components as measured a t 475 minus 510 nm can occur under conditions that transhydrogenation to NAD is inhibited. That the latter reaction was substantially inhibited in these experiments is clear because in the presence of a piericidin block even a slow production of NADH would have resulted in bleaching just as NADPH did under conditions of low electron flux (pH 7.5 and NADPH concentration equivalent to 0.1 K,,,). Similar results were reported for complex I (80). I n addition, it has been shown by Djavadi-Ohaniance and Hatefi (111) that trypsin treatment of submitochondrial particles can distinguish among NADH oxidation, NADPH oxidation, and NADPH-to-NAD transhydrogenation. The exceptional sensitivity of the latter reaction to trypsin was demonstrated earlier by Ernster and his colleagues (119). Taking advantage of this sensitivity, the former investigators have shown that treatment of submitochondrial particles a t Oo with appropriate amounts of trypsin can completely destroy the NADPH-to-NAD transhydrogenase activity without affecting appreciably either the NADH or the NADPH oxidase activity (Fig. 17). Incubation of the particles a t 30° in the presence of trypsin then led to gradual loss of NADPH oxidase activity without affecting NADH oxidase activity. These results clearly demonstrate that the three reactions shown in Fig. 17 are independently affected by trypsin: transhydrogenase activity is rapidly destroyed a t Oo, NADPH oxidase is gradually destroyed a t 30°, while NADPH oxidation is unaffected by trypsin under these conditions. An important question that is raised by these experiments is the relationship between the NADPH dehydrogenase and the NADPH-to-NAD transhydrogenase activities of submitochondrial particles. On the one hand, they are clearly distinguishable on the basis of their sensitivities to trypsin and palmitoyl coenzyme-A. On the other hand, they exhibit common features with regard to their stereospecificities for abstraction of the 4-B hydrogen of NADPH, their extremely large response to pH change, and their copurification into complex I. It is entirely possible that the two activities might be catalyzed by the same enzyme because (a) many nicotinamide110. J. Rydstrorn, A. V. Panov, G. Paradies, and L. Ernster, BBRC 45, 1389 (1971). 111. L. Djavadi-Ohaniance and Y . Hatefi, JBC, in press. 112. K.Juntti, U. B. Torndal, and L. Ernster, in “Electron Transport and Energy Conservation” (J. M. Tager et a l , eds.), p. 257. Adriatica Editrice, Bari, 1970.

4.

m

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

5

(a)

15

25

40

213

55

(b)

FIG. 17. Effect of trypsin on the NADH oxidase, NADPH oxidase and the NADPH-to-NAD transhydrogenase activities of submitochondrial particles. The particles suspended in 0.25 M sucrose and 100 mM sodium phosphate, pH 7.0, were treated with 0.1 mg trypsin per mg particle protein and incubated a t (a) 0" or (b) 30". At the intervals shown samples were removed and assayed at pH 6.0 and 7.0 for the activities shown. Transhydrogenase activity was measured either directly by reduction of 3-acetylpyridine adenine dinucleotide at 375 nm in the presence of cyanide-treated particles or by the increase in the rate of NADPH oxidation by submitochondrial particles after the addition of NAD. (A) NADH + O,, ( 0 ) NADPH + 02,and (0) NADPH + NAD. From Djavadi-Ohaniance and Hatefi (111).

adenine dinucleotide dehydrogenases, including the mitochondrial NADH dehydrogenase, exhibit transhydrogenase activity in the presence of a suitable analog, and (b) the exceptional trypsin sensitivity of the transhydrogenase reaction might be concerned mainly with the NAD ( H ) binding site. For example, it is possible that similar to Rhodospirillum rubrum (113) the mitochondrial transhydrogenase system also involves a soluble protein cofactor. This protein factor or its association with the membranes might be susceptible to the action of trypsin. A more attractive possibility is suggested by the recent work of Vallee and his colleagues (114). They have found that in a number of enzyme-active sites arginyl residues serve as the positively charged recognition sites for negatively charged substrates, and have identified arginyl residues a t the NADH binding site of a number of alcohol dehydrogenases from various sources. A peptide bond involving the carboxyl group of arginine, if present a t the substrate binding site of the mitochondrial transhydrogenase, could be particularly susceptible to attack by trypsin. 113. R. R. Fisher and R. J. Guillory, JBC 244, 1078 (1969). 114. L. G. Lange, 111, J. F. Riordan, and B. L. Vallee, Biochemistry 13, 4361 (1974) .

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YOUSSEF HATEFI AND DIANA L. STIGGALL

7. Energy Conservation and Coupling by Complex I It has long been known that the first site of energy conservation in the respiratory chain is located between NADH and ubiquinone-cytochrome b. Schatz and Racker (44) demonstrated ATP synthesis by submitochondria1 particles a t the expense of NADH oxidation by externally added ubiquinone-1, and more recently Ragan and Racker (36)reconstituted oxidative phosphorylation a t site 1 in a system composed of complex I and appropriate membrane proteins, coupling factors, and phospholipids. The important experiments of Ragan and Racker showed that in principle the isolated and purified complexes are capable of energy conservation and coupling. The studies of Hatefi, Galante, and You (106, 107, 115) have shown that comparable ATP yields (P/O = 2.62.8) are obtained as a result of oxidation of NADH or NADPH by submitochondrial particles. Since the components of complex I reduced by both NADH and NADPH are iron-sulfur centers 2 and 3 and ubiquinone, the above experiments implicate these electron carriers. in site 1 energy conservation. Gutman and his colleagues (116) have shown that in piericidin-treated submitochondrial particles iron-sulfur center 2 remains reduced after exhaustion of added NADH through the piericidin leak. Iron-sulfur center 2 could be reoxidized by addition of ATP. This reaction was sensitive to uncouplers, and appeared to result from energy-linked reverse electron transfer to NAD. On the basis of these observations, Gutman et al. have concluded that coupling site 1 is located on the oxygen side of iron-sulfur center 1 and the substrate side of both iron-sulfur center 2 and the site of roteaone-piericidin block. These conclusions are in general agreement with the results of Hatefi et al. (80,106-108) regarding the NADPH reducible iron-sulfur centers and P/O > 2 obtained during NADPH oxidation. It is also interesting to note that energy-linked transhydrogenation can be induced by ATP or as a result of succinate oxidation in rotenonetreated particles (117). Further, Van de Stadt et al. (118) and Skulachev (119) have presented data regarding energy production in rotenoneblocked particles as a result of transhydrogenation from NADPH to NAD. Therefore, it appears that energy communication with the transhydrogenase reaction also occurs a t or near site 1. This would indeed 115. Y. Hatefi, Y. Galante, and K. S. You, unpublished. 116. M. Gutman, T. P. Singer, and H. Beinert, Biochemistry 11, 556 (1972). 117. C.-P. Lee, G. F. Azzone, and L. Ernster, Nature (London) 201, 152 (1964). 118. R. J. Van de Stadt, F. J. R. M. Nieuwenhuis, and K. Van Dam, BBA 234, 173 (1971). 119. P. Skulachev, Curr. T o p . Bioenerg. 4, 127 (1971).

4.

METAL-CONTAINING

215

FLAVOPROTEIN DEHYDROGENASES (SDH Fe/S) +3omV

Rotenone NADH t) (Center lo.lb)(Center3,4)-(Center

Halfreduction potentiol

-305mV

-245rnV

-20rnV

2) -(Center

1

5 ) +-+(R~eske’s

+4OrnV

Fe/S)-

O2

+ 280rnV

FIG. 18. Thermodynamic profile of iron-sulfur centers in pigeon heart mitochondria : SDH, succinate dehydrogenase ; Rieske’s Fe/S, iron-sulfur protein of complex 111. From Ohnishi (121).

be plausible if as suggested by their similar enzymic features NADPH oxidation and nicotinamide-adenine dinucleotide transhydrogenation shared through a common enzyme the same linkage to the respiratory chain. Ohnishi et al. (120, 121) have suggested that coupling site 1 involves not only iron-sulfur center 2 but also half of iron-sulfur center 1 (designated center l a ) . Their conclusion is based on an apparent change in the reduction potential of these centers induced by ATP as estimated from the measurement of E P R signals in the presence of redox mediators. Similar criteria were used by Wilson and Dutton (122, 123) to identify cytochromes b, and a3 as energy transducing components a t coupling sites 2 and 3, respectively. However, these experiments have been criticized by Caswell (124) and Lambowitz e t al. (125).They feel that complications resulting from improper equilibration of the redox mediators with the electron carriers under study and ATP-induced reverse electron transfer to and from these components have been the underlying basis of the measured redox potential changes brought about by A T P addition. According t o Ohnishi (121) , the half-reduction potentials of the various iron-sulfur centers of the respiratory chain a t pH 7.2 are as shown in Fig. 18. The value for center 1 is essentially in agreement with the results of Orme-Johnson et al. (46, 54) who found that reduced acetylpyridine adenine dinucleotide (Eo’ = -248 mV) can only reduce this center by 50%, while its oxidized form can effectively oxidize iron-sulfur center 120. T. Ohnishi, D. F. Wilson, and B. Chance, BBRC 49, 1087 (1972). 121. T. Ohnishi, BBA 301, 105 (1973). 122. D. F. Wilson and P. L. Dutton, BBRC 39,59 (1970). 123. D. F. Wilson and P. L. Dutton, A B B 136, 583 (1970). 124. A. H. Caswell, A B B 144,445 (1971). 125. A. M. Lambowitz, W. D. Bonner, Jr., and M. K. F. Wikstrom, Proc. N u t . Acud. Sci. U . S. 71, 1183 (1974).

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YOUSSEF HATEFI AND DIANA L. STIGGALL

-.I

NADPHNADP

FeS4

FIQ. 19. Proposed electron transfer pathways for oxidation and reduction of NADH/NAD and NADPH/NADP, and energy coupling site 1 in complex I. Where applicable broken arrows indicate energy-linked electron or hydride ion transfer. FeS, iron-sulfur center.

1 in dehydrogenase preparations. These investigators feel, however, that the reduction potential of center 3 is greater than or equal to that of center 2, since upon titration of complex I with graded amounts of NADH or dithionite reduced centers 2 and 3 appear long before centers 4 and 1 are reduced. Thus, it seems that complex I contains two iron-sulfur centers (1 and 4) with reduction potentials close to that of NADH (Eo’= -315 mV), and two centers (2 and 3) with potentials close to that of ubiquinone (Eo’N +65 mV) . Accordingly, the largest single-step energy drop in the NADH pathway is between iron-sulfur centers 4 and 2 + 3 (AE = -225 mV; AGO‘ for 2e = -10,400 cal), and in the NADPH pathway is, so far as known, between NADPH and iron-sulfur centers 2 3 (AE = -295 mV; AGO’ for 2e = -13,600 cal). I n theory, therefore, the energy liberated a t each of these two steps appears to be compatible with the amount required for ATP synthesis a t site 1 with either NADH or NADPH as substrate. The above considerations are summarized in Fig. 19.

+

B. NADH DEHYDROGENASES OF YEAST The NADH dehydrogenase of yeast is of considerable interest because in Saccharomyces cerevisiae and Saccharomgces carlsbergensis coupling site 1 is absent, whereas in Candida utilis its existence depends on the growth phase of the cells and can be altered by adaptations to culture conditions and by catabolite repression. In 1961, Vitols and Linnane (126) reported that mitochondria isolated from S. cerevisiae showed identical P/O values for oxidation of succinate and NAD-linked substrates. These results were the first indication of the absence of coupling site 1 in S. cerevisiae. Mackler et al. (127) and 126. E. Vitols and A. W. Linnane, J. Biophys. Biochem. Cytol. 9, 701 (1961). 127. B. Mackler, P. J. Collipp, H. M. Duncan, N. A. Rao, and F. M. Huennekens, JBC 237, 2968 (1962).

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Mahler and co-workers (128) isolated respiratory particles from bakers’ yeast, and showed that NADH oxidation by these particles was insensitive to inhibition by Amytal, Seconal, and rotenone. The former authors also demonstrated that, unlike mammalian respiratory particles, the flavin of S. cerevisiae particles was exclusively FAD of which approximately 50% was acid-extractable. Extraction of the remainder required prior digestion of the particles with trypsin. Duncan and Mackler (129) isolated a low molecular weight NADH dehydrogenase from these particles by the acid-ethanol procedure (see above). The preparation contained per mg protein 10.6 nmoles of FAD, 6.3 ng atoms of iron, and no labile sulfide. The molecular weight was determined by equilibrium sedimentation to be approximately 55,000. Thus, it appeared that during isolation the enzyme had lost about 40% of its flavin, assuming one mole of FAD per mole of enzyme. The possibility of iron and labile sulfide loss is also suggested by the use of acid pH, prolonged dialysis, and DEAE-cellulose chromatography for isolation and purification of the enzyme. The dehydrogenase could react with ferricyanide, dichloroindophenol, and cytochrome c as electron acceptors, and in all instances added FAD increased the activity and p-mercuriphenyl sulfonate inhibited it. Thus, in terms of size, acceptor specificity, and mercurial sensitivity, the NADH dehydrogenase from S. cerevisiae respiratory particles appears to be comparable to a similar preparation from mammalian mitochondria. The major difference is that the flavin of the latter is FMN. According to Kim and Beattie (130),the appearance of NADH dehydrogenase activity in mitochondria of glucose derepressed S. cerevisiae is blocked by cycloheximide, but not by chloramphenicol, suggesting that NADH dehydrogenase does not contain products of mitochondria1 protein synthesis. The NADH dehydrogenase system of C . utilis is very similar to that of mammalian mitochondria. Both systems are inhibited by rotenone and piericidin A, conserve energy a t coupling site 1, contain multiple forms of EPR-active iron-sulfur center, have F M N as their flavin prosthetic group, react best with ferricyanide as electron acceptor, and are inhibited a t high substrate concentrations (131-133).Biggs et al. (132) were unsuccessful in dissociating a high molecular weight type enzyme from C. utilis 128. H. R. Mahler, B. Mackler, S. Grandchamp, and P. P. Slonimski, Biochemistry 3, 668 (1964). 129. H. M. Duncan and B. Mackler, Biochemistry 5, 45 (1966). 130. I. C.Kim and D. S. Beattie, Eur. J . Biochem. 36,509 (1973).

131. P.A. Light, C. I. Ragan, R. A. Clegg, and P. B. Garland, FEBS (Fed. Eur. Biochem. Soc.) Lett. 1, 4 (1968). 132. D. R. Biggs, H. Nakamura, E. B. Kearney, E. Rocca, and T. P. Singer, ABB 137, 12 (1970). 133. S. 0. C. Tottmar and C. I. Ragan, BJ 124,853 (1971).

218

YOUSSEF HATEFI AND DIANA L. STIGGALL 0.06 r Wavelength (nm)

h

350

400

450

500

550

600

650

700

Wnvalongth (nm)

FIG.20. Absorption spectra of the purified NADH dehydrogenase of C. utilis at 0.8 mg/ml. Trace (a), oxidized enzyme; trace (b) after addition of 0.1 mM NADH; inset, (b) minus (a). From Tottmar and Ragan (138).

particles by digestion with phospholipase A. However, Tottmar and Ragan (133) have isolated such a preparation with the use of deoxycholate and Triton X-100. The preparation contains per mg protein 0.5-0.6 nmole of flavin ( F M N ) , 15-17 ng-atoms of iron, and 15-17 nmoles of labile sulfide. It reacts efficiently with ferricyanide as electron acceptor, but poorly with menadione, cytochrome c, dichloroindophenol, and ubiquinone-1. The K , and K i of the enzyme with respect to NADH are, respectively, 83 pM and 0.2 m M ; the K , for ferricyanide is 1.0 mM. The preparation exhibits a g = 1.94 E P R signal, which is characteristic of complex I and the high molecular weight dehydrogenases from heart mitochondria. Its absorption spectrum (Fig. 20) is qualitatively comparable to that of the mammalian high molecular weight dehydrogenase, and similar to complex I it can oxidize NADPH a t a slow rate (133) (see Table V I ) . The absence of rotenone-piericidin sensitivity and coupling site 1 in S. cerevkiae and S. carlsbergensis, and their presence in C. utilis have suggested a possible connection between the two phenomena. Light et al. (131) demonstrated that in C. utilis iron-limited growth conditions result in a decrease of mitochondria1 cytochromes and nonheme iron, loss

4.

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FLAVOPROTEIN DEHYDROGENASES

219

of site 1 phosphorylation, loss of sensitivity of NADH oxidation to inhibition by rotenone and piericidin A, and loss of piericidin A binding capacity. These changes were not reflected, however, in the respiratory activity of the mitochondria. Site 1 phosphorylation and rotenone-piericidin sensitivity were recovered when the iron-deficient cells were incubated with FeS04 in the absence of an added carbon source. Others have reported that under various growth conditions, energy conservation a t site 1 reappears upon aeration of deficient cells concomitant with (134) or without (135) the appearance of piericidin sensitivity. These developments have been reviewed by Ohnishi (f.21). More recently, a careful study of the problem has been done by Grossman et al. (136) in which they have followed NADH dehydrogenase activity, appearance and loss of piericidin sensitivity, and the nature of various E P R signals and cytochromes during the exponential and stationary phases of C . utilis growth as well as after catabolite repression. They found that the respiratory particles of cells harvested during the exponential growth phase have very low piericidin sensitivity and NADHferriayanide reductase activity, but high NADH-juglone (5-hydroxy-1,4naphthoquinone) reductase activity (Fig. 21). Successive washing of the particles with low osmolarity buffer resulted in the loss of NADH dehydrogenase activit,y. The EPR spectra of antimycin-treated particles reduced with NADH or dithionite showed a n absence of signals resulting from iron-sulfur centers 1 and 2, but centers 3 4 were present. Signals were also present a t gll = 2.01 and gL = 1.92, which appeared t o result from an iron-sulfur center. The temperature sensitivity of this new species was similar t o that of center 1. Particles of cells in the stationary phase showed an increase in P/O ratio, suggesting the appearance of site 1 coupling, presence of piericidin sensitivity, rise in ferricyanide reductase activity, loss of juglone reductase activity (Fig. 21), appearance of EPR signals resulting from centers 1 and 2, and loss of the new EPR signal seen in exponential phase particles. Further, the dehydrogenase of the stationary phase was stable to washing of the particles. Catabolite repression of late stationary phase cells by addition of ethanol resulted in the loss of the above characteristics and reacquisition of exponential phase properties with regard to enzymic activity, piericidin sensitivity, and the EPR signals associated with NADH dehydrogenase. These changes were prevented by cy cloheximide.

+

134. T. Ohnishi, P. Panebianco, and B. Chance, BBRC 49,99 (1972). 135. R. A. Clegg and P. B. Garland, BJ 124, 135 (1971). 136. S. Grossman, J. G. Cobley, T. P. Singer, and H. Beinert, JBC 249, 3819 (1974).

220

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10

20

30 40 50 Hours FIG.21. Characteristics of NADH oxidation by submitochondrial particles from C. utilis during transformation from exponential to stationary phase. Candida utilis was grown in 1.5% (v/v) ethanol in a fermentor at 30". Cells were harvested a t the times shown for isolation of mitochondria and preparation of submitochondrial particles. NADH oxidase activity is expressed as microatoms of oxygen per min per mg protein at 30". NADH dehydrogenase activity is expressed as micromoles of NADH oxidized per min per mg particle protein a t 25" at V,, with respect to Fe(CN).'-; sensitivity to piericidin A (0.5 nmole/mg protein) is expressed as percent inhibition of NADH oxidase; and turbidity is given as absorbance at 650 nm in 1 cm light path. The pH was maintained during growth by automatic addition of 6 N KOH (pH-stat) at 5.0 until 25 hr, after which no further acid development occurred but the pH rose to between 5.0 and 6.2. From Grossman et al. (136).

The authors feel that during transition from the exponential to the stationary phase, a different type of NADH dehydrogenase is synthesized (136). However, a very interesting possibility suggests itself when one compares the characteristics of NADH dehydrogenase in the stationary phase and exponential phase (or catabolite repressed) particles, respectively, with the properties of complex I and chaotrope-destabilized complex I (see above). Stationary phase particles and complex I are capable of energy conservation a t site 1, are piericidin-sensitive, have high ferricyanide reductase and low naphthoquinone reductase activities (juglone was used with stationary phase particles and menadione with. complex I ) , and exhibit EPR signals resulting from centers 1 (responsible for g = 1.94 signal) and 2. In both cases, the membrane-bound dehydrogenase is stable. In contrast, destabilized complex I and particles from

4.

METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

221

exponential phase or catabolite-repressed C . utilis cells exhibit low ferricyanide reductase and high naphthoquinone reductase activities,' loss of piericidin sensitivity and g = 1.94 E P R signal, and appearance of new EPR signals (46, 136). In the fractions isolated from destabilized complex I, the modified EPR signal is seen in the soluble, low molecular weight dehydrogenase ( 4 6 ) , which presumably originates from the temperature-insensitive center 1 signal of the unperturbed complex. Further, the dehydrogenase activities of both destabilized complex I and exponential phase or repressed particles are unstable. These analogies suggest, therefore, that in C . utilis cells at exponential growth phase or subjected to catabolite repression a t late stationary phase there exist species of NADH dehydrogenase with features akin to the mammalian low molecular weight enzyme derived from destabilized particles. Assuming that the low molecular weight dehydrogenase of mammalian mitochondria is a bona fide component rather than a degradation product, then its different properties when membrane bound might be a consequence of integration into the membrane and interaction with other respiratory components. By analogy, it is conceivable that during the exponential growth phase a species comparable to the low molecular weight dehydrogenase is present and in the process of being assembled into the respiratory chain, while after catabolite repression the membrane becomes degraded and this species of dehydrogenase exhibits once again the properties of the unbound enzyme.

C. NADH DEHYDROGENASE OF Azotobacter winelandii Low molecular weight NADH dehydrogenases have been isolated by DerVartanian (137) from A. winelandii grown under normal and ironlimited conditions. Both preparations are reported to have a molecular weight of 56,500, and to contain FRIN. The enzyme from cells grown under iron-limited conditions contained 1 g-atom of molybdenum, 2 g-atoms of iron, and 2 moles of labile sulfide per mole of FMN, while in the enzyme from cells grown under normal conditions the iron and labile sulfide content was twice as much. The two preparations exhibited different EPR signals but comparable NADH-menadione and NADHferricyanide reductase activities. It is stated that, unlike the mammalian low molecular weight enzyme, the purification of the Azotobacter low molecular weight NADH dehydrogenase is not accompanied by changes in catalytic properties. 137. D. V. DerVartanian, Z. Naturforsch. 27, 1082 (1972).

222

YOUSSEF HATEFI AND DIANA L. STIGGALL

111. Succinate Dehydrogenarer

A. MAMMALIAN SUCCINATE DEHYDROGENASE (EC 1.3.99.1) Succinate dehydrogenase is the only enzyme of the citric acid cycle which is bound to the inner membrane of mitochondria. It is also one of three flavoproteins known in which flavin is covalently linked to the protein. The other two are monoamine oxidase of the outer membrane of liver mitochondria (138) and Chromatiurn cytochrome c-552 (139). Succinate dehydrogenase was solubilized from beef heart mitochondria in 1954 (140) and purified in 1970 (141-143). In the intervening years modified or new procedures for isolation and purification of the enzyme were reported by Bernath and Singer (144), Basford et al. (145),Wang et al. (l46‘),Keilin and King (147, 148),Veeger et al. (149), and Cerletti et al. (160). The preparations of various laboratories differed in their content of covalently bound flavin (hence in degree of purity), nonheme iron, and labile sulfide, and exhibited different enzymic activities, the most important of which was the ability of the enzyme to transfer electrons to the respiratory chain. Consequently, unresolvable controversies developed and strong positions were taken regarding the molecular weight, composition, activities, and regulatory properties of succinate 138. E. B. Kearney, J. I. Salach, W. H. Walker, R. Seng, W. Kenney, E. Zeszotek, and T. P. Singer, Eur. J. Biochem. 24, 321 (1971). 139. W. C. Kenney, D. Edmondson, R. Seng, and T. P. Singer, BBRC 52, 434 ( 1973).

140. T. P. Singer and E. B. Kearney, BBA 15, 151 (1954). 141. Y. Hatefi, K. A. Davis, W. G. Hanstein, and M. A. Ghalambor, A B B 137, 286 (1970). 142. W. G. Hanstein, K. A. Davis, and Y. Hatefi, in “Energy Transduction in

Respiration and Photosynthesis” (E. Quagliariello, S. Papa, and C. S. Rossi, eds.), p. 495. Adriatica Editrice, Bari, 1971. 143. K. A. Davis and Y. Hatefi, Biochemistry 10, 2509 (1971). 144. P. Bernath and T. P. Singer, “Methods in Enzymology,” Vol. 5, p. 597, 1962. 145. R. E. Basford, H. D. Tisdale, and D. E. Green, BBA 24, 290 (1957). 146. T. Y. Wang, C. L. Tsou, and Y. L. Wang, Sci. Sinicn 5, 73 (1956). 147. D. Keilin and T. E. King, Nature (London) 181, 1520 (1958). 148. T. E. King, JBC 238, 4037 (1963). 149. C. Veeger, D. V. DerVartanian, and W. P. Zeylemaker, “Methods in Enzymology,” Vol. 13, p. 81, 1969. 150. P. Cerletti, G. Zanetti, G. Testolin, C. Rossi, F. Rossi, and G. Osenga, in “Flavins and Flavoproteins,” 3rd Int. Symp. (H. Kamin, ed.), p. 629. Univ. Park Press, Baltimore, Maryland, 1971.

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

223

dehydrogenase. These issues have been discussed in a number of reviews by Singer and his colleagues from their laboratory’s standpoint. Their most recent are references 23 and 25. 1. Molecular Properties Singer and co-workers extracted succinate dehydrogenase from mitochondrial acetone powder a t alkaline pH, and applied purification procedures involving adsorption on calcium phosphate gel and ammonium sulfate fractionation (140, 144). Summarizing their results, Singer ( 1 5 ) concluded in 1966 that succinate dehydrogenase had the following molecular properties: “The minimum molecular weight of homogeneous preparations from nonheme iron content is 49,000 and from flavin content 200,000. Physical measurements support a molecular weight of approximately 200,000. This value is also in accord with gel exclusion studies on Sephadex G-200. Thus the enzyme contains 1 mole of flavin and 4 g-atoms of nonheme iron per mole. . . . The sedimentation velocity of the beef heart enzyme at 10-15 mg protein/ml is 6.5 S . . . .” This preparation could oxidize succinate in the presence of ferricyanide or phenazine methosulfate (PMS) as electron acceptor but was unable to transfer electo be unable to interact with the respiratory chain. Wang et al. (146) reported in 1956 the isolation of succinate dehydrogenase from heart muscle preparations treated with succinate and cyanide. The enzyme was extracted with 20% aqueous ethanol a t p H 9.0. The final product, after adsorption on calcium phosphate gel and ammonium sulfate fractionation, was stated to be electrophoretically homogeneous and to contain 1 mole of flavin and 4 g-atoms of iron per 140,000160,000 g of protein. While apparently purer than the preparations of Singer’s group, the cyanide-treated enzyme was also shown subsequently to be unable to interact with the respiratory chain. In 1958, Keilin and King (147) reported that succinate dehydrogenase isolated essentially by the method of Wang et al. (146), but without the use of cyanide, had an important property. Unlike other preparations, the Keilin-King enzyme was capable of electron transfer to the respiratory chain (148). It was subsequently shown by King (148) that the presence of succinate during extraction of the enzyme was essential for its ability to interact with the respiratory chain. Preparations of the enzyme contained 2.4-3.6 nmoles of flavin per mg protein and high levels of nonheme iron and labile sulfide. An average of several preparations showed a ratio of iron to sulfide to flavin of 8.5:8.1: 1 (151). By compari151.

T.E. King, BBRC 16,511 (1964)

224

YOUSSEF HATEFI AND DIANA L. STIGGALL

son to the preparation of Singer and co-workers and Wang et al., the Keilin-King enzyme contained twice as much iron per mole of flavin but less flavin per unit weight of protein. These data suggested the presence in the latter preparation of an additional iron-sulfur protein linking succinate dehydrogenase to the respiratory chain. One difficulty with regard to this possibility was that omission of succinate during enzyme isolation resulted in a preparation with similar composition, absorption spectrum, and dye reductase properties, but with complete lack of the ability to interact with the respiratory chain. Further, Veeger et al. (149) showed that modifications of the Keilin-King procedure yielded a preparation of succinate dehydrogenase which retained the ability to interact with the respiratory chain, but contained 1 mole of flavin, 8 g-atoms of iron, and 4-8 moles of labile sulfide per 200,00CL250,000 g of protein. Thus, it appeared that the ability to reconstitute with the respiratory chain is an important property of succinate dehydrogenase, which was lost in the earlier low-iron preparations. I n 1959, Ziegler and Doeg (152-164) reported the isolation of a particulate preparation from beef heart mitochondria which was capable of electron transfer from succinate to ubiquinone. This preparation was subsequently recognized by Hatefi et al. (27, 29) to be one of the four electron transfer complexes of the respiratory chain and is now generally referred to as complex 11. Preparations of complex I1 contain approximately 5 nmoles of covalently bound flavin per mg protein, and 7-8 g-atoms of iron and 7-8 moles of labile sulfide per mole of flavin. I n addition, it was shown by Ziegler and Doeg that complex I1 contained cytochrome b at a molar concentration comparable to flavin. These data cast strong doubt on the molecular weight of 200,000 for succinate dehydrogenase since, on the basis of its flavin content, complex I1 had a similar molecular weight while containing in addition cytochrome b and possibly other proteins. In agreement with this, Baginsky and Hatefi (155,156) showed in 1969 that a preparation of succinate dehydrogenase with a flavin content of 6-7 nmoles/mg protein could be isolated from complex 11. I n spite of these indications, it was generally assumed, however, that succinate dehydrogenase had been purified and its molecular weight was 200,000. The enzyme was finally purified in 1970 by selective resolution of complex I1 with chaotropic agents (141-143). It was shown to have a flavin 152. D. M. Ziegler and K. A. Doeg, BBRC 1, 344 (1959). 153. D. M. Ziegler and K. A. Doeg, ABB 97,41 (1962). 154. D. M. Ziegler, in “Biological Structure and Function” (T. W. Goodwin and 0. Lindberg, eds.), Vol. 2, p. 253. Academic Press, New York, 1961. 155. M. L. Baginsky and Y. Hatefi, BBRC 32,945 (1968). 156. M. L. Baginsky and Y. Hatefi, JBC 244,5313 (1969).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

225

content of 10.3 +- 4% nmoles per mg protein, a molecular weight of about 100,000, two unlike subunits, an iron:labile su1fide:flavin ratio of 7-8:7-8:1, very high dye reductase activity, and the ability to interact with the respiratory chain. Certain features of the chaotrope-induced resolution of complex I1 will be described because (a) the procedure is novel and interesting and ( b ) insofar as the authors have been able to ascertain this is the only published procedure which yields pure succinate dehydrogenase (156~). The molecular and enzymic properties of the above preparations are summarized in Table IX. a. Resolution of Complex I I with Chotropic Ions. The purification of succinate dehydrogenase is a simple process, which involves selective extraction of the enzyme from complex I1 in the presence of a chaotropic agent followed by precipitation with ammonium sulfate (72, 143). Before the addition of ammonium sulfate to the resolved complex, it is necessary to separate the soluble enzyme from the remainder of complex 11, which is particulate, by centrifugation. Otherwise, the addition of ammonium sulfate will reverse the resolution process, as will be seen below. Figure 22 shows the effect of several chaotropic agents on the resolution of complex 11. The ordinate is percent succinate-ubiquinone (solid lines) or SUCcinate-PMS (dotted line) reductase activity, and the abscissa is time. 156a. Subsequent to the announcement of the above findings by Hatefi’s group (141, I @ ) , two laboratories (167, 165) have claimed the purification of succinate dehydrogenase by modifications of earlier procedures. Righetti and Cerletti (167) have stated that using an earlier method (150) they have obtained pure succinate

dehydrogenase “containing 7-8 nmoles of peptide bound flavin.” Purification details for this preparation, which according to its stated flavin content can be at best 80% pure, have, not been given. The earlier procedure (150), according to which the above enzyme was made, showed in the best fraction a flavin content of 5.5-5.7 nmoles/mg protein. The other preparation claimed to be pure (23, 26) is that of Coles et 01. (15,s).This preparation was made by the original procedure of Singer and co-workers (144) with subsequent purification by chromatography on Sephadex G-200 columns (details not given). However, Coles et al. (165) stated that “Although the peak fractions from such columns appeared homogeneous in sucrose gradients, their histidyl flavin content was no higher (4 to 5 nmoles/mg) than previously reported ( 5 nmole/mg).” This agreed with the earlier results of Singer’s (16) quoted above that their molecular weight of 200,000 was “in accord with gel exclusion studies on Sephadex G-200.” Coles et al. further stated that “correction for impurities” observed on polyacrylamide gels stained with Coomassie Blue “raised the histidyl flavin content to 8 to 9 nmoles/mg. . . .” Such calculated values referred to elsewhere should not be misunderstood, however, to represent the actual flavin content of the preparation, which was not more than 5 nmoles/mg protein, thus indicating a t best 50% purity. 157. P. Righetti and P. Cerletti, FEBS (Fed. Eur. Biochem. Soc.) LetL.13, 181 (1971). 158. C. J. Coles, H . D. Tisdale, W. C. Kenney, and T. P. Singer, Physiol. Chem. Phys. 4, 301 (1972).

TABLE IX

MOLECULAR AND ENZYMIC PROPERTIES OF REPRESENTATIVE PREPARATIONS OF SIJCCINATE DEHYDROOENASE

Preparation Bernath-Singera wang et al! King" Veeger et a1.d Complex 118 Davis-Hatefie

Molecular weight

Flavin (nmole/mg protein)

Flavin :iron : labile sulfide ratio

200,000

4.2-5.0

1:4:?

-6,000

1.3

1:4:? 1:8.5:8.1 1:8:4-8

2,00O-3,500 -6,000

0.58 0.3

1:7-8:7-8 1:7-8 :7-8

10,000-11,000 10,000-11,000

140,OOO-160,000

329,000

200,000-250,000 200, OOO

97,000

6-7 3.04 4-5 5.0 10.3

Turnover number at

v,PMS

-3,900

KZa (mM)

0.3

0.3

Reconstitution activity Absent Absent Present

0

Present Present Present

X

From Singer (16)and Bernath and Singer (I&). The latest preparation of Coles el al. (168)haa similar properties. Turnover number calculated from data in Wang et al. (146). ~Flavinand flavin:iron:labile sulfide are average values from (161).Turnover number and molecular weight calculated from highest activity in King (171)and average flavin content, respectively. d Turnover number calculated from V , activity at 25" in DerVertanian et al. ($18). From Hate6 et d. (%), and references therein. a

<

sm

2H z &U E9 z

P

r m

2

P

E

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

227

FIG.22. Resolution of complex I1 with respect to succinate dehydrogenase by various chaotropes. Complex I1 was suspended in 50 mM Tris-HC1, p H 8.0. After addition of 0.6 M chaotrope the concentration of complex I1 was 8 mg/ml. After addition of the salts, samples were taken a t the intervals indicated and assayed for succinate-Q, and succinate-PMS reductase activities. Solid lines, succinateubiquinone-2(&) reductase activity ; dotted line, succinate-PMS reductase activity. Resolution temperature, 0" ; assay temperature, 38". The complex I1 preparations used in the experiments of this and subsequent Figs. 24 and 25 had specific activities between 40 and 45 pmoles Q2 reduced by succinate per min per mg protein. From Davis and Hatefi (169).

It is seen t h a t before addition of chaotropes, both activities were stable. When 0.6 M chaotrope was added ubiquinone reductase activity diminished, indicating resolution of the complex. However, PMS reductase activity remained unchanged because complex 11-bound and soluble succinate dehydrogenases have the same PMS reductase activities. Both the extent and the rate of resolution are functions of the concentration and potency of the chaotrope used. Further addition of 0.9 M NaClO, resulted in further resolution of complex I1 as evidenced by further loss of ubiquinone reductase activity. Once again PMS reductase activity was essentially unaltered. The small change is probably because the enzyme, unprotected in these experiments with succinate and dithiothreitol, was allowed to stand in the presence of 1.5 M NaC10, for many minutes. When the above protective agents are added, the small activity loss is prevented. I t has also been shown that the amount of succinate dehydrogenase solubilizcd is a linear function of the concentration of complex I1 (the range tested was 2.5-10 mg complex I1 protein/ml). The selectivity of the resolution of complex I1 with respect to succinate dehydrogenase is depicted in Fig. 23 on SDS-polyacrylamide gels stained

228

YOUSSEF HATEFI AND DIANA L. STIGGALL

FIQ.23. Profile of the purification procedure of succinate dehydrogenase depicted on SDS-polyacrylamide gels stained with Coomassie Blue. 1, complex 11; 2, first extraction of succinate dehydrogenase with 0.4 M NaClO,; 3, second extraction of succinate dehydrogenase with 0.75 M NaClO,; 4, particulate fraction of complex I1 remaining after twice removal of succinate dehydrogenase. From Davis and Hatefi (143).

with Coomassie Blue. Tube 1 is the starting complex 11. Tube 2 shows the two subunits of succinate dehydrogenase plus traces of impurity (about 6%) extracted with 0.4 M NaC104, and tube 3 shows the result of a second extraction of complex I1 with 0.75 M NaC104. The latter is essentially pure succinate dehydrogenase. Tube 4 is the remainder of complex I1 after the two successive extractions with 0.4 and 0.75 M NaClO,. It is seen in tube 4 that the stain intensities of the various polypeptide bands of complex I1 are essentially unchanged, except for the two bands corresponding to the extracted subunits of succinate dehydrogenase. As indicated by the results of Fig. 22, the resolution of complex I1 with respect to succinate dehydrogenase is an equilibrium process. At

4.

229

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

I

4

NaCIOe

1

'

o

'4 ' (a)

'

8'

-!/'

'

0

Min

'

'

4 (b)

'

'

1

8

FIG.24. Effect of temperature on the resolution of complex 11. Complex I1 was suspended in a solution containing 30 mM Tricine, 100 mM boric acid, 20 mM succinate and 5 mM dithiothreitol. pH of the buffer was adjusted to 8.1 with NaOH. Complex I1 was resolved by addition of 1.0 M NaClO, at the temperatures indicated. After addition of NaC10, the concentration of complex I1 was 8 mg/ml. (a) Resolution at O", lo", and 30". (b) Increased resolution and reconstitution as the temperature was changed from (0) 0" to ( 0 ) 30" and from 30" to O", respectively. Assays were performed a t 38". From Davis and Hatefi (169).

a given concentration of chaotrope, the extent of resolution increases with temperature (Fig. 24a) and is reversible when the temperature is decreased (Fig. 24b). Enthalpy and unitary entropy changes calculated from the data of Fig. 24 are, respectively, +4 kcal/mole and -15 eu. Chaotrope-induced resolution, as discussed elsewhere (70-72) , results from the destructuring effect of chaotropes on water and consequent weakening of the hydrophobic attractions of the system under study. Thus, it might be expected that increasing the structure of the aqueous medium by water structure-forming ions might shift the equilibrium in the direction of unresolved (or reconstituted) complex I1 (159).This is shown in Figs. 25a and 25b. I n Fig. 25a, complex I1 is first resolved t o the extent of about 50% by addition of 0.7M NaC104. Then various concentrations of ammonium sulfate are added. Since sulfate is a water structure-forming ion, the resolution is reversed and complete reconstitution is achieved a t about 0.46 M sulfate. Figure 25b (upper panel) shows the effect of other structure-forming (or antichaotropic) ions, and Fig. 25b 159. K.A. Davis and Y. Hatefi, ABB 149,505 (1972).

230

YOUSSEF HATEFI AND DIANA L. STIGGALL

0.7M1 NaC104

[(NHdaSOI]

-: ; ;7f[: 'ol

.* .r

80-

x

8

~1057~

-

0

t

8 -

@

---a---DilUtion ,/-

60-

* 40-

-

t

(NH4)tSO4

Fm. 25(a). Resolution of complex I1 by NaC104, and reconstitution by increasing concentrations of ammonium sulfate as indicated. Complex I1 was suspended in a solution containing 50 mM Tris-HC1, pH 8.0, 20 mlll succinate, and 5 mM dithiothreitol. After the addition of NaC104, the concentration of complex I1 was 8 mg/ml. Other conditions were the same as in Fig. 22. From Davis and Hatefi (169).

(lower panel) shows that removal of C10,- by precipitation as KC10, also results in reconstitution. That the degree of reconstitution is not a function of ionic strength is shown in Fig. 25b. I n agreement with the above, Davis and Hatefi (159) have also shown that chaotrope-induced resolution of complex 11in D,O, which is believed to be a more structured solvent than H,O, is considerably inhibited as compared to the same reaction carried out in H,O. The solvent isotope effect of D,O on the stability of membranous structures has been studied by Hanstein et al. (73). b. Subunits. It was shown in 1970 that succinate dehydrogenase is composed of two unlike subunits (141-143). These subunits could be separated by SDS-acrylamide gel electrophoresis or by resolution of succinate dehydrogenase under special conditions. The molecular weights of the subunits estimated from a number of separate experiments were 70,000 f 7% and 27,000 f 5% (143). Subsequent results of Righetti and Cerletti (157) have indicated molecular weights of 68,500 and 30,000, and those of Coles et at. (158) give molecular weights of 70,000 and 30,000, respectively, for the two subunits. The larger subunit carries the covalently bound flavin (143). Since treatment of succinate dehydrogenase with SDS for gel electrophoresis destroys the iron-sulfur chromophore and results in the loss of labile sulfide, a technique was devised by Davis and Hatefi (141-143)

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

23 1

min

FIU.25(b). Resolution of complex I1 by NnC104 and reconstitution by (upper panel) various nntichaotropic salts, and (lowrr panel) removal of C104- with K’. Conditions were the same as in Fig. 25a. At the concentrations used here and in Fig. 25~1,the antictiaotropic salts had no effect on the original activity of unresolved complex 11. From Davis and Hatefi ( 1 5 0 ) .

to resolve the enzyme in such a manner that the distribution of iron and labile sulfide in the two subunits could be determined (159a). The technique uses a combination of potent chaotropes and cold temperature, taking advantage of the inverse relationship of temperature with the strength of hydrophobic interactions. Thus, succinate dehydrogenase was 159a. I n spite of the known fact about the effect of SDS, Righetti and Cerletti (167) made the following surprising statement in their note on the subunits of succinate dehydrogenase: “The polypeptides were eluted from the gel and SDS was removed from the protein moiety. . . . Both subunits contain iron and labile sulfide. . . .”

232

YOUSSEF HATEFI AND DIANA L. STIGGALL

treated a t pH 7.0 with the chaotrope sodium trichloroacetate (71)and subjected to repeated freeze-thawing a t 77OK (liquid N,) and room temperature, respectively. The larger subunit formed an insoluble aggregate, while the smaller subunit remained in solution. They were separated by centrifugation. Figure 26 shows the effect of increasing concentrations of the sodium salts of trichloroacetate (TCA) , SCN-, ClO,-, and of guanidinium hydrochloride (Gu-HCl) on the resolution of succinate dehydrogenase as measured by the loss of PMS reductase activity. It is seen that, as expected, the extent of resolution increases in a cooperative manner with the increased concentration of chaotropes, and that addition of high concentrations of chaotropes without freeze-thawing had no effect on PMS reductase activity (Fig. 26, top right corner). The subunits separated by the above technique indicated a distribution of iron and labile sulfide as shown in Table X. Thus the larger subunit contained flavin, iron, and labile sulfide in the approximate ratio of 1 :4 :4, while the smaller subunit appeared to have the characteristics of a soluble iron-sulfur protein. The absorption spectra of succinate dehydrogenase and its two subunits analyzed to show the contributions of flavin and the iron-sulfur chromophore in each preparation are given in Fig. 27. c. Molecular Weight. Since flavin is covalently bound in succinate

0.1

0.2

0.3

0.4

0.5

Cawntration [MI

FIQ.26. Effect of chaotropic agents and freeze-thawing on the resolution of succinate dehydrogenase. The enzyme, at a protein concentration of 1.48 mg/ml in 50 mM sodium phosphate (pH 7.0) and 20 m M succinate, was treated with various chaotropic agents as shown in the figure, and frozen in liquid nitrogen and thawed at room temperature three times. Since in the presence of higher concentrations of chaotropes the larger subunit precipitated after freeze-thawing, all samples were centrifuged for 1 min at 35,000 rpm, and only the clear supernatant was assayed for PMS reductase activity. F-T,freeze-thawing. From Hanstein et al. (166).

4.

233

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

TABLE X MOLECULAR

PROPERTIES O F

SUCCINATE DEHYDROGENASE' Flavin

Molecule

Molecular weight

Succinate dehydrogenase Large subunit Small subunit

97,000 f 4% 70,000 k 7% 27,000 k 5%

Iron

Labile sulfide

nmoles (ng-atoms)/mg protein 10. 3b 12-13 < 0.5"

70-80 45-55 95-110

70-80 45-55 90-100

From Davis and Hatefi (143). 12 preparations. c From the small subunit obtained by chaotrope-induced resolution of succinate dehydrogenase ( 1 4 3 ) . The small subunit obtained by resolution and column chromatography in presence of SDS is completely free of flavin. a

* Average of

Wavelength (nm)

(A)

(6)

(C)

FIG.27. Absorption spectra of succinate dehydrogenase (A), its larger subunit (B), and its smaller subunit (C). ( A ) Trace 1, oxidized succinate dehydrogenase; trace 2, flavin contribution to the spectrum of oxidized enzyme; trace 5, iron-sulfur contribution to the spectrum of oxidized enzyme ; trace 4, oxidized enzyme treated with dithionite ; trace 3, oxidized enzyme treated with sodium mersalyl and dithionite. Protein = 1.48 mg/ml. (B) Trace 1, oxidized; trace 2, flavin contribution to trace 1 ; trace 3, after treatment of 1 with sodium mersalyl and dithionite. Protein = 3.0 mg/ml. (C) Trace 1, oxidized; trace 2, after treatment with mersalyl; trace 3, after treatment of 2 with dithionite. Protein = 0.8 mg,'ml. From Davis and Hatefi ( 1 4 3 ) .

234

YOUSSEF HATEFI AND DIANA L. STIGGALL

dehydrogenase, its concentration in a preparation would permit the calculation of a minimum molecular weight, The flavin content of succinate dehydrogenase as reported by Davis and Hatefi (143) for a number of preparations is 10.3 k 4% nmoles/mg protein. This value indicates a minimum molecular weight of 97,000 f 4%, which agrees with the sum of the subunit molecular weights (70,000 & 7% plus 27,000 f 5% = 97,000 6.6%). The relative mobility of succinate dehydrogenase on Agarose columns corresponded to a molecular weight of 105,000 k 7.6%. The molecular weight of about 100,000 for succinate dehydrogenase was confirmed subsequently by others (25, 158) for the enzyme isolated from complex I1 by the method of Davis and Hatefi. The above data also suggested that the ratio of the two subunits in succinate dehydrogenase must be close to unity. According to Coles et al. (158) and Righetti and Cerletti (1573, their preparations contain more of the small subunit. This has not been confirmed by others, but a 2 : 1 ratio in favor of the smaller subunit (or an impurity which would coelectrophorese with the smaller subunit) would account for the low flavin content (7-8 nmoles/mg protein) reported by the latter authors. The sedimentation constant (s2,) of succinate dehydrogenase between 4 and 12 mg protein per ml was determined by Davis and Hatefi (143) to be 9.22 S. This is considerably higher than the value of 6.5 S given by Singer (15) and co-workers for their low iron, 50% pure preparation. The high sedimentation constant was considered by Davis and Hatefi to be either the result of the density and shape of the enzyme (159b) or polymerization in solution. Ultracentrifugation of the enzyme in the presence of potent depolymerizing agents (SDS, cetyldimethylethylammonium bromide, Triton X-100, or 0.5 M sodium trichloroacetate) a t pH 7.0, 8.0, and 9.0 showed no change in the sedimentation constant. These results and the relative mobility of succinate dehydrogenase on Agarose columns did not agree with the polymerization possibility. HOWever, Coles et al. (158) felt that succinate dehydrogenase purified according to Davis and Hatefi exists in solution in a monomer-dimer equilibrium, and that a t protein concentrations of 1-10 mg/ml the molecular weight obtained by sucrose gradient centrifugation and chromatography on Sephadex G-200 is 175,000 & 10,000. d. Prosthetic Groups. Succinate dehydrogenase contains one mole of FAD per mole, and the flavin is covalently linked to the larger subunit

*

159b. D-Amino acid oxidase (Michaelis complex with benzoate) has been reported ( f / f ~ ) of 1.013

to have a molecular weight of 115,000 k 500, a frictional coefficient (nearly spherical), and an S = 11.00 (160).

160. K . Yagi and T. Ozawa, BBA 62, 397 (1962).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

235

FIG.28. Structure of histidyl-8-a-FAD (left) and the sequence of the flavin pentapeptide (right) of succinate dehydrogenase. From Singer et al. ( 2 6 ) . (143). As a result of collaborative work among the laboratories of Ehrenberg, Hemmerich, arid Singer (161-163), the structure of the bound flavin was elucidated to be 8 ~(N-3-histidyl) flavin adenine dinucleotide. The structure of the histidyl flavin and the sequence of a flavin-bound pentapeptide isolated by proteolytic digestion are shown in Fig. 28. Singer et al. (23,25) have discussed the details of these studies. I t has been shown recently by Ohnishi et al. (164) that a t temperatures below 20°K, succinate dehydrogcnase or complex I1 exhibits E P R signals indicative of two iron-sulfur centers, designated centers S-1 and S-2. Center S-1 ( g = 2.03, 1.94, and 1.92 a t temperatures above 20°K) is the same as that originally reported by Beinert and Sands (1666).Center S-2 exhibits signals a t the same field positions but with different line shapes. These signals become prominent at temperatures below 20°K. The halfreduction potentials (En&)given for these two centers a t pH 7.4 are 0 +- 10 and -260 +- 15 mV, respectively. The presence of two species of EPR-active iron-sulfur centers in succinate dehydrogenase agrees with the earlier finding of two iron-sulfur containing subunits. In the larger subunit the iron : labile sulfide: flavin ratio suggests that this subunit might contain a 4 iron-4 labile sulfide cluster as in clostridial ferredoxin. In the smaller subunit, the data of Davis and Hatefi indicate the presence of approximately 3 g-atoms of iron and 3 moles of labile sulfide per mole. Since these values are clearly higher than 2 and the loss of iron and labile sulfide as a result of resolution of the enzyme into subunits is quite prob161. P. Hemmerich, A. Ehrenbcrg, W. H . Walker, L. E . G . Eriksson, J. Salach, P. Bader, and T. P. Singer, FEBS ( F e d . Eur. Biochem. Soc.) L e t t . 3, 37 (1969). 162. J. Salach, W. H. Walker, T. P. Singer, A. Ehrenberg, P. Hemmerich, S. Ghisla, and U. Hartman, Eur. J. Biochem. 26, 267 (1972). 163. T. P. Singer, J. Salach, W. H. Walker, M. Gutman, P. Hemmerich, and A. Ehrenherg, i n “Flavins and Flavoproteins,” 3rd Int. Symp. (H. Kamin, ed.), p. 607. Univ. Park Press, Baltimore, Maryland, 1971. 164. T. Ohnishi, D. U. Winter, J. Lim, and T. E. King, BBRC 53, 231 (1973). 165. H. Beinert and R. H. Sands, BBRC 3, 41 (1960).

236

YOUSSEF HATEFI AND DIANA L. STIGGALL

TABLE XI DYE REDUCTASE PROPERTIES OF SUCCINATE DEHYDROOENASE Parameter Succinate -+ PMSa activity Succinate -+K3Fe(CN)aactivity Turnover number at Vz::

::v

KEMs(mM)

K z c o(mM) 0

Value 67-7gb 13. 5bsc 10,000-11,000d 100-1 l o b -0.48 ~ 0 . 3 ~

At 1.65 mM PMS.

* Micromoles of succinate oxidized X min-1

X mg-I protein at 38". At 3 mM ferricyanide. d Moles of succinate oxidized X min-1 X mole-' enzyme at 38".

able, the smaller subunit might also have a clostridial-type iron-sulfur center .

2. Enzymic Properties a . Succinate Dehydrogenase. Soluble succinate dehydrogenase can oxidize succinate in the presence of PMS or ferricyanide as electron acceptor. The highest activities reported for various preparations of succinate dehydrogenase are those of the preparation of Davis and Hatefi (143, 166). These values are shown in Table XI. The turnover number of the enzyme a t V,,, with respect to PMS concentration is 10,000-11,000 moles succinate oxidized per minute per mole of enzyme at 3 8 O . This value is the same as the turnover number of complex I1 (per mole of flavin) with either PMS or ubiquinone as electron acceptor (153, 154, 156). It is also the same as the turnover number reported by Ziegler and Doeg (153) and by King (167) for succinate oxidation by submitochondrial particles. The values given by the former authors are 10,000 with ubiquinone, 9,700 with PMS, and 10,100 with oxygen as electron acceptors. According to King, the turnover numbers of the succinoxidase system of intact heart muscle preparations and reconstituted preparations are 7,500 and 10,000, respectively. The latter value is the same as that given by Veeger et al. (149) for reconstituted particles made with their preparation of succinate dehydrogenase. In contrast to the consistent results of the above authors, Singer (15,23,25) insisted that the turnover number of succinate dehydrogenase in heart mitochondria is 17,000-18,000, and concluded that 166. W. G. Hanstein, K. A. Davis, M. A. Ghalambor, and Y . Hatefi, Biochemistry 10, 2517 (1971). 167. T. E. King, Advan. Enzymol. 28, 155 (1966).

4. METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

237

complex I1 and soluble succinate dehydrogenases have, therefore, lost activity as a result of preparative modification. Surprisingly, however, the paper referred to by Singer for the high turnover number of 17,00018,000 is also by Singer et al. (168), and in the latter paper the turnover number of succinate dehydrogenase in beef heart mitochondria a t V,,, with respect to PMS and 3 8 O is 11,000-13,000 and in complex I1 it is 11,300-11,900 moles succinate oxidized per mole of bound flavin. The only other high turnover number found in the literature is that of Cerletti et al. (169) which does not give details. These stated differences might also be based on the values used for the concentration of succinate dehydrogenase flavin in heart mitochondria and respiratory particles. Neither Singer et al. (168) nor Cerletti et al. (169) gave the flavin values used in these papers for the calculation of their turnover numbers. However, literature values for submitochondrial particles range from 0.11-0.14 (15) to 0.28 (170) nmole/mg protein. The ferricyanide reductase activity given in Table XI was measured in the presence of 3 mM ferricyanide and is far from V,,,. As shown by King (17 1 ) ,higher ferricyanide concentrations are strongly inhibitory. Others have stated, however, that the extrapolated turnover number of the enzyme is the same with either PMS or ferricyanide as electron acceptor (172).The K , values given in Table XI for PMS and succinate are essentially the same as the values reported by King (167) and King and Takemori (173).The K, for succinate is also the same as has been found for complex 11, submitochondrial particles (166),Keilin-Har,tree heart muscle preparations, and King's reconstitutable enzyme ( 1 7 3 ) .The value given by Singer (15) for the succinate-PMS reductase activity of heart homogenates, determined under similar temperature ( 3 8 O ) and pH conditions as in Table XI, is 1.3 mM. The I(, given by Singer et al. (25) for succinate a t 2 2 O to 2 5 O is 0.5 mM. According to Veeger et al. (149),succinate dehydrogenase can also oxidize L-chlorosuccinate, L-methyl succinate, D-malate, and L-malate. Brodie and Nicholls (174) have reported that monofluorosuccinate and 2,2-difluorosuccinate are also oxidized by Keilin-Hartree preparations yielding oxaloacetate as a final product. D-Chlorosuccinate, D-methyl suc168. T. P. Singer, J. Hauber, and 0. Arrigoni, BBRC 9, 150 (1962). 169. P. Cerletti, R. Strom, M. Giordano, F. Balastero, and M. A. Giovenco, BBRC 14, 408 (1963). 170. P. V. Blair, T. Oda, and D. E. Green, Biochemistry 2, 756 (1963). 171. T. E. King, JBC 238, 4032 (1963). 172. W. P. Zeylemaker, A. D. M. Klaase, E. C. Slater, and C . Veeger, BBA 198, 415 (1970). 173. T. E. King and S. Takemori, JBC 239,3559 (19643. 174. J. D. Brodie and P. Nicholls. BBA 198,423 (1970).

238

YOUSSEF HATEFI AND DIANA L. STIGGALL

TABLE XI1 DISSOCIATION AND INHIBITION CONSTANTS OF COMPETITIVE INHIBITORS OF SUCCINATE DEHYDROOENASE"

Inhibitor Oxaloacetate Malonate Fumarate Methylene succinate Maleate Bicarbonate Glyoxylate Acetoacetate Glycolate Formate 0

KD (mM)

Ki (mM)

0.004 0.028 2.6 6.3 7.0

0.0015 0.018 1.3 1.8 6.2 12 21 40 120 120

80 -

From Zeylemaker et al. (I"$?) and DerVartanian and Veeger (176).

cinate, malonate, methylene succinate, maleate, acetoacetate, and oxaloacetate are competitive inhibitors (149). I n addition bicarbonate, formate, glycolate, and glyoxylate are also competitive inhibitors, and appear to bind a t two sites on the enzyme (172).The inhibitor constants ( K i ) and the dissociation constants ( K D ) of a number of these compounds as reported by Veeger and his colleagues (172, 175) are shown in Table XII. It is seen that among these oxaloacetate binds strongly to the enzyme and is a very potent competitive inhibitor. The role of oxaloacetate in controlling the activity of succinate dehydrogenase is discussed below. DerVartanian and Veeger ( 1 75,176) have studied the effect of various substrates, inhibitors and analogs on the spectral properties of their preparation of succinate dehydrogenase. Spectral changes in the presence of competitive inhibitors are thought to arise from charge-transfer complex formation between the electron-donating inhibitor and the enzyme. Both the membrane-bound and the soluble succinate dehydrogenases are capable of catalyzing fumarate reduction in the presence of a suitable electron donor such as FMNH,. I n the mammalian enzyme, this activity is not more than a few percent of the succinate-PMS reductase activity (23, 2 5 ) . Ringler and Singer (177) have also shown that in brain mito175. D. V. DerVartanian and C. Veeger, BBA 92,233 (1964). 176. C. Veeger, D. V. DerVartanian, J. F. Kalse, A. de Kok, and J. F. Koster, in "Flavins and Flavoproteins," Symp Adn Flavoproteins (E.C. Slater, ed.), p. 242. Elsevier, Amsterdam, 1966. 177. R. L. Ringler and T. P. Singer, JBC 234,2211 (1959).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

239

chondria treated with antimycin A, the reduction of fumarate to succinate could be linked to the oxidation of a-glycerophosphate to dihydroxyacetone phosphate. A seemingly interesting observation reported by Singer is that alkali treatment of respiratory particles a t pH 9.3-9.4 results in a much greater loss of succinate dehydrogenase activity than fumarate reductase activity. However, i t should be remembered that the electron donors and acceptors in these assays are not exactly the same. Therefore, the difference might have resulted from preferential destruction by alkali of the PMS reductase site. b. Complex ZZ. Preparations of complex I1 catalyze the oxidation of succinate by either PMS ( a t V,n,,) or ubiquinone a t a rate of about 45-55 pmoles/min/mg protein a t 38O. Thus the turnover number in either reaction is about 10,000. It has been shown by Ziegler and Doeg (153) that the PMS reductase activity of complex I1 is the same in the presence or absence of added ubiquinone-2 (complex I1 preparations are essentially devoid of bound ubiquinone). This is not in agreement with the conclusions of others on ubiquinone effect in submitochondrial particles (see, however, Section III,A,4). It has been shown recently that the mitochondria1 electron transport system contains a t least three different b-type cytochromes (178).Two of these cytochromes are found in complex 111, and under appropriate conditions are reducible with substrates. The third b-type cytochrome was discovered by Davis et al. ( l 7 8 ) , and shown to fractionate exclusively into complex 11. At 77OK, the cytochrome b of complex I1 exhibits a double band a t 557.5 and 550 nm, a prominent p band a t 531 nm, and a Soret band at 422 nm (Fig. 29). Cytochrome b,,,., appears to have a low reduction potential. It is not detectably reduced by succinate in either complex I1 or respiratory particles, but its dithionite reduced form is rapidly oxidized by either fumarate or ubiquinone. The role of this cytochrome in mammalian mitochondria is not known. Davis et al. (178) have suggested that it might be an electron entry point for an unknown ancillary tributary of the respiratory chain. Further, Bruni and Racker (179) have shown that a preparation of cytochrome b is required for reconstitution of succinate-ubiquinone reductase activity (see below). c. Reconstitution. The reconstitution experiments involving succinate dehydrogenase may be divided into three types. These are recombination of succinate dehydrogenase with alkali-inactivated particles, reconstitution of succinate-ubiquinone reductase activity with partially purified components, and reversible resolution of complex 11. (Y

178. K. A. Davis, Y . Hatefi, K. L. Poff, and W. L. Butler, BBA 325,341 (1973). 179. A. Bruni and E. Racker, JBC 243,962 (1968).

240

YOUSSEF H A T E F I AND DIANA L. STIGGALL

I

400 420

440

460 400 500 Wavelength, nm

520

540

560

FIQ.29. Reduced minus oxidized spectrum of cytochrome bEnr.sin complex 11. Complex I1 at 1.76 mg/ml of 40 mM potassium phosphate, pH 7.4, was treated with a small amount of dithionite in order to reduce its succinate dehydrogenase and minor (-10%) complex I11 contaminant, and its spectrum was recorded. This spectrum was then subtracted from that of complex I1 fully reduced with dithionite. From Davis et a.!. (f78).

It has been known since the early 1950’s that treatment of respiratory particles a t alkaline pH ( >9.0) destroys succinoxidase activity. King and his colleagues showed that addition of the Keilin-King type of succinate dehydrogenase to these particles restored a substantial amount of succinoxidase activity (147, 148, 167). The alkali inactivation of the particles was shown subsequently to result from inactivation of the particlebound enzyme (16, 166, 180) rather than loss of succinate dehydrogenase as originally believed (148, 167). The work of Hanstein et a2. (166) with pure succinate dehydrogenase clarified many aspects of this type of reconstitution. Their results may be summarized as follows. Incubation of submitochondrial particles a t pH 9.3 and 38O under an argon atmosphere resulted in a rapid, exponential loss of succinoxidase activity. Presence of succinate during incubation retarded inactivation considerably, and concomitant presence of dithiothreitol offered additional protection (Fig. 30A). All the decay curves extrapolated to a zero time activity corresponding to 90% ,of the activity of untreated particles (Fig. 30A). This 90% activity was totally recovered upon addition of sufficient amounts of succinate dehydrogenase to the alkali-inactivated particles. Titration 180. T. Kimura, J. Hauber, and T. P. Singer, Nature (London) 198,362 (1968).

4.

241

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

Min at pH 9.3 and 38'

Min ot 0'

(A)

(8)

FIG.30. (A) Kinetics of inactivation of succinate oxidase activity of submitochondrial particles (ETP) a t pH 9.3 and 38". E T P at 18.2 mg/ml of deoxygenated 035 M sucrose plus 10 mM Tris-HC1 (pH 8.0) was brought to pH 9.3 at 38" with a predetermined amount of 1 N NaOH, incubated under an atmosphere of argon and sampled for activity as indicated. Where present in the incubation mixture, succinate was 20 mM and dithiothreitol (DTT) 5 mM. Succinoxidase assays were conducted at 30". All samples were neutralized with one volume of 100 mM sodium phosphate (pH 7.3), containing 20 mM succinate and incubated 3 min at 38" before assay. To those not containing succinate in the original mixture 40 mM succinate was added along with phosphate buffer at the time of neutralization, so that the final succinate concentration in all samples was 20 mM. From Hanstein et al. (166). (B) Loss of reconstitution activity of soluble succinate dehydrogenase at 0" as a function of time as compared to the activity of the same preparation when bound to alkali-treated submitochondrial particles (ETP). Succinoxidase activity was asNo ETP. sayed as in ( A ) . (U) SD/ETP = 0.10, ( A ) SD/ETP = 0.30,(0)

of the particles with increasing amounts of succinate dehydrogenase (Fig. 31) indicated that full activity restoration required the addition of a t least 5 times as much succinate dehydrogenase as was present in the particles. The same stoichiometry was found for reconstitution of activity in alkali-treated complex 11. These results suggest that the equilibrium of the system involving free and particle-bound succinate dehydrogenase under these conditions (i.e., in the presence of alkali-treated particles) is in favor of the free enzyme. Another possibility that cannot be ruled out is that only 20% of the added succinate dehydrogenase was capable of reconstitution. When succinate dehydrogenase was added to alkaliinactivated complex I1 and the reconstituted complex isolated by centrifugation, it was found that the active complex contained twice as much flavin and the corresponding amount of additional protein (Table XIII). These results showed, therefore, that in this type of reconstitution a double-headed particle is formed containing one complement of active

242

YOUSSEF HATEFI AND DIANA L. STIGGALL SD/alk-ETP

Ql

t

1.6

'

0;2

,

0:3

. $

1.2-

aa

b

tI : 0.8

F

M S.A./mg alk-ETPprole S.A./w SDpotsin 0-0 S.A./mq total protein

-

tP 0."4i

I 8

.

'

16

, I

24

alk- ETP/SD

Fra. 31. Reconstitution of succinate oxidase activity. Fresh succinate dehydrogenase was dissolved at a concentration of 7.8 mg/ml in 50 mM phosphate (pH 7.5), containing 20 mM succinate and 5 mM dithiothreitol, then immediately divided in 50 pl quantities into small test tubes, frozen in liquid nitrogen, and stored at -70". E T P preparations stored as centrifuged pellets were suspended a t a concentration of 9.1 mg/ml in 50 mM Tris (pH 8.01, containing 0.66 M sucrose. They were inactivated a t pH 9 8 essentially as in Fig. 30A (none), readjusted to pH 7.7,and supplied with 20 mM succinate (protein concentration at this point was 8.35 mg/ml). Appropriate amounts of the alkali-treated E T P were then added to test tubes containing frozen succinate dehydrogenase. The components were mixed a t room temperature with the help of a stirring rod, incubated 2.5 min at 30°, and assayed at 30" for succinate oxidase activity. alk-ETP, alkali-treated ETP. From Hanstein et al. (166).

(added) and one of inactive (alkali-treated) succinate dehydrogenase (166).

As originally shown by King (148),the reconstitution, but not the PMS reductase, activity of succinate dehydrogenase is extremely labile. Hanstein et al. (166) showed that at Oo the reconstitution activity of the soluble, but not the reconstituted, enzyme decays exponentially with a half-life of 75 min (Fig. 30B). However, a t < - 6 5 O this activity was indefinitely stable. King (148) showed that during extraction of the enzyme from particles the presence of succinate was necessary for retention of reconstitution activity. Davis and Hatefi (149) used both succinate and dithiothreitol during the chaotrope-induced resolution of complex I1 and salt precipitation of succinate dehydrogenase. They showed, however, that dithiothreitol was much more effective than succinate for complete preservation of reconstitution and PMS reductase activities (166). King (148, 161) had shown that except for reconstitution activity succinate dehydrogenases extracted in the presence and absence of succinate were identical in composition, spectra, and dye reductase properties. However,

4.

243

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

RECONsTlTUTION

Preparation Complex 11, p H 9.3, 0 rnin a t 38" Complex 11, p H 9.3, 20 min a t 38" Complex I1 succinate dehydrogenase Complex I1 succinate dehydrogenase, spun 60 min Supernatant Pellet

+ +

OF

TABLE XI11 SUCCIN~4T~-UnIaUINONE REDUCT.4SE ACTIVITY"

Protein (mg)

Flavin (nmoles / me)

Succ' PMS (per mg total protein)

Succ-t Q (per mg total protein)

22.6

22.2

5.6

5.6

5.6

4.92

<0.9

<0.8

+ 12.6

8.Gb

43.0

7.7

6.7

48.8 24.4

<1.0 23.9

10.0 7.9

a Conditions: Complex I1 was inactivated by 20 min incubation a t pH 9.3 and readjusted to p H 7.6 (protein 11.25 mg/ml). Succinate dehydrogenase was prepared and stored as an ammonium sulfate-precipitated pellet for 1 day a t -70". I t was dissolved in 50 mM Tris-HC1 (pH 8.0) containing 20 m M succinate and 3 m M dithiothreitol before using (protein 21.1 mg/ml). Alkali-treated complex I1 (0.5 ml) and succinate dehydrogenase (0.6 ml) were mixed together, assayed, and centrifuged for 60 min a t 49,000 rpm. The supernatant and the pellet were then separated, the latter was suspended in Tris-succinate-dithiothreitol buffer and both fractions were assayed as indicated. Activities shown are expressed as micromoles of succinate oxidized X min-1 x mg-1 of total protein a t 38". * Calculated. From Hanstein el al. (168).

Baginsky and Hatefi (155, 156) showed that loss of reconstitution activity appears to be related to a damage in the iron-sulfur system of the enzyme which is not detectable by assay for iron and labile sulfide content. They obtained a preparation of succinate dehydrogenase from complex I1 which exhibited no reconstitution activity but had an iron:labile su1fide:flavin ratio close to 8:8:1. They were then able to reactivate this enzyme for reconstitution by treating it with Na2S, ferrous ions, and mercaptoethanol, essentially in the same manner as apoferredoxin had been previously converted to ferredoxin (181, 182). The reactivated preparation was able to reconstitute with alkali-treated submitochondrial particles or complex 11. Analyses showed that the preparation had acquired additional iron and labile sulfide, but control experiments indicated that reconstitution activity was not a spurious effect. The reactiva181. J. Hong and J. C. Rabinowitz, BBRC 29,246 (1967). 182. K.McCarthy and W. Lovenberg, JBC 243,6439 (1968).

244

YOUSSEF HATEFI AND DIANA L. STIGGALL

tion of succinate dehydrogenase by the above technique was confirmed by others (182a). As stated above, preparations of succinate dehydrogenase contain two EPR-active iron-sulfur centers in agreement with the fact that the enzyme is composed of two iron-sulfur containing subunits. The temperature-insensitive E P R signal a t g = 1.94 (EmN 0 mV) is present in high iron-labile sulfide preparations which are inactive (156) and active (143) for reconstitution. I n addition, this signal was originally observed by Beinert and Sands (165) in succinate dehydrogenase preparations of Singer et al. and Basford et aZ., which according to Beinert and Sands contained per mg protein 1.0 nmole flavin plus 4.7 ng-atoms iron and 4 nmoles flavin plus 8 ng-atoms iron, respectively. These results indicate that (a) reconstitution activity may not be related to the g = 1.94signal, and (b) that the signal is present in preparations with iron :flavin ratios of 4:l and 2:l. The latter conclusion is only qualitatively correct, of course, since titration and signal integration were not performed in these early experiments, and possible heterogeneity in the enzyme preparations tested cannot be ruled out. However, Ohnishi et al. (164) stated that the stability of center S-1 mirrors the stability of the dye reductase activity of the enzyme, whereas the half-reduction potential of center S-2 in soluble succinate dehydrogenase (King type) showed a strong tendency to decrease from -260 mV to approximately -430 mV. They felt that these results suggest that center 5-2 may be involved in linking the enzyme to the respiratory chain. This conclusion does not necessarily imply that center 5-2 is an intermediate electron carrier since succinate does not reduce this low potential iron-sulfur center. The second type of reconstitution is demonstrated by the work of Bruni and Racker (179).These investigators reconstituted a succinate-ubiquinone reductase system from a King-type preparation of succinate dehydrogenase (2.6nmoles flavin per mg protein), a preparation of cytochrome b (24-27nmoles heme per mg protein) solubilized and purified with the use of bile salts and SDS, and mitochondria1 or soybean phospholipids. The highest succinate-ubiquinone reductase activity achieved was 980 moles succinate oxidized per minute per mole of succinate dehydrogenase flavin. While this activity is only 10% of the turnover number of complex 11, it is still quite appreciable for this type of reconstitution. Since the preparations of succinate dehydrogenase and cytochrome b used were not pure, it is not known what is the minimum number of components needed for reconstitution of succinate-ubiquinone reductase activity. The role and the exact nature of the b-type cytochrome used in these experi182a. T. E. King, D. Winter, and W. Steele, in “Structure and Function of Oxidation Reduction Enzymes” (A. Akeson and A. Ehrenberg, eds.), p. 519. Pergamon, Oxford, 1972.

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGJCNASES

245

ments are also not known (in complex I1 the ratio of cytochrome b to flavin is approximately 1 :1; in the Bruni-Racker reconstituted preparation this ratio was 3 2 : l ) . On the other hand, there is no evidence for the existence of a component in complex I1 which might act as an intermediate electron carrier between succinate dehydrogenase and ubiquinone. It is entirely possible that, as implied by Bruni and Racker, succinate dehydrogenase bound to a membranous structure containing phospholipids might be able to interact with ubiquinone. Recently, Beinert and his colleagues (183, 184) have isolated an iron-sulfur protein from mitochondria with a molecular weight of about 100,000. They stated that preparations of complex I1 contain an iron-sulfur component with similar EPR characteristics. However, they agreed with the previous work of others (143) that the SDS-polyacrylamide gel pattern of complex I1 is devoid of a species with the mobility characteristics of their 100,000 molecular weight iron-sulfur protein. The third type of reconstitution has been discussed already in Section III,A,l,a. It involves reversal of the chaotrope-induced resolution of complex I1 with respect to succinate dehydrogenase either by removal (or dilution) of the added chaotrope or by addition of an antichaotropic ion to the resolved system. The basis of this resolution-reconstitution process is the disordering and restructuring of the water in which complex I1 is suspended. The process in either direction is relatively rapid, and complete structural and functional reconstitution is easily achieved (Figs. 25a and b). Earlier studies of Hatefi et al. ($7, 29) have also shown that complex I1 interacts with complexes I, 111, and IV to reconstitute the entire respiratory chain. However, contrary to claims in the literature (167, 173) based on experiments in which highly impure preparations of succinate dehydrogenase and complex I11 were used, pure succinate dehydrogenase and complex I11 do not interact to reconstitute a succinate-cytochrome c reductase system (185). These results indicate that appropriate components of complex I1 are needed t o link succinate dehydrogenase to the mitochondria1 ubiquinone-cytochrome c reductaee complex.

3. Inhibitors and Modifiers It has been known since the work of Hopkins and his colleagues in 1938 (186) that succinate dehydrogenase contains -SH groups essential for the catalytic activity of the enzyme, and that substrates and competitive inhibitors protect the enzyme against inhibition by -SH reagents. 183. F. J. Rueicka and H. Beinert, BBRC 58,556 (1974). 184. H. Beinert, B. A. C. Ackrell, E. B. Kearney, and T. P. Singer, BBRC 58, 564 (1974). 185. K. A. Davis and Y. Hatefi, BBRC 44, 1338 (1971). 186. F. G. Hopkins, E. Morgan, and C. Lutwak-Mann, BJ 32, 1829 (1938).

246

YOUSSEF H A T E F I AND DIANA L. STIGGALL

However, it is not known whether the active thiol of succinate dehydrogenase is a t or near the enzyme active site. At low levels of mercurials, the inhibition can be reversed, but a t high levels mercurials irreversibly destroy the iron-sulfur chromophore of the enzyme (149, 187).The flavin spectra of succinate dehydrogenase and its larger subunit in Fig. 27 were obtained by treating the preparations with solid sodium mersalyl. I n addition to mercurials, N-ethylmaleimide and certain alkyl bromides also inhibit the enzyme. Similar to other iron-sulfur proteins, succinate dehydrogenase is slowly inactivated in the presence of o-phenanthroline, but Tiron and bathophenanthroline are much less effective (166). 2-Thenoyltrifluoroacetone (TTFA), which is also considered to interact with iron in the respiratory chain, is a strong and relatively specific inhibitor of succinate-ubiquinone reductase activity, but does not inhibit PMS reductase activity in complex I1 and soluble succinate dehydrogenase. Since in complex I1 and the reconstituted system of Bruni and Racker (179) the only succinate-reducible species known are succinate dehydrogenase and ubiquinone, Baginsky and Hatefi (156) have suggested that the site of action of TTFA might be on succinate dehydrogenase itself. I n view of our present knowledge regarding the two iron-sulfur-containing subunits of the enzyme, it is further possible that PMS and TTFA do not interact with the same iron-sulfur species. According to Rossi et al. (188) TTFA also inhibits the succinate dehydrogenase activity of submitochondrial particles. However, these data involve assay complications, which will be discussed in Section 111,A14. Since the isolation of succinate dehydrogenase from cyanide-treated particles (146), a considerable amount of work has been done on the effect of cyanide on succinate dehydrogenase. Treatment of particles with cyanide diminishes succinate-PMS reductase activity by about 50% and causes a large increase in the apparent I<,, of the dye. These effects are prevented by succinate (189).The effect of cyanide is more drastic on electron transfer from succinate to methylene blue or the respiratory chain. These reactions are almost completely destroyed by cyanide. Experiments with partially purified succinate dehydrogenase showed, however, that cyanide treatment did not alter the PMS reductase activity (190). The reactive species has been shown to be CN-, which binds strongly to succinate dehydrogenase (189-1 91). According to Zanetti et 187. V. Massey, BBA 30, 500 (1958). 188. E. Rossi, B. Norling, B. Persson, and L. Ernster, Eur.

J. Biochem. 16, 508 (1970). 189. A. Giuditta and T. P. Singer, JBC 234, 666 (1959). 190. C.-P. Lee and T. E. King, BBA 59,716 (1962). 191. G. Zanetti, Y. M. Galante, P. Arosio, and P. Cerletti, BBA 321, 41 (1973).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

247

al. (191) the stoichiometry is 2 moles of CN- per g-atom of iron. It has been postulated that interaction of cyanide with succinate dehydrogenase causes conformational changes which alter the dye reductase properties of the enzyme and labilize its linkage to the respiratory chain (189, 191). Other details of the cyanide effect are found in earlier reviews (15, 251, and will be discussed below. 4. Regulatory Properties

It has been known since the early studies of Kearney (192) that SUCcinate dehydrogenase undergoes reversible activation by substrates, competitive inhibitors, and phosphate. The activation of succinate dehydrogenase was shown to be a characteristic of both the soluble and particle-bound enzyme and a slow process requiring many minutes of incubation with the activator a t ambient or higher temperatures (activation energy = 31-33 kcal/mole). It has been suggested that the enzyme exists in a free equilibrium between the unactivated and the activated forms, and that the activator interacts with the latter and establishes a new equilibrium in favor of the activated state of the enzyme (23, 25, 193; see also 194 for an expanded mechanism). I n addition to succinate, malonate, fumarate, and phosphate, i t has been reported in recent years that succinate dehydrogenase is activated by ATP, ITP, IDP (195), reduced ubiquinone-10 (195-197), succinyl coenzyme-A (198), formate, ClO,-, I-, Br-, C1-, NO,-, SO,2-, acid pH ( I % ) , FMNH,, light irradiation of enzyme in the presence of EDTA (200), 2,4-dinitrophenol (201), and phospholipids (202-205; see also 20, 192. E. B. Kearney, JBC 229,363 (1957). 193. T. Kimura, J. Hauber, and T. P. Singer, JBC 242,4987 (1967). 194. R. G. McDonald-Gibson and M. B. Thorn, BJ 114,755 (1969). 195. M. Gutman, E. B. Kearney, and T. P. Singer, Biochemistry 10, 4763 (1971). 196. M. Gutman, E. B. Kearney, and T. P. Singer, BBRC 42, 1016 (1971). 197. M. Gutman, E. B. Kearney, and T. P. Singer, Biochemistry 10, 2726 (1971). 198. E. B. Kearney, M. Mayr, and T. P. Singer, BBRC 46, 531 (1972). 199. E. B. Kearney, B. A. C. Ackrell, M. Mayr, and T. P. Singer, JBC 249, 2016 (1974). 200. J. I. Salach and T. P. Singer, JBC 249,3765 (1974). 201. L. Susheela and T. Ramasarma, BBA 242, 532 (1971). 202. P. Cerletti, R. Strom, and M. G. Giordano, BBRC 18, 259 (1965). 203. P. Cerletti, M. A. Giovenco, M. G. Giordano, S. Giovenco, and R. Strom, BBA 146, 380 (1967). 204. P. Cerletti, P. Caifa, M. G. Giordano, and G. Magni, in “Flavins and Flavoproteins,” 2nd Int. Symp. (K. Yagi, ed.), p. 178. Univ. Park Press, Baltimore, Maryland, 1968. 205. P. Cerletti, P. Caifa, M. G. Giordano, and M. A. Giovenco, BBA 191, 502 (1969).

248

YOUSSEF HATEFI AND DIANA L. STIGGALL

24, 26, 206-208). Among the nucleotides ATP does not activate complex

I1 and soluble preparations, while ITP and IDP do. IDP is much more effective than ITP, but its effect relative to ATP could not be tested because it does not readily penetrate the mitochondria1 membrane. The activating effect of formate is probably related to its property as a competitive inhibitor (see above). The order of effectiveness of the monovalent ions has been reported to be C10,-, I-, NO,- > Br- > C1-, which is reminiscent of their order of potency as chaotropes (70-72). These ions are effective a t much lower concentrations when used a t pH values a t or below pH 6. It was reported by Hanstein et al. (166) that pure succinate dehydrogenase prepared by perchlorate extraction of complex I1 in the presence of succinate and dithiothreitol was fully activated. However, when reconstituted with alkali-inactivated submitochondrial particles the enzyme became deactivated and required preincubation a t 30° with succinate before maximal succinoxidase activity was attained. Ignoring the latter observation, Singer and his colleagues chose to fault the pure enzyme for being fully active as isolated on the assumed ground that it had lost its regulatory properties (23, 26, 26, 168). Subsequent studies of Coles et al. (209) showed, however, that the above preparation could be deactivated by removal of succinate and perchlorate. When isolated from complex I1 in the absence of succinate (as stated above succinate is not crucial for preservation of reconstitution property of the Davis-Hatefi enzyme) , then deactivation was achieved rapidly upon filtration of the enzyme through a Sephadex column. Such deactivated preparations could be reactivated again by the usual procedures. Most preparations of succinate dehydrogenase contain tightly bound oxaloacetate apparently in a 1:1 molar ratio with respect to the deactivated fraction (207,208). According to King and his colleagues (210, S l l ) , oxaloacetate binds to a sulfhydryl group on the larger subunit of the enzyme to abolish enzymic activity. Kearney and her colleagues have shown that the tightly bound oxaloacetate can be dissociated by various activators of the enzyme, such as succinate, malonate, IDP, ITP, and high concentrations of anions a t elevated temperatures but not in the 206. P. Cerletti and A. Manzocchi, Acta Vitaminol. Enzymol. 27, 5 (1973). 207. T. P. Singer, G. Oestereicher, and P. Hogue, Plant Physiol. 52, 616 (1973). 208. G. Oestereicher, P. Hogue, and T. P. Singer, Plant Physiol. 52, 622 (1973). 209. C. J. Coles, H. D. Tisdale, W. C. Kenney, and T. P. Singer, JBC 249, 381 (1974). 210. A. Vinogradov, D. Winter, and T. E. King, BBRC 49, 441 (1972). 211. D. B. Winter and T. E. King, BBRC 58,290 (1974).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

249

cold or by precipitation and gel exclusion of the enzyme (212, 213). The activation energies for activation and release of oxaloacetate were found to be the same. However, with some activators it was found that oxaloacetate release showed a considerable lag as compared t o recovery of enzymic activity. Moreover, oxaloacetate-free preparations could be reversibly deactivated a t alkaline pH, and activation by anions or FMNH, showed a lower activation energy than given above (199, 200). Thus, Ackrell et al. (213) suggested that oxaloacetate binding is not the cause of deactivation but a consequence of the deactivated conformation. That activation-deactivation of succinate dehydrogenase might involve conformational changes is suggested by ( a ) greater sensitivity of the activated enzyme to inhibition by N-ethylmaleimide and bromopyruvate, which has been interpreted as exposure of -SH groups ( S l d ) , and (b) the report that activation involves a free-energy change of only about 1.8 kcal/mole but an entropy change of 51 eu ( 2 5 , 2 6 ) .An important consideration with regard to the above studies is that activation-deactivation experiments involving incubations of 30 min or longer a t elevated temperatures, treatment a t p H 6, and filtration through Sephadex have been carried out using the dye reductase activity of the enzyme as the sole criterion. While the results are interesting and in some cases applicable to the membrane-bound enzyme, it is very doubtful that the reconstitution activity of soluble succinate dehydrogenase would survive the above treatments. Activation studies with ATP and reduced ubiquinone-10 are interesting because these experiments involve membrane-bound succinate dehydrogenase and the nature of the activators suggests physiological regulation. .4TP activation does not appear to involve membrane energization and reverse electron transfer, because it is insensitive to inhibition by oligomycin. The possibility that ATP induces the formation of the activator succinyl-CoA by reversal of the succinyl-CoA synthetase reaction is complicated by the finding that in metabolic states of the mitochondrion (e.g., the active state 3) where succinyl-CoA is known to accumulate, succinate dehydrogenase is in the deactivated state ( 2 3 ) .However, the deactivated state of succinate dehydrogenase under these conditions is believed to be governed mainly by the oxidized state of ubiquinone ( 1 9 7 ) . The possible role of ubiquinone in controlling succinate dehydrogenase was first suggested in 1970 by Rossi et al. (188), who made the following observations on succinate-PMS reductase activity of lyophilized, ubiquinone-depleted (by pentane extraction of the lyophilized particles),

<

212. E. B. Kearney, B. A. C. Ackrell, and M. Mayr, BBRC 49, 1115 (1972). 213. B. A. C. Ackrell, E. B. Kearney, and M. Mayr, JBC 249, 2021 (1974). 214. B. M. Sanborn, N. T. Feldberg, and T. C. Hollocher, BBA 227, 219 (1971).

250

YOUSSEF HATEFI AND DIANA L. STIGGALL

and ubiquinone-replenished submitochondrial particles. I n lyophilized and ubiquinone-replenished particles, TTFA and cyanide pretreatment diminished VmaXwith respect to PMS by about 50%. In ubiquinonedepleted particles, V,,, was already diminished by 50% and further treatment with TTFA or cyanide had no additional effect. These results were taken by the authors and by Singer and co-workers (23, 25) to indicate (a) an effect of ubiquinone on succinate dehydrogenase activity; (b) that as originally suggested by Singer’s laboratory there are two PMS reduction sites in the particles, one of which is abolished by TTFA or cyanide pretreatment of the particles; and (c) that ubiquinone is probably the second PMS reduction site in particles. Regarding point ( a ) , the experiments of Rossi et al. appear to involve an assay complication, however. The final electron acceptor in the succinate-PMS reductase assay was 2,6-dichloroindophenol, and both PMS and dichloroindophenol accept electrons in submitochondrial particles from components on the oxygen side of the TTFA block (214a). Thus, addition of TTFA, extraction of ubiquinone, or pretreatment of the particles with cyanide would abolish the latter route for electron transfer to PMS and dichloroindophenol, leaving only the pathway of dye reduction through succinate dehydrogenase itself. Therefore, the loss of 50% activity as a result of extraction of ubiquinone is probably referable to interruption of electron flow to the additional sites a t which PMS and dichloroindophenol are reduced, rather than to an activating effect of ubiquinone on succinate dehydrogenase. Gutman e t al. (196, 197) have shown, however, that reduced ubiquinone might, indeed, be a regulator of succinate dehydrogenase activity. Substrates (NADH and a-glycerophosphate) , which increased the level of reduced ubiquinone in particles, activated succinate dehydrogenase as did added reduced ubiquinone-10. Similarly, the resting state of mitochondria achieved by respiratory control showed high succinate dehydrogenase activity, which was rapidly diminished by the release of respiratory control with ADP or uncouplers (195). The greater activity of succinate dehydrogenase in the resting state has been interpreted as resulting from the accumulation of reduced ubiquinone under these conditions. The results of Gutman et al. are not subject to the complications of the work of Rossi et al., even though the succinate dehydrogenase assay was the same, i.e., with PMS and dichloroindophenol as electron acceptors. This 214a. Our preliminary results have shown that about 27% of the succinoxidase or succinate + P M S 2.6-dichloroindophenol activity of subrnitochondrial particles ran be measured in the absence of PMS with dichloroindophenol as electron acceptor. This activity is enchanced in the presence of added cytochrome c, and inhibited by antimycin A (80%) or TTFA.

+

4.

METAL-CONTAINING FLAVOPHOTEIN DEHYDROGENASES 05

Succtnate activation

5

I 10

1 I5

25 1

-

-

20

25

Time ( m i d

FIG.32. Activation of succinate dehydrogenase b y reduced ubiquinone-10. Phosphorylating submitochondrial particles ( ETPH) were resuspended in a sucroseTris-Mg buffer, pH 7.4, a t 4 mg protein/ml. Antimycin A ( I nmole/mg) and cyanide (1 m M ) were added and the sample placed under an atmosphere of NZ t o prevent autoxidation of reduced ubiquinone-10. Ubiquinone-10 was reduced with borohydride, neutralized with dilute acetic acid, and shaken till the first appearance of the yellow color of oxidized uhiquinone-10 t o ensure removal of unreacted borohydride, all a t 0". Activation of succinate dehydrogenase was started by adding 50 fi1 of either reduced ubiquinone-10 (curve A) or of ubiquinone-I0 (curve B) in absolute ethanol to 3 ml of enzymr giving 175 m M final concentration of the quinone. Curve C, no addition. Samples were withdrawn a t intervals and assayed immediately at 17". The horizontnl arrow indicates the maximal activation reached with succinate as activator. From Gutman et al. ( 1 9 7 ) .

is because ( a ) all the preparations used by Gutman et al. contained ubiquinone, and (b) in the experiments with NADH and added reduced ubiquinone-10 thc activation of succinate dehydrogenase was time-dependent (Figs. 32 and 33). 5. Mechaniswi Study of the reaction mechanism of succinate dehydrogenase has been complicated, partly because of the activation-deactivation properties Df the enzyme, and partly because most of the carly preparations studied had low iron content and low activities. Kinetic studies with activated, soluble preparations have led Zeylemaker et al. (215) to propose the fol215. W. P . Zeglrmaker, D. V. DerVartanian, C. Veeger, and E. C. Slater, BBA 178, 213 (1969).

252

YOUSSEF HATEFI AND DIANA L. STIGGALL

Time (min)

FIG.33. Activation of succinate dehydrogenase by NADH. A preparation of phosphorylating submitochondrial particles ( E T P d (succinoxidase activity = 1.18 pmoles succinate per min per mg at 30") was washed by centrifugation in a sucrose-Tris-Mg buffer (pH 7.4) and resuspended in the same buffer at 1 mg of protein/ml. Antimycin A (1 nmole/mg protein) was added to slow the rate of aerobic oxidation of NADH, followed by 0.25 mM NADH. Oxidation of the latter at 23" was monitored spectrophotometrically at 340 nm (dashed line). Samples were removed periodically and assayed immediately for succinate dehydrogenase activity in the presence of 0.33 mg of PMS/ml (solid line). A t 16 min a second aliquot of 0.25 mM NADH was added. From Gutman et al. (197).

lowing sequence for the partial reactions involving the interaction of substrate with the enzyme: E +S=ESr ESr 2 ESri ESrr A,, + E P f EP=E+P

+

Aped

where E, S, ESI, ESrr,A, and P are enzyme, substrate, ES complexes I and 11, electron acceptor, and product, respectively. Stereospecificity of the enzyme for dehydrogenation of succinate is trans irrespective of which hydrogen pair H,H, or H,H, is removed (216, 217). Monohalogen-substituted acids with the L-configuration have only one trans pair of hydro216. D. V. DerVartanian, W. P. Zeylemaker, and C. Veeger, in "Flavins and Flavoproteins," Symp. Adn Flavoproteins (E. C. Slater, ed.), p. 183. Elsevier, Amsterdam, 1966. 217. J. Rktey, J. Seibl, D. Arigoni, J. W. Cornforth,'G. Ryback, W. P. Zeylemaker, and C. Veeger, Eur. J . Biochem. 14,232 (1970).

4. METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

253

COOH

COOH

gens and serve as substrates. The D-stereoisomers do not serve as substrates but bind as competitive inhibitors. Electron paramagnetic resonance studies of DerVartanian et al. (218) with the Veeger et al. succinate dehydrogenase (149) indicated that maximal radical concentration occurred a t a level corresponding to only about 20-300/0 of the bound flavin, and that the g = 1.94 signal accounted for a t most 1 equivalent of unpaired electrons per mole of flavin. Recent work of Beinert et al. (184) with the activated soluble and particulate preparations indicates that succinate reduces not more than 60% of the ferredoxin-type iron-sulfur center of succinate dehydrogenase within the turnover time of the enzyme, and that flavin semiquinone formation neither precedes nor significantly lags behind the reduction of the ironsulfur center. The larger subunit by virtue of carrying the bound flavin is undoubtedly involved in substrate binding and electron transfer, and the semiquinone form of the enzyme is considered to be a catalytically significant intermediate. Iron-sulfur center S-1, which is substrate reducible and has an E m N 0 mV according to Ohnishi et al. (164), might be the species that interacts with flavin. The role of iron-sulfur center 5-2 with Em N -260 mV is not known. Preparations of succinate dehydrogenase with as little iron as 2 or 4 g-atoms/mole of flavin exhibit PMS reductase (albeit low) activity, g = 1.94 signal a t >20°K, and activation-deactivation effects. Therefore, it appears that these phenomena do not require the intactness of both iron-sulfur centers. The presence of PMS reductase activity in the 2-iron preparations further suggests the possible presence in these preparations of a stable 2 iron-2 sulfur center or of a fraction with higher iron-sulfur content which is responsible for the g = 1.94 signal and PMS reduction. The two subunits resolved by Davis and Hatefi (143, 166) by the use of chaotropes and freeze-thawing were inactive, separately and in combination, for succinate oxidation or fumarate reduction. However, the possibilities have not been fully explored. Nor has this sort of resolution been performed on the cyanide-treated enzyme to see whether one or the other subunit can be preferentially modified. Further work in these areas might 218. D. V. DerVartanian, C. Veeger, W. H. OrmeJohnson, and H. Beinert, BBA 191, 22 (1969).

254

YOUSSEF HATEFI AND DIANA L. STIGGALL

provide important clues to the reaction mechanism of succinate dehydrogenase and the role of the subunits.

B. SUCCINATE DEHYDROGENASE IN MICROORGANISMS All aerobic organisms, including yeast, appear to have a membranebound succinate dehydrogenase containing iron and covalently bound flavin (15, 16, 25). In contrast, the enzyme in anaerobic organisms is found in the cytoplasm and appears to be more effective as a fumarate reductase, a modification which is in accord with the physiological requirements of the organism. I n facultative anaerobes such as E. coli and S. cerevisiae, both the membrane-bound succinate dehydrogenase and the cytoplasmic fumarate reductase are found, their synthesis and concentration depending on the growth conditions. The succinate dehydrogenase of yeast mitochondria was isolated by Singer et al. (15, 219) in 1957, and stated to have a molecular weight of 200,000 and an iron:flavin ratio of 4:1, similar to the mammalian enzyme. These studies antedated, however, the purification of mammalian succinate dehydrogenase by Davis and Hatefi. Therefore, the exact molecular weight and composition of the yeast enzyme will have to be reexamined in light of present information. Hatefi et al. (220) have isolated the succinate dehydrogenase of Rhodospirillum rubrum by extraction of chromatophores with NaC10,. The enzyme has two subnits of molecular weights of approximately 60,000 and 25,000 (Fig. 34) (22Oa). Both contain iron-sulfur chromophores (221), and the larger subunit carries the covalently bound flavin. I n the intact enzyme, the ratio of flavin:iron:labile sulfide is approximately 1:8 :8. The enzymic properties of this prokaryotic enzyme are also very similar to the mammalian succinate dehydrogenase. Ferricyanide and PMS are reduced a t comparable V,,,,, rates, and the former inhibits above 1 mM. K , values with respect to succinate, PMS, and ferricyanide are 0.23, 0.11, and 0.3 mM, respectively (220). At 77OK, the succinate-reduced enzyme exhibits a free radical signal a t g = 2.00 and an iron-sulfur type of signal a t g = 1.93 (220).The absorption spectrum of the R . rubrum enzyme is very similar to that of mammalian mitochondria. A very interesting observation is that the R . rubrum succinate dehydrogenase can cross-interact with alkali-inactivated mammalian respiratory 219. T. P. Singer, V. Massey, and E. B. Kearney, ABB 69, 405 (1957). 220. Y . Hatefi, K. A. Davis, H. Baltscheffsky, M. Baltscheffsky, and B. C. Johansson, ABB 152, 613 (1972).

220a. K. A. Davis, I. P. Crawford, and Y . Hatefi, in preparation.

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

255

FIG.34. Electrophoresis of Rhodospirillum rubrum succinate dehydrogenase on SDS-polyacrylamide gel. The protein bands were visualized with Coomassie blue. From Davis et al. ( 2 2 0 ~ ) .

particles to reconstitute succinoxidase activity (Fig. 35). This activity is inhibited by TTFA. According to Hatefi et al. ( 2 2 0 ) , affinity of the R. rubrum enzyme for reconstitution with the mammalian respiratory chain is also similar to that of the mammalian succinate dehydrogenase. Rhodospirillurn rubrum succinate dehydrogenase can also reconstitute with alkali-inactivated R. rubrum chromatophores, but cross-interaction of the latter with the mammalian enzyme, though it occurs, is not equally efficient ( 2 2 1 ) . Tisdale et al. (222) have shown that several isoeymes of fumarate reductase occur in brewer’s yeast, ranging in molecular weight from 34,000 to 112,000. The predominant species had a molecular weight of 62,000221. Y. Hatefi, unpublished. 222. H. Tisdale, J. Hauber, G. Pragcr, P. Turini, and T. P. Singer, Eur. J. Biochem. 4, 472 (1968).

256

YOUSSEF HATEFI AND DIANA L. STIGGALL

42rnin

FIQ.35. Reconstitution of succinoxidase activity of alkali-inactivated bovine heart ETP with bovine and R . rubrum succinate dehydrogenases (SD). Left-hand trace : 114 pg alkali-treated ETP (alk-ETP) and 37 pg bovine SD per ml. Right-hand trace: 172 pg alk-ETP and 46 pg R. rubrum SD per ml. I n both cases alk-ETP and SD a t 10 times the concentrations indicated above were premixed with 10 m M succinate and preincubated for 3 min a t 30" before addition t o the assay mixture. Alk-ETP, bovine SD, or R . rubrum SD d o n e resulted in no oxygen uptake. Where indicated, 5.9 m M TTFA and 0.15 mM PMS were used. Assay temperature 30". S.A., specific activity. From Hatefi et nl. (220).

63,000. The enzyme contains noncovalently bound FAD, nonheme iron, possibly copper, but no labile sulfide. Electron paramagnetic resonance studies showed signals resulting from copper and high-spin ferric ions a t g = 4.3.

IV. ~-Glycerol-3-phosphateDehydrogenase (EC 1.1.99.5 1

The oxidation of L-glycerol 3-phosphatc to dihydroxyacetone phosphate is catalyzed by two different enzymes. One is the cytoplasmic NAD-linked a-glycerophosphate dehydrogenase, and the other is the niitochondrial enzyme, which appears to contain flavin and iron. The latter enzyme was first studied by Green in 1936 (223). It was shown to be associated with respiratory particles, and widely distributed in animal tissues. The highest concentration of the enzyme was found in the brain. Lardy and co-workers (264) studied the enzyme in deoxycholatesolubilized particles obtained from skeletal muscle, confirmed the finding 223. D.E. Green, BJ 30, 629 (1936). 224. T. Tung, I,. Anderson, and H. A. Lardy, ABB 40,191 (1952).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

I

400

500

257

600

Wavelength nm

FIG.36. Absorption spectrum of partially purified a-glycerophosphate dehydrogenase, protein concentration 13 mg/ml. (0) Spectrum of oxidized enzyme. The difference spectra (shown in the insert) were recorded after the addition of a few granules of sodium hydrosulfite ( 0 )or 5 pmoles substrate (0) in a total volume of 0.2 ml. I n the difference spectra, a decrease in optical density indicates bleaching. From Ringler (226).

of Green with regard to the specificity of the enzyme for L-a-glycerophosphate as substrate, and demonstrated that the reaction product was dihydroxyacetone phosphate. a-Glycerophosphate dehydrogenase has been solubilized by treatment of pig brain mitochondria with phospholipase A (225). Only partial purification of the enzyme has been achieved. The best preparations of Ringler and Singer (225-227) were shown to contain 1 mole of flavin per 2.1 X 10” g of protein and 1 g-atom of nonheme iron per 3.5 X lo5 g of protein. I t has been claimed that the flavin is FAD (227). The absorption spectrum of the above preparation is shown in Fig. 36. Phenazine methosulfate, ferricyanide, 2,6-dichloroindophenol, and methylene blue have been used as electron acceptors. With PMS a s electron acceptor, the best preparations of Ringler and Singer (226) from pig brain were shown to oxidize a-glycerophosphate a t 38O and p H 7.6 a t a rate of 3.4 pmoles/min x mg protein. Under these conditions K, for a-glycerophosphate was shown to be 9.5 mM, the same as the particle-bound enzyme (223). Dihydroxyacetone phosphate is a competitive inhibitor of the mammalian enzyme; K , = 0.18 m M a t 3 8 O and p H 7.6. Attempts a t reversing the action of a-glycerophosphate dehydrogenase in the presence of dihydroxyacetone phosphate plus reduced FMN, leucobenzylviologen, 225. R. L. Ringler, JBC 236, 1192 (1961). 226. R . L. Ringler and T. P. Singer, “Methods in Enzymology,” Vol. 5, p. 432, 1963. 227. T. P. Singer, “The Enzymes,” 2nd ed., Vol. 7, p. 345,1963.

258

YOUSSEF HATEFI AND DIANA L. STIGGALL

or leucomethylviologen as electron donor have not been successful (226). a-Glycerophosphate dehydrogenase is believed to be located in the outer phase of the mitochondria1 inner membrane (228). The enzyme appears to donate electrons to the respiratory chain beyond the level a t which Amytal inhibits NADH oxidation (229). It has been shown with the use of pentane-extracted mitochondria that electron transfer from the enzyme to the respiratory chain, but not to dyes, requires the presence of ubiquinone, and is inhibited by antimycin A (230). Therefore, i t appears that a-glycerophosphate dehydrogenase interacts with the mitochondrial electron transport system a t the level of ubiquinone, which is also the point of convergence of complexes I, 11, and I11 (29, 33, 231). I n agreement with the above findings, Ringler and Singer (177) have made the important observation that in antimycin-treated brain mitochondria the oxidation of a-glycerophosphate to dihydroxyacetone phosphate can be linked by way of succinate dehydrogenase to the reduction of fumarate. Further, Szarkowska and Drabikowska (232) have demonstrated the reduction of exogenous ubiquinone-6 by the particle-bound and the phospholipase-solubilized a-glycerophosphate dehydrogenase from pig brain. Both systems were inhibited by 3-phosphoglycerate. The best rate reported by these investigators for the soluble enzyme is 0.24 pmole QG reduced/min x mg protein at 37O and pH 7.2, which is only 7% of the PMS reductase activity of similar preparations. This low activity might in part have resulted from assay difficulties, which are usually encountered when water-insoluble homologs of ubiquinone are used as electron acceptor. It is also possible that (a) by analogy to the early preparations of succinate dehydrogenase (see Section 111), the preparation of a-glycerophosphate dehydrogenase used in these studies was damaged with respect to ubiquinone reduction, and (b) the reduction of ubiquinone by the dehydrogenase occurs by way of an unknown electron carrier in which the enzyme preparation used was deficient. Mitochondria from the flight muscle of house flies, Musca dornestica, have been shown to oxidize a-glycerophosphate a t exceptionally high rates (233, 234). This activity was shown to be inhibited by EDTA. It is believed that in these and other mitochondria the combined action of the soluble and the particle-bound a-glycerophosphate dehydrogenases 228. 229. 230. 231. 232. 233. 234.

M. Klingenberg, Eur. J. Biochem. 13, 247 (1970). B. Chance and B. Sacktor, ABB 76, 509 (1958). J. I. Salach and A. J. Bednarz, ABR 157, 133 (1973). Y. Hatefi, Clin. Chem. 11, 198 (1965). L. Szarkowska and A. K. Drabikowska, LifeSci. 7,519 (1963). R. W. Estabrook and B. Sacktor, JBC 233, 1014 (1958). B. Sacktor and D. G. Cochran, ABB 74,266 (1958).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES ouler

membranes

259

inner

Cytosol Glycerol- 1 -P

Dihydroxyacetone-P

It

FIG.37. The a-glycerophosphate cycle for the oxidation of extramitochondrial NADH by the mitochondrial respiratory chain. From Klingenberg (928).

provides the principal route for the transfer of reducing equivalents from cxtramitochondrial NADH to the mitochondrial electron transport SYStem (228, 229, 233-237). This process has been termed the “a-glycerophosphate cycle” (Fig. 37). Lee and Lardy (238) and others (239-242) have found that in the rat the a-glycerophosphate dehydrogenase activity of mitochondria from liver, kidney, adipose tissue, and heart were increased severalfold upon feeding of desiccated thyroid glands. The activity increase appeared to be organ-specific and particularly marked in liver, which showed a 20-fold increase after 10 days. The activity of the enzyme in brain, lung, spleen, stomach, small intestine, and testis was not appreciably increased. Thyroidectomy resulted in the decrease or disappearance of particle-bound a-glycerophosphate dehydrogenase activity in several organs, and a single injection of triiodothyronine restored the activity within 48 hr. The increased activity of a-glycerophosphate dehydrogenase induced by the thyroid hormone appears to result from synthesis of new enzyme (239).The possible role of this striking, organ-specific effect of the thyroid hormone has been discussed in relation to increased carbohydrate degradation in response to the increased oxidation of extramitochondrial NADH by the a-glycerophosphate cycle (238), as well as in relation to its effect on phospholipid synthesis ( 243) . 235. R. W. Estabrook and B. Sacktor, ABB 76, 532 (1958). 236. B. Sacktor, L. Packer, and R. W. Estabrook, .4BB 80, 68 (1959). 237. B. Sacktor and A. Dick, JBC 237,3259 (1962). 238. Y.-P. Lee and H. A. Lardy, JBC 240, 1427 (1965). 239. Y.-P. Lee, A. E. Takernori, and H. Lardy, JBC 234, 3051 (1959). 240. H. A. Lardy, Y.-P. Lee, and A. Takemori, Ann. N . Y. Acad. Sci. 86, 506 (1960). 241. 0. Z. Sellinger and K.-L. Lee, BBA 91, 183 (1964). 242. G. H. Isaacs, B. Sacktor, and T. A. Murphy, BBA 177, 196 (1969). 243. W. R. Frisell and J. R. Cronin, ir, “Electron and Coupled Energy Transfer in Biological Systems” (T. E. King and M. Klingenberg, eds.), Vol. 1, Part A, p. 177. Dekker, New York, 1971.

260

YOUSSEF HATEFI AND DIANA L. STIGGALL

Flavin-containing a-glycerophosphate dehydrogenases have been found also in Streptococcus faecalis (244) and Propionibacterium arabinosum (245). The enzyme from 8. faecalis is reported to contain FAD, have a K , for a-glycerophosphate of 4 mM, and a pH optimum of 5.8. I n addition t o dyes, this enzyme can interact directly with molecular oxygen to form H,O,. The preparation from P . arabinosum is particle-bound, has a K , for a-glycerophosphate of 26 p M , and is claimed to contain flavin and nonheme iron.

V. Choline Dehydrogenase (EC 1.1.99.1 1

The oxidation of choline to betaine is catalyzed by two enzymes. First, choline is oxidized to betaine aldehyde by a n enzyme which is found in mitochondria in membrane-bound form. This enzyme is believed to be a flavoprotein containing nonheme iron. Betaine aldehyde is then oxidized to betaine by a soluble enzyme, which is NAD-linked. Betaine aldehyde dehydrogenase appears to be present both in mitochondria and the soluble fraction of liver (243,246‘). The existence of choline dehydrogenase was first demonstrated by Mann and Quastel in 1937 (247, 248) in extracts of rat liver and kidney. These authors also obtained evidence that the first oxidation product of choline was betaine aldehyde. Others showed subsequently that choline oxidase activity resided in the mitochondria1 fraction of rat liver and is linked to the respiratory chain (249, 250). Detergents (251, 252), solvent treatment of fragmented mitochondria (253), and venom phospholipase (254-256‘) have been used for extraction and solubilization of choline dehydrogenase. Among these, the best method reported to date appears to be the digestion of acetone-powdered mitochondria with venom phospholipase. Choline dehydrogenase, partially purified from phospholipase extracts of rat liver mitochondria, contains 1 mole of flavin and 4 g-atoms of nonheme iron per 850,000 g protein. The flavin is claimed to be acid244. 245. 246. 247. 248. 249. 250. 251. 252. 253. 254. 255. 256.

N. J. Jacobs and P. J. VanDemark, ABB 88,250 (1960). N. Sone, J . Biochem. (Tokyo) 74,297 (1973). J. L. Glenn and M. Vanko, ABB 82, 145 (1959). P. J. G. Mann and J. H. Quastel, BJ 31, 869 (1937). P. J. G. Mgnn, H. E. Woodward, and J. H. Quastel, BJ 32, 1025 (1938). C. J. Kensler and H. Langemann, JBC 192, 551 (1951). J. N. Williams, Jr., JBC 194, 139 (1952). J. N. Williams, Jr. and A. Sreenivasan, JBC 203, 899 (1953). M. Korgenovsky and B. V. Auda, BBA 29,463 (1958). K. Ebisuzaki and J. N. Williams, Jr., BJ 60, 644 (1955). G . Rendina and T. P. Singer, BBA 30,441 (1958). G. Rendina and T. P. Singer, JBC 234, 1605 (1959). T. Kimura and T. P. Singer, “Methods in Enzymology,” Vol. 5, p. 562, 1962.

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extractable FAD, and the enzyme preparation is reported to contain trace amounts of a b-type cytochrome. For assay of the activity of membrane-bound enzyme, molecular oxygen, cytochrome c, ferricyanide, PMS, and dichloroindophenol can be used (254).The soluble enzyme reacts only with the latter two electron acceptors (227, 255, 256), thus indicating that other acceptors interact indirectly by way of the niitochondrial electron transport system. With PMS as electron acceptor, the K , for choline a t 38O and p H 7.6 is about 7 mM (227, 255). The best preparations of Kimura and Singer (256) oxidize choline a t a rate of 5.3 pmoles/min x mg protein a t 3 8 O . I n addition to choline, the particulate enzyme has been reported to oxidize arsenocholine (248) and other choline analogs (25‘7).Choline dehydrogenase is very sensitive to thiol inhibitors, and choline has been reported to protect against inhibition by p-mercuribenzoate (258).The oxidation of choline is competitively inhibited by betaine aldehyde ( K i = 2 mM) (256).Nitrogen mustard has also been reported to be a strong competitive inhibitor ( 2 5 9 ) , but others have reported no inhibition of choline dehydrogenase by nitrogen mustard ( 2 6 0 ) . Information regarding the involvement of flavin and iron in enzyme catalysis is not available. Rothschild et al. (258) have reported that dialysis of rat liver particles resulted in the loss of choline-cytochrome c reductase activity, which could be restored by addition of FAD but not FMN. However, these results have not been substantiated by others (255). Singer has stated that the difference spectrum of the enzyme “shows bleaching by substrate in both the flavin and the iron regions” (227).This spectrum has not been published. Relation to the Electron Transport System

It was shown by Strength et al. (261) that the oxidation of choline by a particulate preparation from rat liver was considerably enhanced upon addition of NAD. Others showed that choline oxidation by isolated rat liver mitochondria was completely inhibited by Amytal when oxygen, cytochrome c, ferricyanide, or methylene blue was the electron acceptor (262-664). Choline dehydrogenase activity of particles and soluble prepa257. I. C. Wells, JBC 207, 575 (1954). 258. H. A. Rothschild, 0. Cori, and E. S. G. Barrbn, JBC 208, 41 (1954). 259. E. S. G. Bnrron, G. R. Bartlett, and Z. B. Miller, J . E x p . M e d . 87, 489 (1948). 260. A . Sivak, A. J. Mahoney, Jr., and W. I. Rogers, Biochem. Pharmacol. 16, 1919 (1967). 261. D. R. Strength, J . R. Christensen, and L. J. Daniel, JBC 203, 63 (1953). 262. L. Packer, R. W. Estabrook, T. P. Singer, and T. Kimura, JBC 235,535 (1960). 263. L. Ernster, 0. Jalling, H. Low, and 0. Lindberg, Exp. Cell R e s . Suppl., 3, 124 (1955). 264. 0. Rendina and T. P. Singer, F e d . Proc., F e d . A m e r . SOC.Exp. Biol. 18, 308 (1959).

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YOUSSEF HATEFI AND DIANA L. STIGGALL

rations was reported to be insensitive to Amytal when assayed with P M S as electron acceptor (227, 262). Bianchi and Azzone (265) confirmed the findings of Strength et al. and showed that choline oxidation by intact rat liver mitochondria, but not by swollen mitochondria, was partially inhibited by rotenone. They further demonstrated that addition of choline to liver mitochondria in the presence of A D P resulted in the reduction of intramitochondrial nicotinamide nucleotides, and that under anaerobic conditions choline oxidation could be linked to the reduction of oxaloacetate to malate in the absence of an energy supply. These and other results led to the conclusion that reducing equivalents from choline dehydrogenase to the respiratory chain of intact mitochondria passed in part through the rotenone- and Amytal-sensitive site 1 of phosphorylation in the NADH oxidase pathway, and in part through another Amytal-sensitive, but rotenone-insensitive . point to ubiquinone and cytochrome b (243).

Kimura e t al. (266) showed that, unlike a-glycerophosphate, the oxidation of choline to betaine aldehyde in anaerobic mitochondria could not be linked to fumarate reduction. They also reported t h a t the choline oxidase activity of rat liver mitochondria was partially resistant to inhibition by antimycin A and quinoline oxide. They concluded, therefore, t h a t the mitochondria1 choline and succinate oxidase pathways were separate. The two chains were interlinked only between cytochrome cI and oxygen, and the choline chain involved an autoxidixable cytochrome b. These complications and the NAD stimulation of choline oxidation were resolved to a considerable extent by the work of Feinberg et al. (267) and Estabrook and his colleagues (268). The former group showed that the NAD stimulation was abolished when semicarbazide was present during choline oxidation. Under these conditions, semicarbazide interacted with, and prevented the NAD-linked oxidation of, betaine aldehyde which was formed as a result of choline oxidation. I n the absence of semicarbazide, choline oxidation was stimulated by NAD. These and other workers concluded that the rotenone inhibition of choline oxidation, which occurred in intact mitochondria, even in the presence of semicarbazide, involved an interference with the rate of entry of choline into intact mitochondria. Estabrook and co-workers (268) showed very clearly ( a ) that the antimycin-resistant oxygen uptake by mitochondria in the presence of choline resulted from inhibitor-resistant oxidation of endogenous substrates ; (b) that rotenone has little effect on choline oxidation by submitochondrial 265. 266. 267. 268.

G. Bianchi and G. F. Azzone, JBC 239, 3947 (1964). T. Kimura, T. P. Singer, and C. J. Lusty, BBA 44, 284 (1960). R. H. Feinberg, P. R. Turkki, and P. E. Witkowski, JBC 242, 4614 (1967). D. D. Tyler, J. Gonze, and R . W. Estabrook, ABB 115, 373 (1966).

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particles, whereas under the same conditions Amytal exerts a strong inhibitory effect; (c) that contrary to the previous report of others, the oxidation of choline by submitochondrial particles in the presence of PMS as electron acceptor was inhibited by Amytal; (d) that the concentration of Amytal needed for 50% inhibition of NADH (or 3-hydroxybutyrate) and choline oxidation by submitochondrial particles were 0.2 and 0.7 mM, respectively; and (e) that on the basis of spectra recorded a t 77OK, the nature and the degree of cytochromes reduced in r a t liver mitochondria were the same when the system was allowed to reach anaerobiosis as a result of succinate oxidation in the presence of rotenone or choline oxidation in the presence of rotenone and malonate. It has been shown that the oxidation of choline by isolated r a t liver mitochondria is biphasic (269). The initial phase of choline oxidation is slow and coupled to the uptake of inorganic phosphate. The ensuing phase is 3-5 times faster and not coupled t o phosphorylation. The slow phase can be extended in the presence of Mg2+and A D P or ATP. These compounds are considered t o control the permeability of mitochondria to choline ( 2 7 0 ) .Calcium ions and conditions which result in mitochondrial swelling and membrane disruption have been shown to increase choline oxidation (266, 271).

VI. lactate Dehydrogenases

Three types of lactate dehydrogenase are found in yeast, which may be considered as metal-containing flavoproteins. These are L-lactate :cytochrome c reductase or cytochrome b,, D-lactatc dehydrogenase, which is found in anaerobic yeast, and D-1actate:cytochrome c reductase, which is associated with the mitochondria of aerobic cells. A.

L (+)-LACTATE: CYTOCHROME c OXIDOREDUCTASE

(CYTOCHROME b,) ( E C 1.1.2.3) This enzyme [also known a t L (+)-lactate dehydrogenase] was first extracted from bakers’ yeast by Bernheim in 1928 (272). Bach et al. (273) showed in 1942 that lactate dehydrogenase copurified with a species of cytochronie b, which contained protoheme as prosthetic group. The 269. 270. 271. 272. 273.

T. Kagawa, D. R. Wilken, and H. A. Lardy, JBC 240, 1836 (1965). D. R. Wilken, T. Kagawa, and H. A. Lardy, JBC 240, 1843 (1965). G. R. Williams, JBC 235, 1192 (1960). F. Bernlieini, BJ 22, 1179 (1928). S. J. Bach, M. Dixon, and L. G. Zerfas, Nature (London) 149, 48 (1942).

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YOUSSEF HATEFI AND DIANA L. STIGGALL

cytochrome was designated cytochrome b, by Keilin and co-workers (274). The enzyme was crystallized by Appleby and Morton in 1954 (275, 276) and shown to contain FhlN in amounts equimolar to heme. This also marked the first crystallization of a cytochrome. These studies were confirmed by others (277) and extended to show that lactate dehydrogenase was specific for L (+) -lactate, and inhibitable by p-mercuribenzoate and Atebrin. Appleby and Morton (278) showed subsequently that the crystalline enzyme contained 5-6% DNA, but that the polynucleotide was not essential for activity. The crystalline preparation containing DNA is known as type I cytochrome b,, and the preparation from which DNA has been removed is known as type I1 cytochrome b,. These early studies have been reviewed (279-282). 1. Physical Properties

Cytochrome b, is found as a soluble protein in the autolysates of Saccharonayces cerevisiae. The crystalline preparations of Appleby and Morton (278) were shown to sediment as a single peak in the ultracentrifuge. Minimum molecular weight based on amino acid analysis and a heme extinction coefficient of 232 mM-l cm-’ was calculated to be 53,000 (283). The heme extinction coefficient was then corrected to 183 mM-l cm-’, and the minimum molecular weight per mole of heme recalculated to be 58,600 ( 2 8 4 ) . It was concluded that cytochrome b, is a tetrameric structure. This conclusion agreed with the results of X-ray diffraction studies on type I and type I1 crystals, which indicated molecular weights of 235,000 2 10,000 and 234,000 & 8,000, respectively, for these two preparations of cytochrome b, (285).The oxidized and reduced spectral bands of cytochrome b, are given in Table XIV. Subsequent studies showed that reduction and carboxymethylation of crystalline cytochrome b, yielded two unlike subunits of approximately 274. S. J. Bach, M. Dixon, and D. Keilin, Nature (London) 149, 21 (1942). 275. C. A. Appleby and R. K. Morton, Nature (London) 173, 749 (1954). 276. C. A. Appleby and R. K. Morton, BJ 71, 492 (1954). 277. E. Boeri, E. Cutolo, M. Luzzati, and L. Tosi, ABB 56, 487 (1955). 278. C. A. Appleby and R. K. Morton, BJ 75,258 (1960). 279. T. P. Singer, “The Enzymes,” 2nd ed., Vol. 7, p. 345, 1963. 280. A. P. Nygaard, “The Enzymes,” 2nd ed. Vol. 7, p. 557, 1963. 281. T. P. Singer, C. Gregolin, and T. Cremona, in “Control Mechanisms in Respiration and Fermentation” (B. Wright, ed.) , p. 47. Ronald Press, New York, 1963. 282. A. P. Nygaard, in “Control Mechanisms in Respiration and Fermentation” (B. Wright, ed.), p. 27. Ronald Press, New York, 1963. 283. C. Jacq and F. Lederer, E w . J . Biochem. 12, 154 (1970). 284. P. Pajot and 0. Groudinsky, Eur. J. Biochem. 12, 158 (1970). 285. C. Monteilhet and J. L. Risler, Eur. J. Biochem. 12, 165 (1970).

4.

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METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

TABLE XIV SPECTRAL PROPERTIES OF CYTOCHROME b t (TYPE II)O Oxidized form Band a

B r

s

uv

Reduced form

x

B

x

€

(nm)

(mM-1 cm-1)

(nm)

(mM-' cm-l)

560 530 413 360-365 275

9.2 11.3 129.5 34.4 83.5

557 528 424 328 269

30.9 15.6 183 39

88

From Pajot and Groudinsky (284).

21,000 and 36,000 daltons (286).These subunits had different amino acid compositions, and other results suggested that the heme binding site is on the heavy chain. At this time, Baudras (287) showed that L ( + ) -lactate: cytochrome c reductase isolated from the yeast Hansenula anomala was very similar to the Saccharamyces enzyme in molecular weight and heme and flavin content, but was considerably more stable and six times more active. Moreover, unlike the Saccharamyces enzyme, the activity of Hansenula cytochrome b, was inhibited in the presence of excess substrate. The Hansenula cytochrome b, appeared to be composed of four subunits of approximately 61,000 ? 5,000 daltons each (288). Baudras and Spyridakis (689) suggested, therefore, that the 21,000- and 36,000dalton subunits of the Saccharomyces enzyme were the result of artifactual splitting during isolation and crystallization of the type I cytochrome b,. The differences between the Hansenula and the Saccharomyces preparations of cytochrome b, were resolved by Jacq and Lederer (290) who showed that, when prepared in the presence of the protease inhibitor phenylmethylsulfonyl fluoride, the Saccharomyces enzyme does not crystallize as before, and shows a subunit size comparable to that of the Hansenula cytochrome b,. The enzyme so prepared had considerably improved stability and enzymic properties, and was inhibited at high lactate concentrations. It was concluded that the uncleaved, physiological form of Saccharomyces cytochrome b, has a molecular weight of 230,000, and is composed of four identical subunits, each associated with one FMN 286. 287. 288. 289. 290.

F. Lederer and A.-M. Simon, Eur. J . Biochem. 20, 469 (1971). A. Baudras, Biochimie 53, 929 (1971). F. Labeyrie and A. Baudras, Eur. J. Biochem. 25,33 (1972). A. Baudras and A. Spyridakis, Biochimie 53, 943 (1971). C. Jacq and F. Lederer, Eur. J. Biochem. 25, 41 (1972).

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YOUSSEF HATEFI AND DIANA L. STIGGALL

TABLE XV PARAMETERS OF INTACT A N D CLEAVED CYTOCHROME bp MOLECULAR Parameter N-Terminal residues

C-Terminal residues

Intact

Cleaved a-Subunit

iLys

a-Subunit

I

Glu

Ala

Val

LYE Ala ASP

/%Subunit ( Ala Minimum molecular weight per heme

58,100+7% (amino acids)

53,000 i~ 3% (amino acids) 58,600 f 2% (dry weight)

Molecular weight of peptide chains

57,500

=-Subunit 33,000-36,000 8-Subunit 21,000

Molecular weight

220,000 f 10% (gel filtration)

220,000 f 10% (gel filtration) 234,600 f 4% (crystallography) 240,000 f 4% (ultracentrifugation)

From Jacq and Lederer (291).

and one heme ($991).The amino acid composition of the enzyme prepared in the presence of phenylmethylsulfonyl fluoride has been determined, and it has been shown that alanine and glutamic acids are the C- and Nterminal residues, respectively (Table XV) (2991).These results indicated that, by comparison, the early crystalline preparations involved nearly 10% loss of peptide material, and circular dichroism spectra a t the Soret region of cytochrome b, showed a modification of the heme environment in the cleaved enzyme (2991). 2. Cytochrorne b, Core

Tryptic hydrolysis of cytochrome b, yields a polypeptide fragment which carries the heme and has a molecular weight of approximately 291. C. Jacq and F. Lederer, Eur. J. Biochem. 41,311 (1974).

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FLAVOPROTEIN DEHYDROGENASES

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11,000 (292). This material, designated cytochrome b, core, resembles the whole enzyme in its reduction potential, light absorption, and EPR spectra. Moreover, cytochrome b, core was shown to resemble soluble preparations of liver microsomal cytochrome b, in several respects, including cytochrome absorption spectrum, extinction coefficient, reduction potential, E P R signals at alkaline pH, and proton NMR spectra of the oxidized and reduced preparations (293, 294). Recently, Guiard et al. (295) have shown that the amino acid sequence of cytochrome b, core is very similar to that of microsomal cytochrome b,. They have also indicated that the amino acid sequence of cytochrome b, core is compatible with the peptide chain folding recently determined by others for cytochrome b,, and thus affords a similar heme environment as well. Cytochrome b, is reduced in microsomes by the enzyme cytochrome b, reductase, which is a flavoprotein. I n cytochrome b,, both the flavin and the heme are found in association with the same polypeptide chain. Thus, Guiard et al. (295) considered the possibility of a common ancestral origin for cytochromes b, and b,. They suggested that a pair of genes coding for cytochrome b, and cytochrome b, reductase might have fused in the course of evolution leading to cytochrome b,. 3. Enzymic Properties

Cytochrome b, is stereospecific for L ( + ) -lactate. It also oxidizes other a-hydroxymonocarboxylic acids a t slow rates (280, 298). As electron acceptors ferricyanide, methylene blue, 2,6-dichloroindophenol, 1,a-naphthoquinone 4-sulfonate, and cytochrome c have been used. This wide acceptor specificity is characteristic of a number of flavoproteins, which are generally capable of reducing quinoid structures and ferric compounds (297). However, as will be seen below, cytochrome c is considered to be the physiological electron acceptor for the yeast L-lactate dehydrogenase. Much of the available enzymic work on cytochrome b, has been performed on type I and type I1 enzymes which, as mentioned above, appear to have suffered limited proteolysis and peptide cleavage of the subunits 292. F. Labeyrie, 0. Groudinsky, Y. Jacquot-Armand, and L. Naslin, BBA 128, 492 (1966). 293. H. Watari, 0. Groudinsky, and F. Labeyrie, BBA 131, 592 (1967). 294. R. Keller, 0. Groudinsky, and K. Wiithrich, BBA 328, 233 (1973). 295. B. Guiard, 0. Groudinsky, and F. Lederer, Proc. Nat. Acad. Sci. U . S., 71, 2539 (1974). 296. R. H. Symons and L. A . Burgoyne, “Methods in Enzymology,” Vol. 9, p. 314, 1966. 297. M. Dixon, BBA 226,269 (1971).

268

YOUSSEF HATEFI AND DIANA L. STIGGALL

during purification. These preparations are very unstable and their enzymic properties as compared to crude yeast extracts reflect the structural damage they have sustained during purification (290, 291 ) . Comparative data regarding molar activities, K , values for substrate and cytochrome c, and inhibition by high levels of substrate have been published for the intact and cleaved Saccharomyces enzymes as well as for the intact cytochrome b, isolated from Hansenula anomalu (287, 289-291). It is generally agreed that the rate-limiting step is the transfer of reducing equivalents from substrate to the enzyme, that the initial reaction rate is first order with respect to substrate concentration, that flavin is the first electron acceptor (298-300), and that the transfer of electrons from flavin to the heme occurs intramolecularly (300). Anaerobic titration with L-lactate has indicated that the enzyme accepts three electrons (301). It has also been shown by EPR studies that upon reduction of the enzyme with L-lactate, a flavin semiquinone is formed to the extent of about 20% of the flavin content of the enzyme (301). However, it is not known whether the flavin semiquinone is a kinetic intermediate during enzyme catalysis. Ferricyanide appears to accept electrons from both the flavin and the heme (299-302), and it is believed that heme is required for cytochrome c reduction. Forestier and Baudras (30.2)have reported that, by treatment with guanidinium chloride, preparations of cytochrome b, could be rendered partially deficient in flavin and heme. Thus, enzyme preparations were obtained which contained 65-75% flavin and variable amounts of heme from about 12 to 100%. The low heme preparations showed considerably greater loss of cytochrome c reductase than ferricyanide reductase activity. When preparations with increasing content of heme relative to flavin were tested, both the ferricyanide and the cytochrome c reductase activities increased as a linear function of heme to flavin ratio (up to heme: flavin = 1) , but the increase in the heme content had a much greater effect on the cytochrome c reductase activity of the enzyme. The apoenzyme of cytochrome b , has been prepared. However, reconstitution with FMN, heme, and F M N plus heme in all cases resulted in extremely 298. M. Iwatsubo, A. Baudras, A. di Franco, C. Capeillere, and F. Labeyrie, in “Flavins and Flavoproteins,” 2nd Int. Symp. (K. Yagi, ed.), p. 41. Univ. Park Press, Baltimore, Maryland, 1968. 299. A. Baudras, C. Capeillere-Blandin, M. Iwatsubo, and F. Labeyrie, in “Strbcture and Function of Oxidation Reduction Enzymes” (A. Akeson and A. Ehrenberg, eds.), p. 273. Pergamon, Oxford, 1972. 300. R. K. Morton and J. M. Sturtevant, JBC 239, 1614 (1964). 301. K. Hiromi and J. M. Sturtevant, JBC 240, 4662 (1965). 302. J.-P. Forestier and A . Baudras, in “Flavins and Flavoproteins,” 3rd Int. Symp. (H. Kamin, ed.), p. 599. Univ. Park Press, Baltimore, Maryland, 1971.

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low activities (SOZ-SO4). Now that intact preparations of cytochrome b, are available, further efforts on these lines might yield clearer results regarding the roles of flavin and heme in the reduction of cytochrome c and artificial acceptors. The kinetics of cytochrome c reduction by L-lactate dehydrogenase are somewhat complicated because the enzyme binds cytochrome c strongly M ) (299, 300, 305). The cleaved enzyme binds one mole of (K, = cytochrome c per mole, but the intact preparations of Haiisenula bind 4 moles of cytochrome c per mole of enzyme, i.e., one mole of cytochrome c per subunit (287). The cytochrome b,-cytochrome c adduct of Saccharoinyces can be crystallized. Examination of the crystals have suggested that the crystal lattice of cytochrome b, can accommodate cytochrome c without an apparent change in the crystal structure (299, 305). The association of cytochrome c with L-lactate dehydrogenase does not depend on the oxidation-reduction state of either cytochrome, and similar to L-lactate protects the enzyme against denaturation by 3 M urea (299, 305). The interaction of cytochrome c with L-lactate dehydrogenase is considered to be specific. I n addition to the above results, i t has been shown that lysozyme, which is similar to cytochrome c in size and charge, does not compete for the binding of cytochrome c to the enzyme (299, 305). The role of L-lactate dehydrogenase in the physiology of aerobic yeast is not clear. It has been shown that its presence in yeast depends on the availability of oxygen (306), and that in the presence of antimycin A, which inhibits electron transfer to cytochrome c from NADH-linked substrates, L-or D-lactate can partially support the growth of Saccharomyces cerevisiae (307). Under these conditions, cyanide inhibited the growth. Therefore, i t has been concluded that L- and D-lactate-cytochrome c reductases can feed electrons to the respiratory chain at the level of cytochrome c and provide energy through the third site of oxidative phosphorylation (30’7).

B. D(-)-LACTATE:CYTOCHROME c OXIDOREDUCTASE (EC 1.1.2.4) This enzyme is tightly associated with the mitochondria of aerobic yeast. Similar to L-lactate: cytochrome c reductase, it is produced in yeast R. K. Morton and K. Sheplcy, Biochem. Z . 338, 122 (1963). M. Mevel-Ninio, P. Pajot, and F. Labeyrie, Biochimie 53, 35 (1971). A. Baudras, M. Krupas, and F. Labeyrie, Eur. J. Biochem. 20, 58 (1971). F. Labeyrie and M. Somlo, “Homologous Enzymes and Biochemical Evolution Colloquium” (Nguyen van Thoai*and J. Roche, eds.), p. 93. Gordon & Breach, New York, 1968. 307. P. Pajot and M. Claisse, Proc. Znt. Congr. Biochem., 9,1973 p . 239 (1973). 303. 304. 305. 306.

270

YOUSSEF HATEFI AND DIANA

L.

STIGGALL

during oxygen adaptation. It was suggested that both the D- .and the L-lactate cytochrome c reductases arise during oxygen adaptation from the D-2-hydroxyacid dehydrogenase of anaerobic yeast. However, this hypothesis has not found experimental support (308-310). 1. Physical Properties D-Lactate: cytochrome c reductasc has been extensively purified from the respiratory particles of bakers’ yeast by two different methods (308, 311-313). One method involves the treatment of particles with acetone and n-butanol, and the other involves treatment with Triton X-100, phospholipase A, and bacterial protcase. The latter method appears to result in greater purification, and higher yield, activity, and stability of the enzyme (312, 313). According to Gregolin and Singer (312), purified preparations of D-lactate: cytochrome c reductase contain 1 mole of FAD per 50,000 5,000 g protein, and 1 g-atom of Zn2+ per 22,000-27,000 g protein. They have concluded that the flavin content and the sedimentation constant of S = 6.8 suggest that the enzyme has a molecular weight of about 100,000 and contains 2 moles of FAD and 6 6 g-atoms of Zn2+ per mole. These conclusions are subject to change, however, because the diffusion constant and the partial specific volume of the enzyme are not known, and partial loss of flavin during purification of the enzyme cannot be ruled out.

*

2. Enzymic Properties D-Lactate :cytochrome c reductase can oxidize D-2-hydroxymonocarboxylic acids, but only D-lactate and D-2-hydroxybutyrate are oxidized at appreciable rates. The enzyme exhibits a similar high specificity for electron acceptors. It reacts with cytochrome c and phenazine methosulfate as electron acceptors, but not with ferricyanide, methylene blue, 2,6dichloroindophenol, and menadione (308, 312, 313). With Dlactate as substrate and a t V,,, with respect to acceptor, phenazine methosulfate is reduced a t 30° eight times as fast as cytochrome c (308). The K , values a t 30° and pH 7.5 are D-lactate, 0.29 mM; ~-2-hydroxybutyrate, 1.4 mM; phenazine methosulfate, 4.5 mM; and cytochrome c, 5.4 p M . The turnover number of the enzyme, isolated with the use of Triton 308. C. Gregolin and T. P. Singer, BBRC 4, 189 (1961). 309. A. P.Nygaard, JBC 236, 1585 (1961). 310. T. P. Singer, E. B. Kearney, C. Gregolin, E. Boeri, and M. Rippa, BBA 54, 52 (1961). 311. A. P.Nygaard, JBC 236, 920 (1961). 312. C.Gregolin and T. P. Singer, BBA 67, 201 (1963). 313. T. P. Singer and T. Cremona, ‘‘Methods in Enzymology,” Vol. 9, p. 302, 1966.

4.

27 1

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

TABLE XVI I N H I B I T O R S O F D ( -)-LACTATk::

CYTOCHROME

C REDUCTASEa

Concentration (M)

Inhibitor p-Mercuriphenyl sulfonate p-Mercuriphenyl sulfonate H202

Oxalate Oxalate Oxalate EDTA EDTA o-Phenanthroline* o-Phenan throline"

x x I x 6 x 1x 5 x 4 x 1x 3.5 x 3.5 x 5 5

10-7

10-3 10-3 10-8 10-6 10-3 10-3 10-2 10-3 10-3

Inhibition (%) 60 60 0 22 50 92 25 51 95 90

~

From Gregolin and Singer (319). * Overnight dialysis a t p H 6.5 against the indicated concentration of inhibitor. Incubated for 15 min a t 30'. a

X-100 and phospholipase A, is reported to be 90,000 moles lactate/ min x mole flavin at 30° and pH 7.5. The reaction of the enzyme does not appear to be reversible (312). D-Lactate :cytochrome c reductase is inhibited by p-mercuriphenyl sulfonate salts, metal chelators, and dicarboxylic acids such as oxalate and oxaloacetate (Table XVI) (312, 314, 315). According to Nygaard (314), salts (cations) inhibit a t the acceptor site, and dicarboxylic acids a t the substrate site. Cremona and Singer (315) have studied the inhibitions by metal chelators and by oxalate. They recognized two types of inhibition. One type of inhibition is that which is caused by E D T A or oxalate. This kind of inhibition is reversed immediately upon dilution of the enzyme-inhibitor mixture. The second is that which results from addition of o-phenanthrolinc. Enzyme preparations treated with o-phenanthroline bind 2 moles of the chelator per mole of Zn2+.This complex is stable and inactive, and does not result in the release of Zn2+.The inactive o-phenanthroline-enzyme complex can be reactivated by dialysis, addition of divalent metal ions such as Zn2+, Co2+,Mn2+, and Fez+, or by incubation at elevated temperaturcs ( 5 4 5 O ) (312, 3 1 5 5 1 7 ) . It has been shown that heat treatment does not involve the release of o-phenanthroline. The authors suggested that thermal reactivation of the o-phenan314. A. P. Nygaard, JBC 236, 2128 (1961). 315. T. Cremona and T. P. Singer, JBC 239, 1466 (1964). 316. A. Ghiretti-Magaldi, T. Cremona, T. P. Singer, and P. Bernath, BBRC 5, 334 (1961). 317. T. Crernona and T. P. Singer, BBA 57, 412 (1962).

272

POUSSEF HATEFI AND DIANA L. STIGGALL

throline-enzyme complex is the result of a change in the conformation of the enzyme molccule. Other studies have suggested to these authors that Zn?+is involved in the binding of substrate to the enzyme (312). It has been shown that by treatment with ammonium sulfate a t acid pH, flavin can be partially removed from the enzyme (318). Addition of FAD, but not FMN, reactivated the enzyme. Zinc is not removed under these conditions, and its addition is not required for reactivation. The metal appears to be very tightly bound to the enzyme (312) ; its removal without protein denaturation has not been achieved.

c. D-2-HYDROXYACID DEHYDROGENASE (EC 1.1.99.6) It was discovered in 1958 that anaerohically grown yeast contains a form of lactate dehydrogenase which is different from the D- and L-lactate:cytochrome c reductases of aerobic yeast (306, 319). The enzyme has been partially purified (320, 321), and shown to contain flavin (320-322). Gel filtration studies have suggested a molecular weight of about 100,000 (320, 321). Preparations of the enzyme oxidize several D-2hydroxyacids to the respective keto acids in a reversible manner (320). For the forward reaction, ferricyanide, 2,6-dichloroindophenol, menadione, and methylene blue have been used as electron acceptors, and for the reverse reaction leucomethyl viologen and FMNH, are effective electron donors (320).A number of L-2-hydroxyacids and 2-keto acids have been shown to be competitive inhibitors. Oxalate, cyanide, o-phenanthroline, and EDTA are also potent inhibitors (320, 321, 323, 324). The inhibition by metal chelators develops slowly and is reversed by addition of Zn2+,Co2+,Mn2+,or Fez+ (320, 323-326). Substrates prevent the inhibition by chelators a t concentrations considerably lower than their respective K, values (327). It has been suggested that EDTA inactivation involves the removal of a metal, most probably Zn2+,from the substrate binding site of the enzyme (325, 326, 328, 329). However, others have 318. C. Gregolin and T. P. Singer, BBA 57,410 (1962). 319. P. P. Slonimski and W. Tysarowski, C . R. Acad. Sci. 246, 1111 (1958). 320. T. Cremona, JBC 239, 1457 (1964). 321. J. Rytka and W. Tysarowski, Acta Biochim. Pol. 12,229 (1965). 322. M. Iwatsubo, BBA 77, 568 (1963). 323. E. Boeri, T. Cremona, and T. P. Singer, BBRC 2,298 (1960). 324. A. Curdel, L. Naslin, and F. Labeyrie, C. R. Acad. Sci. 249, 1959 (1959). 325. A Curdel and F. Labeyrie, BBRC 4,175 (1961). 326. A. Curdel, C. R . Acad. Sci. 254, 4092 (1962). 327. F. Labeyrie and E. Stachiewicz, BBA 52, 136 (1961). 328. E. Stachiewicz, F. Labeyrie, A. Curdel, and P. P. Slonimski, BBA 50, 45 (1961). 329. M. Iwatsubo and A. Curdel, BBRC 6, 385 (1961).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

273

shown that, similar to D-lactate :cytochrome c reductase, EDTA-treated D-2-hydroxyacid dehydrogenase can be reactivated by dialysis or by incubation a t elevated temperatures in the absence of added metals (320, 330, 331). The latter authors believe that chelator treatment results in the formation of an inactive enzyme-chelator complex without the removal of metal. This complex can be reactivated by addition of metal ions or dialysis, which presumably will result in the removal of the chelator, or by heat treatment, which converts the inactive complex to an active form. A similar mechanism has been invoked for the inhibition of D-lactate :cytochrome c reductase by o-phenanthroline. Thus, the presence and possible role of Zn2+and the nature of the flavin prosthetic group of D-Zhydroxyacid dehydrogenase have yet to be unambiguously demonstrated. Howeger, it might be added that the enzyme can be inactivated by treatment with ammonium sulfate a t acid pH, and reactivated by FAD, but not by F M N (329). Further, the flavin in EDTA-inactivated preparations is not reduced by D-lactate, but addition of Zn? results in rapid bleaching at 450 nm (330). These results have been considered as evidence that the flavin prosthetic group is FAD, and that the metal is necessary for the reduction of flavin by substrate. Soluble D-lactate dehydrogenases with enzymic properties similar to those of the D-2-hydroxyacid dehydrogenase of anaerobic yeast have been isolated from rabbit kidney mitochondria (322-334) and from a species of Mycobacterium ( 3 3 5 ) .It is not clear whether these enzymes are metalcontaining flavoproteins.

VII. Nitrite Reductases (EC 1.6.6.4)

Nitrate reduction and assimilation is a fundamental biological process in plants and various microorganisms. In this process nitrate is reduced ultimately to ammonia. Thus, as shown in Eq. ( 5 ) , the reduction of nitrate to ammonia requires eight electron or hydrogen equivalents. HN03

+8H+

NH.7

+ 3H20

(5)

The first reduction product of nitrate is nitrite. This reaction is catalyzed 330. A. Ghiretti-Magaldi, T. Cremona, T. P. Singer, and P. Bernath, BBRC 5, 334 (1961). 331. T. Cremona and T. P. Singer, Nature (London) 194,836 (1962). 332. P. K. Tubbs, BBRC 3, 513 (1960). 333. P. K. Tubbs and G. D. Greville, BJ 81, 104 (1961). 334. P. K. Tubbs, BJ 82,36 (1962). 335. T. Szumilo and M. Szymona, Physiol. Chem. Phys. 4, 407 (1972).

274

YOUSSEF HATEFI AND DIANA L. STIGGALL

by the molybdenum- and FAD-containing enzyme, nitrate reductase, which is discussed in Volume XII, Chapter 6, p. 402. Enzyme systems which catalyze nitrite reduction have been observed in bacteria (336-343), fungi (344-351), green algae (348, 352-355) , and higher plants (344, 356-362). While the assimilatory nitrite reductases convert nitrite to NH,, the denitrifying organisms reduce it to nitric oxide (338, 340, 363) or nitrogen gas (336). Examples of denitrifying nitrite reductases are the enzymes of Pseudomoms denitnficans ( S d O ) , and P . aeruginosa (338, 364-366), which convert nitrite to nitric oxide, and of P. stutzeri (336), which reduces nitrite to NO and N,. The nitrite reductase of P. denitrificuns has been partially purified. The enzyme reduces 336. C. W. Chung and V. A. Najjar, JBC 218,617 (1956). 337. D. Spencer, H. Takahashi, and A. Nason, J. Bacterial. 73, 553 (1957). 338. G. C. Walker and D. J. D. Nicholas, BBA 49, 350 (1961). 339. R. A. Lazzarini and D. E. Atkinson, JBC 236, 3330 (1961). 340. B. C. Radcliffe and D. J. D. Nicholas, BBA 153,545 (1968). 341. 0. Prakash and J. C. Sadana, ABB 148,614 (1972). 342. J. M. Vega, M. G. Guerrero, E. Leadbetter, and M. Losada, BJ 133, 701 (1973). 343. C. D. Cox, Jr. and W. J. Payne, Can. J. Microbial. 19, 861 (1973). 344. A. Nason, R. G. Abraham, and B. C. Averback, BBA 15, 159 (1954). 345. J. Rivas, M. G. Guerrero, A. Paneque, and M. Losada, Plant Sci. Lett. 1, 105 (1973). 346. D. J. D. Nicholas, A. Medina, and 0. T. G. Jones, BBA 37,468 (1968). 347. K. Yamafuji, Y. Osajima, H. Omura, and 8. Hatano, Enzymologia 21, 245 (1960). 348. E. Kessler, Annu. Rev. Plant. Physiol. 15, 57 (1964). 349. K. A. Cook and G. J. Sorger, BBA 177,412 (1969). 350. R. H. Garrett, BBA 264, 481 (1972). 351. M. A. Lafferty and R. H. Garrett, Abstr. 7Jrd. Annu. Meet Amer. Sac. Microbiol. p. 194 (1973). 352. E. Kessler and F. C. Czygan, Experientia 19, 89 (1963). 353. M. G. Guerrero, J. Rivas, A. Paneque, and M. Losada, BBRC 45, 82 (1971). 354. W. G. Zumft, BBA 276, 363 (1972). 355. A. Hattori and I. Uesugi, Plant Cell Physiol. 9, 689 (1968). 356. G. G. Roussos and A. Nason, JBC 235,2997 (1960). 357. R. H. Hageman, C. F. Cresswell, and E. J. Hewitt, Nature (London) 193, 247 (1962). 358. K. W. Joy and R. H. Hageman, BJ 100, 263 (1966). 359. K. Asada, G . Tamura, and R. S. Bandurski, JBC 244, 4904 (1969). 360. J. Cardenas, J. L. Barea, J. Rivas, and C. G. Moreno, FEBS (Fed. Eur. Biochem. Sac.) Lett. 23, 131 (1972). 361. M. J . Dalling, N. E. Tolbert, and R. H. Hageman, BBA 283, 505 (1972). 362. M. J. Dalling, N. E. Tolbert, and R. H. Hageman, BBA 283, 513 (1972). 363. A. Nason, Bacterial. Rev. 26, 16 (1962). 364. T. Yamanaka, A. Ota, and K. Okunuki, BBA 44,397 (1960). 365. T. Yamanaka and K. Okunuki, BBA 67,379 (1963). 366. T. Yamanaka, Nature (London) 204, 253 (1964).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

275

nitrite to nitric oxide in the presence of NADH or NADPH and FMN, FAD, or riboflavin. It can also use artificial electron donors, such as reduced benzyl viologen or leucomethylene blue, in the absence of flavins. Inhibitor studies have suggested the involvement of metals and active thiol. Walker and Nicholas (338)have reported the isolation and 600-fold purification of an enzyme from P. aeruginosa, which reduces nitrite to nitric oxide. The preparation contained 1.5 nmoles of FAD per mg protein, a c-type cytochrome and an absorption band a t 630-635 nm, suggestive of copper. As electron donors, reduced FMN, FAD, riboflavin, pyocyanine, and methylene blue were effective, but not NADH, NADPH, or reduced cytochrome c. The preparation required phosphate or sulfate for maximal activity. The cytochrome and the 630-635-nm band were reduced under anaerobic conditions with a suitable electron donor and readily oxidized by nitrite. The K , for NaNO, is reported to be 3.1 X lo-” M . The presence of an active thiol in the enzyme is indicated by p-mercuribenzoate inhibition and glutathione reactivation. Yamanaka and co-workers (364-366) have crystallized a cytochrome oxidase from P . aeruginosa which oxidizes Pseudomonus ferrocytochrome 0551. It is also capable of nitrite reduction with a turnover number of 4000 moles nitrite reduced under anaerobic conditions to nitric oxide per minute a t 37O. It is an adaptive enzyme, nitrate being essential for its biosynthesis. The enzyme has a molecular weight of 120,000, with two subunits of equivalent molecular weight, 2 heme c and 2 heme d groups per mole (Fig. 38) (36%). Nitrite reductase activity is 94% inhibited M KCN, but only by CO. The lack of CO inhibition appears by 8 X to be related to the fact that the enzyme has a greater affinity for nitrite than for carbon monoxide. Nitrite reduction in assimilatory nitrate-reducing Neurospora crassa, Tomlopsis nitratophila, Azotobacter vinelandii, and Azotobacter chroococcum appears to be catalyzed by enzyme systems which require flavin and metals. The enzyme from N . crassa has been partially purified, and its molecular weight has been estimated to be 300,000 (344, 346, 351, 367). The enzyme reduces both nitrite and hydroxylamine to ammonia and utilizes NADH or NADPH as electron donor. It is reported to be a FAD-dependent enzyme and to contain iron, copper, and active thiol (346, 367). Three moles of NADH are oxidized per mole of nitrite reduced to ammonia. It has been suggested that the reduction of nitrite occurs in three steps, each involving two electrons. Thus, hyponitrite and hydroxylamine have been proposed as successive intermediates in the re366a. J. C. Gudat, J. Singh, and D. C. Wharton, BBA 292, 376 (1973); D. C. Wharton, private communication. 367. A. Medina and D. J. D. Nicholas, BBA 25, 138 (1957).

276

YOUSSEF HATEFI AND DIANA L. STIGGALL

Wavelength (nm)

FIG.38. The absorption spectra of crystalline Pseudomonas cytochrome oxidase. The crystals were dissolved in 0,2 M phosphate buffer (pH 7.0). (---) Oxidized, (-) reduced with sodium dithionite. From Yamanaka and Okunuki (366).

duction of nitrite to NH, (36'7). The nitrite reductase of N . crassa is inducible by nitrate or nitrite and repressed by ammonia (350). The nitrite reductase of Torulopsis nitratophila is specific for NADPH and FAD, and can utilize reduced benzyl or methyl viologen as electron donor, but not reduced flavins (345). With NADPH as electron donor, nitrite reduction is inhibited by cyanide and mercurials. Michaelis constants for FAD and nitrite have been reported to be 45 n M and 19 p M , respectively. Unlike the Neurospora enzyme, the nitrite reductase of T. nitratophila could not reduce hydroxylamine in the presence of NADPH and FAD. That hydroxylamine might not be an obligatory intermediate, or occur as a free intermediate, in the reduction of nitrite to ammonia is suggested by the properties of nitrite reductases of Azotobacter chroococcum and Escherichia coli. The former is an adaptive enzyme, the formation of which requires nitrate or nitrite in the culture (342).It is FAD-dependent and presumably contains metals and p-mercuribenaoate inhibitable

4.

METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

277

thiols. It reduces nitrite to ammonia in the presence of NADH as electron donor and does not appear to produce hydroxylamine as an intermediate. Cyanide competitively inhibits the reduction of nitrite with a K i= 32 nM. The K , values for nitrite and NADH are 5.5 and 15 p M , respectively. The enzyme is inhibited upon preincubation with NADH. Nitrite protects against NADH inhibition and reverses it. Sucrose density gradient centrifugation has suggested a molecular weight of 67,000 for the A . chroococcum enzyme. The E. coli enzyme can reduce nitrite and hydroxylamine to ammonia a t the expense of NADPH (339). However, with the use of 15N-nitrite it was shown that hydroxylamine was not an intermediate in the reduction of nitrite. No cofactor requirements were shown for the E . coli enzyme, but similar to other flavin and metal requiring nitrite reductases it was inhibited by cyanide and mercurials. The nitrite reductase of Azotobacter vinelandii ( A . agile) was extracted in soluble form by Nason and his colleagues (337). The preparation reduced nitrite and hydroxylamine in the presence of reduced nicotinamide-adenine dinucleotides and required flavin for maximal activity. FAD was shown to be specific for nitrite reduction, whereas both FAD and FMN were active for hydroxylamine reduction. The hydroxylamine reductase activity of the preparation was enhanced in the presence of Mn2+.Ammonia was shown to be the product of nitrite reduction, but the product of hydroxylamine reduction was not identified. Another nitrite and hydroxylamine reductase, which had a MnZ+requirement, was also isolated and partially purified in Nason’s laboratory from soybean TABLE XVII THEPHYSICAL PROPERTIES OF PURIFIED NITRITEREDUCTASE FROM Achromobacter fscherio Sedimentation constant,, slo,w Diffusion coefficient, D ~ o . ~ Molecular weight (Archibald procedure)b Molecular weight (calculated from diffusion and sedimentation constants)b Heme content (nmoles/mg protein) Minimum molecular weight (from heme content) Iron content Minimum molecular weight (from iron content) Isoelectric point

5.2 s 5.56 X lo-’ om2 sec-l 95,000 +_ 4,000 84,000 19 52,500 0.102-0.105% 54,000 Around p H 4,. 5

From Prakash and Sadana (341). In the calculation, a value for the partial specific volume of 0.73 ml/g for nitrite reductase is assumed.

278

YOUSSEF HATEFI AND DIANA L. STIGGALL

I

t

0.9

4 20

I!

i!

Wavelength (nm)

FIQ.39. The absorption spectra of Achromobacter fischeri nitrite reductase. Spectra were recorded in 0.05 1cI phosphate buffer, p H 7.5, at 0.41 mg enzyme protcin/ml. (-) Oxidized, (- * -) reduced with dithionite, (---I NO,- (or hydroxylamine) added to the dithionite reduced enzyme. From Prakash and Sadana (341).

leaves (356).However, the enzyme preparation did not require flavin, but had an absolute requirement for an unidentified, heat-stable factor, which had an absorption peak a t 312 and 315 nm, respectively, in 0.1 N HC1 and 0.1 M pyrophosphate, pH 7.0. The peak shifted to 358 nm in 0.1 N NaOH. The nitrite reductase system of Achromobacter fischeri appears to be composed of two separable enzymes (341). The first enzyme is a flavin reductase and utilizes NADH or NADPH to reduce FMN or FAD. The second interacts with the flavin reductase and converts nitrite and hydroxylamine to ammonia. The nitrite reductase enzyme has a molecular weight of 95,000 f 4,000 (Table XVII), contains two heme c per mole, and is inhibited by p-mercuribenzoate, cyanide, and carbon monoxide.

4. METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

279

The latter inhibition is reversed by light. Urea inactivation-reactivation studies showed parallel loss and recovery of nitrite and hyroxylamine reductase activities, and nitrite was shown to inhibit hydroxylamine reduction. These results have suggested that the enzyme has a common binding site for nitrite and hydroxylamine. The absorption spectra of the A . fischeri enzyme (oxidized, reduced, and reduced plus nitrite or hydroxylamine) are shown in Fig. 39.

VIII. Adenylyl Sulfate Reductases (EC 1.8.99.2)

Two major pathways are known for the reduction of sulfate. One is the assimilatory pathway, which reduces sulfate to the extent necessary for satisfying the nutritional requirements of the organism. I n this pathway, which has been extensively studied in yeast by Robbins and Lipmann (368) and Bandurski and his colleagues (369, 370), sulfate is first activated in the presence of ATP by the enzyme ATP-sulfurylase t o form adenosine 5'-phosphosulfate (APS). Then in a second reaction, APS is phosphorylated in the 3' position by ATP to form 3'-phosphoadenosine 5'-phosphosulfate (PAPS) ATP APS

+ Solz+ ATP

+ +

APS PP PAPS ADP

(6) (7) In the presence of appropriate enzymes, the sulfate group of PAPS can be donated to various acceptors, such as carbohydrates, steroids and phenols, or become reduced to sulfite for assimilatory purposes. Figure 40 shows a unified scheme for sulfate and sulfite assimilation by algae as proposed by Abrams and Schiff (371). The second pathway by which sulfate is reduced is the dissimilatory pathway in which sulfate is the terminal electron acceptor and leads to the formation of large quantities of H,S. During the dissimilatory reduction of sulfate, APS is formed as in Eq. ( 6 ) . Then APS is reduced directly to sulfite and AMP by the enzyme APS-reductase. Table XVIII shows the data of Peck (372) on the pathway of sulfate reduction in various microorganisms. Adenylyl sulfate (APS) reductase is a flavoprotein, which contains iron and possibly acid-labile sulfide. It catalyzes the reduction of APS in the -+

+

368. P. W. Rohbins and F. Lipmann, JACS 78, 6409 (1956). 369. L. G. Wilson, T. Asahi, and R. S. Bandurski, JBC 236, 1822 (1961). 370. K. Torii and R. S. Bandurski, BBA 136, 286 (1967). 371. W. R. Abrams and J. A. Schiff, Arch. Mikrobiol. 94, 1 (1973). 372. H. D. Peck, Jr., J. Bacteriol. 82, 933 (1961).

SULFATE ESTERS

h3

TRANSFERASES

m

*

0

I

I

n

I

I

-o-s-o-2 ;

;

SUFATE OUTSIDE

1 1 1

[Cor-s-]

-0-2-0-

0

I

SUFATE INSIDE

.

SULFURYLASE

!

P-PI

I

I

-

FERREDOXIN OXIDIZED

AMP

'R-S-

I I

X

I

t

--+-I

E -0-s - 0SULFITE OUTSIDE

FIG.40. A proposed unified scheme of sulfate assimilation in algae. Adenylyl sulfate (APS) transfers the sulfo group via APS-sulfotransferase to form Car-SSO; (Car = carrier), which is reduced further by thiosulfonate reductase to Car-SS- which yields the thiol group of cysteine. In addition, if sulfite is released from Car-S-SOa- (i.e., by thiol or from mutated sites) or if it enters the cell from outside, i t can be reduced via a separate sulfite reductase. From Abrams and Schiff (371).

r

4.

281

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

TABLE XVIII PATHWAY OF SULFATE REDUCTION I N VARIOUS TYPES OF MICROORGANISMS~ APS reductaseb Met,hyl viologenc

Organism

PAPS reductaseb Methyl viologen

NADPH

0.4

2.4

3.7

0.0

3.4

2.8

0 0.6 0 0 0 0 0 0 0 0

0 5.6 0.1 3.3 4.0 12.2 0.1 8.7 0.3 0

1.1 7.3 8.9 0.6 21.4 0 0 0 0

0 0.4 0

0 0.1

1. Assimilatory sulfate reducers

Escherichia coli (grown aerobically) E. coli (grown anaerobically) Yeast Aerobacter aerogenes Proteus mirabilis P . vulgaris Pseudomonas hydrophila Aeromonas punctata Clostridium kluyveri C. pasteurianum Rhodopseudomonas spheroides R . palustris 2. Dissimilatory sulfate reducers Desulfovibrio desulfuricans Clostridium nigrificans Vibrio cholinicus 3. Sulfur oxidizers Thiobacillus thioparus T . thiooxidans T . denitri’cans Chromatium sp.

1,750 310 907

0

640 162 1,260 0.3

From Peck (372). Specific activity is expressed aa nanomoles of acid-volatile sulfur formed per hr per mg protein. No activity was observed with NADPH. a

presence of an appropriate electron donor [reduced methyl viologen, or reduced cytochrome c3 in Desuljovibrio vulgaris (373, 37Sa) ] to sulfite and AMP [Eq. ( S ) ] . APS

+ 2e

S0a2-

+ AMP

(8) It can also catalyze the reverse reaction when ferricyanide or cytochrome c is used as electron acceptor (374, 375). The phosphosulfate bond of 373. H. D. Peck, Jr., Proc. N a t . Acad. Sci. U . S. 45,701 (1959). F!

373a. D. V. DerVartanian and J. LeGall, B B A 346,79 (1974). 374. H. D. Peck, Jr., B B A 49, 621 (1961). 375. R. M. Lyric and I. Suzuki, Can. J. Biochem. 48, 344 (1970).

282

YOUSSEF HATEFI AND DIANA L. STIGGALL

APS is energy-rich (AGO = 18-19 kcal/mole). Therefore, the reversal of reaction (8) is rather interesting, because it can capture oxidation energy and convert it to a biologically utilizable form. For example, the enzyme ADP-sulfurylase can catalyze the synthesis of ADP from APS and inorganic phosphate as shown in Eq. (9) (376, 377). AD P-sulfurylase

APS

+ Pi,

'

ADP

+ SOa2-

(9)

APS reductase is found in dissimilatory sulfate reducing bacteria, such as Desulfovibrio and Desulfotomaculum, in certain Thiobacilli, in Thiocapsa roseopersicina, and in the alga Chlorella pyrenoidosa. Table XIX, compiled by Schiff (378), gives the properties of various APS reductases from plants and microorganisms. I n Thiobacilli and Desulfovibrio, APS reductase constitutes as much as 1-5% of the cell protein, which suggests the important role of this enzyme in the metabolism of these organisms (375). The APS reductase of Desulfovibrio vulgaris has been extensively studied by Peck and his co-workers. The enzyme is reported to have a molecular weight of 220,000, and to contain 1 mole of FAD and 6-8 g-atoms of nonheme iron per mole (379). The oxidized and reduced absorption spectra of the enzyme are shown in Fig. 41. Spectrophotometric studies have shown that in the absence of AMP the enzyme is partially bleached between 350 and 500 nm upon addition of sulfite. The rate of bleaching achieved with sulfite was shown by stopped-flow kinetic measurements to be comparable to the turnover number of the enzyme when sulfite oxidation was assayed in the presence of ferricyanide as electron acceptor. These findings, plus the increased absorption of the sulfitetreated enzyme a t 320 nm, have suggested to Peck and co-workers that sulfite oxidation involves the interaction of sulfite with the enzyme to form a flavin-sulfite adduct in position N-5 of the isoalloxazine ring (379). The authors pointed out that these results are analogous to the data of Massey and co-workers (380, 381) on the effect of sulfite on various flavoproteins. The latter authors found similar spectral changes when sulfite was added to glucose oxidase, D- and L-amino acid oxidases, oxynitrilase, lactate oxidase, and glycollate oxidase. They concluded that the flavoproteins which are capable of interacting with oxygen (APS re376. H. D. Peck, Jr., JBC 237, 198 (1962). 377. H. D. Peck, Jr., T. E. Deacon, and J. T. Davidson, BBA 96, 429 (1965). 378. J. A. Schiff and R. C. Hodson, Annu. Rev. Plant Physiol. 24, 381 (1973). 379. G. B. Michaels, J. T. Davidson, and H. D. Peck, Jr., BBRC 39,321 (1970). 380. V. Massey, F. Miillcr, R. Feldberg, M . Schurnan, P. A . Sullivan, L. G. Howell, S. G. Mayhew, R. G. Matthews, and G. P. Foust, JBC 244, 3999 (1969). 381. F. Miiller and V. Massey, JBC 244, 4007 (1969).

TABLE XIX PROPERTIES OF ADENYLYL SULF.ATEREDUCTASES FROM PLANTS A N D MICROORGANISMS~

Organism Bacteria Desulfovibrio vulgaris T hiobacillus thioparus

Enzyme

Electron donor or acceptor

PH optimum

K,

MW

Remarks Contains 1 mole FAD, 6-8 g-atoms nonheme iron

APS reductase

Fe(CN)?

7.4

S032-,2 m M

220,000

APS reductase

Fe(CN)F

7.4

170,000

APS reductase

Cytochrome c

9.5

APS reductase

Fe(CN)2-

7.2

APS reductase

Fe(CN)63-

8.0

APS reductase

Cytochrome c

9.0

S032-, 2.5 m M AMP, 0.1 m M S032-, 0.017 m M AMP, 0.0025 m M SO3$-, 1.5 m M AMP, 0.041 m M S032-, 1.5 m M AMP, 0.073 m M S032-, 0.093 m M AMP, 0.059 m M

Fungi Saccharom yces cerevisiae

PAPS reductase

NADPH

7.5 (tris)

Algae Chlorella p yrenaidosa

APS reductase

Thiol

Thiobacillus denitri’cans Thiocapsa roseopersicina

a

From Schiff and Hodson (378).

-

170,000 -

180,000 180,000

Contains 1 mole FAD, 8-10 g-atoms nonheme iron Contains 1 mole FAD, 6-11 g-atoms nonheme iron Contains 1 mole FAD, 4 gatoms nonheme iron, 2 gatoms heme iron. Purified 60-80-fold ; homogeneous upon ultracentrifugation Partially purified into 3 fractions A, B, C. Some activity with APS. Fraction A purified 60-fold, fraction C to apparent homogeneity in ultracentrifugation

-

330,000

Partially purified. PAPS is active in the presence of a 3’-nucleotidase

284

YOUSSEF HATEFI AND DIANA L. STIGGALL

o’6 0.5

1 -

0.4

-

w. C

5 03-

::

n

a

0.2-01 0.1

-

300

3K)

460

450

500

550

600

nrn

FIa. 41. Absorption spectrum of purified APS reductase from Desulfovibrio vulA : difference spectrum obtained from tracing of “oxidized” and “reduced” enzyme. Insert B: spectrum obtained after boiling APS reductase and removing protein by centrifugation. From Peck et al. (377). garis. The enzyme concentration was 2.5 mg/ml. Insert

ductase reacts slowly with oxygen) can form a flavin-sulfite adduct, and that the N-5 position of the isoalloxazine ring is very likely involved. Addition of AMP to the sulfite-treated APS reductase resulted in further bleaching between 350 and 500 nm. Peck et al. (382, 383) have shown by EPR spectroscopy near liquid helium temperature that addition of either sulfite or AMP alone does not result in the formation of an iron signal a t g = 1.94. However, when AMP and sulfite are added together, a g = 1.94 signal is produced, which is approximately 80% of that obtained when the enzyme is reduced with dithionite. Thus, the authors suggested that APS reductase catalyzes an intramolecular electron transfer during sulfite oxidation as shown in Fig. 42 from Peck et al. ( 382).

Whereas Peck and his co-workers have not reported the presence of acid-labile sulfide in the APS reductase of D.vulgaris, Lyric and Suzuki (376) have shown that the enzyme from Thiobacillus thioparus contains 4-5 moles of labile sulfide per mole. The T . thioparus enzyme appears to have a molecular weight of 170,000, and contains, in addition to labile sulfide, 1 mole of FAD and 8-10 g-atoms of iron per mole. That the en382. H. D. Peck, Jr., R. Bramlett, and D. V. DerVartanian, 2. Nuturforsch. B 27, 1084 (1972). 383. R. N. Bramlett and H. D. Peck, Fed. Proc., Fed. Amer. SOC.E z p . Biol. 32, 668 (1973).

4.

METAL-COKTAINING FLAVOPROTEIN DEHYDROGENASES

X

=

285

nonheme iron centers

FIG.42. A proposed mechanism for APS reductase. From Peck e t al. (388).

zyme of Peck et al. very likely contains labile sulfide is suggested both by its absorption spectrum and by its characteristic iron-sulfur signal centered a t g = 1.94. Another APS reductase of interest is that which has been isolated by Triiper and Roger (384) from Thiocapsa roseopersicina. The enzyme is reported to have a molecular weight of 180,000 and to contain 1 mole of flavin (presumably FAD), 4 g-atoms of nonheme iron, 6 moles of labile sulfide, and 2 c-type hemes per mole. The spectral properties of the enzyme are shown in Fig. 43. It utilizes cytochrome c and ferricyanide as 0.7

1

iL17nm

Wavelength (nm)

FIG.43. Absorption spectra of the purified APS reductase from Thiocapsa roseopersicina: ox, oxidized enzyme; red, enzyme reduced with 1 mg sodium dithionite per ml. From Triiper and Rogers ( 3 8 4 ) . 384. H. G. Truper and L. A. Rogers, J. Bacterial. 108, 1112 (1971).

286

YOUSSEF HATEFI AND DIANA L. STIGGALL

electron acceptors, and the reaction to cytochrome c is especially sensitive to thiol inhibitors. The heme groups of the enzyme are suggested to be involved in electron transfer from sulfite to added cytochrome c. However, it has not been shown that these heme groups can be reduced by treatment of the enzyme with substrate ( 3 8 4 ~ ) .

IX. Sulfite Reductases (H,S:NADPH Oxidoreductases) (EC 1.8.1.2)

As pointed out in the preceding section, sulfate assimilation in yeast has been shown to involve the activation of sulfate by ATP successively to adenosine 5’-phosphosulfate and once again to 3’-phosphoadenosine 5’-phosphosulfate. The latter is then reduced in the presence of NADPH to sulfite and 3’,5’-diphosphoadenosine (372). Enzymes catalyzing the 6-electron reduction of sulfite to sulfide have been observed in bacteria (339,385-397), yeast (398-401), fungi (402-404), and higher plants (359, 405). These enzymes may be divided into two classes depending on 384a. Dr. H. G. Truper has informed us that the APS reductase of Chlorobium limicola, recently purified in his laboratory, does not contain any heme groups, but is otherwise similar to the APS reductases of sulfate reducing bacteria and Thiobacilli. 385. M. Ishimoto, J. Koyama, and Y. Nagai, J . Biochem. (Tokyo) 42, 41 (1955). 386. J. Mager, BBA 41,553 (1960). 387. J. Dreyfuss and K. J. Monty, JBC 238, 3781 (1963). 388. J. M. Akagi, BBRC 21, 72 (1965). 389. J. LeGall and N. Dragoni, BBRC 23, 145 (1966). 390. L. M. Siegel and H. Kamin, in “Flavins and Flavoproteins,” 2nd Int. Symp. (K. Yagi, ed.), p. 15.Univ. Park Press, Baltimore, Maryland. 1968. 391. N. Gilboa-Garber and J. Mager, BBA 220,602 (1970). 392. P. A. Trudinger, J . Bncteriol. 104, 158 (1970). 393. W.D. Hoeksema and D. E. Schoenhard. J . Bacteriol. 108, 154 (1971). 394. L. M.Siegel, H. Kamin, D. C. Rueger, R. P. Presswood, and Q. H. Gibson, in “Flavins and Flavoproteins,” 3rd Int. Symp. (H. Kamin, ed.), p. 523. Univ. Park Press, Baltimore, Maryland, 1971. 395. K. Kobayashi, E.Takahashi, and M. Ishimoto, J . Biochem. (Tokyo) 72, 879 (1972). 396. J.-P. Lee, J. LeGall, and H. D. Peck, Jr., J . Bacteiiol. 115, 529 (1973). 397. L. M. Siegel, M. J. Murphy, and H. Kamin, JBC 248, 251 (1973). 398. T. Wainwright, BJ 83, 39P (1962). 399. N. Naiki, Plant Cell Physiol. 6,179 (1965). 400. A. Yoshimoto and R. Sato, BBA 153, 555 (1968). 401. K. Prabhakararao and D. J. D. Nicholas, BBA 180,253 (1969). 402. A. Yoshimoto, T. Nakamura, and R . Sato, J . Biochem. (Tokyo) 50,553 (1961). 403. A. Yoshimoto, T . Nakamura, and R. Sato, J . Biochem. (Tokyo) 62, 756 (1967). 404. L. M. Siegel, F. J. Leinweber, and K. J. Monty, JBC 240, 2705 (1965). 405. G. Tamura, J . Biochem. (Tokyo) 57,207 (1965).

4. METAL-CONTAINING

FLAVOPROTEIN DEHYDROGENASES

287

COOH I

FIQ.44. Postulated structural formula for the siroheme prosthetic group. From Murphy et al. (418).

whether or not they can use reduced nicotinamide adenine dinucleotide (specifically NADPH) for the reduction of sulfite. The NADPH-sulfite reductases appear to contain flavin, nonheme iron, acid-labile sulfide, and a novel heme (extractable by acid acetone) of the isobacteriochlorin type with characteristic a-absorption peak a t 582-589 nm. This heme, in which two adjacent pyrrole rings are reduced, has been named “siroheme” (Fig. 44). The sulfite reductases, which cannot utilize NADPH as reductant, are generally of smaller molecular weight, do not require flavin, but exhibit the cytochrome-like absorption peaks comparable to those of the siroheme-containing enzymes. Sulfite reduction by this group of enzymes is usually studied in the presence of appropriate dyes (e.g., reduced methyl viologen) as electron donors. Enzymic and genetic studies have suggested that NADPH-sulfite reductases are composed of a flavoprotein (NADPH dehydrogenase) , and a hemoprotein (sulfite reductase) which can utilize reduced methyl viologen as electron donor. A. NADPH-SULFITEREDUCTASES NADPH-sulfite reductases are found in E . coli (386, 390, 391, 397, 406-416)) Salmonella typhimurium (587, 394, 417-419)) yeast (598-401, 406. F. J. Leinweber and K. J. Monty, BBA 63, 171 (1961). 407. L. M. Siegel and H. Kamin, “Methods in Enzymology,” Vol. 17B, p. 539, 1971. 408. L. M. Siegel, E. J. Faeder, and H. Kamin, 2.Naturjorsch. B 27,1087 (1972).

288

YOUSSEF HATEFI AND DIANA L. STIGGALL

420-424), and Neurospora crassa (404, 425, 426). The E. coli enzyme

has been purified and extensively studied by Kamin, Siegel, and their colleagues (390, 397, 407-416). The enzyme has a molecular weight of 670,000, and contains 4 moles of FAD, 4 moles of FMN, 20-21 g-atoms of iron, 14-15 moles of acid-labile sulfide, and 3 4 moles of heme per 670,000 g protein (390, 397). The absorption spectrum of E. c d i NADPH-sulfite reductase is shown in Fig. 45. The oxidized enzyme (trace A ) has absorption maxima a t 278, 386, 455, 587, and 714 nm. The 455-nm peak results largely from flavin and is bleached upon treatment of the enzyme with NADPH (trace B) or dithionite (trace C ) . Electron paramagnetic resonance studies have shown a signal centered a t g = 6, which is characteristic of high-spin ferric heme, and only under special conditions a signal a t g = 1.94, characteristic of an iron-sulfur center, has been observed (413). The enzyme catalyzes electron transfer from NADPH to sulfite, nitrite, hydroxylamine, cytochrome c, ferricyanide, dichloroindophenol, menadione, FMN, FAD, and molecular oxygen. It is also capable of transhydrogenation from NADPH to acetylpyridine adenine dinucleotide phosphate, and electron transfer from reduced methyl viologen (MVH) to sulfite, nitrite, hydroxylamine, or NADP. All the NADPH-dependent reductions, except the reduction of acetylpyridine adenine dinucleotide phosphate, are inhibited by p-mercuriphenyl sulfonate, but not the reduction of sulfite, nitrite, and hydroxylamine by MVH. The reduction of the latter compounds by NADPH or ~

~~~

409. M. J. Murphy, L. M. Siegel, H. Kamin, D. V. DerVartanian, J.-P. Lee, J. LeGall, and H. D. Peck, Jr., BBRC 54, 82 (1973). 410. M. J. Murphy and L. M. Siegel, JBC 248, 6911 (1973). 411. M. J. Murphy, L. M. Siegel, and H. Kamin, JBC 248, 2801 (1973). 412. M. J. Murphy, L. M. Siegel, S. Tove, and H. Kamin, Proc. Nut. Acad. Sci. U. S. 71, 612 (1974). 413. L. M. Siegel, P. S. Davis, and H. Kamin, JBC 249, 1572 (1974). 414. L. M. Siegel and P. S. Davis, JBC 249, 1587 (1974). 415. E. J. Faeder, P. 9. Davis, and L. M. Siegel, JBC 249, 1599 (1974). 416. M. J. Murphy, L. M. Siegel, and H. Kamin, JBC 249, 1610 (1974). 417. J. Dreyfuss and K. J. Monty, JBC 238, 1019 (1963). 418. L. M. Siegel, E. M. Click, and K. J. Monty, BBRC 17, 125 (1964). 419. L. M. Siegel and K. J. Monty, BBRC 17,201 (1964). 420. T.Wainwright, BJ 103, 56p (1967). 421. A. Yoshimoto and R. Sato, BBA 153,576 (1968). 422. K.Prabhakararao and D. J. D. Nicholas, BBA 218, 122 (1970). 423. A. Yoshimoto and R. Sato, BBA 220, 190 (1970). 424. A. Yoshimoto, N. Naiki, and R. Sato, “Methods in Enzymology,” Vol. 17B, p. 520, 1971. 425. F. J. Leinweber, L. M. Siegel, and K. J. Monty, JBC 240, 2699 (1965). 426. F.J. Leinweber and K. J. Monty, JBC 240, 782 (1965).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

so

I 400

I 450

I

MO

I 5

~

I

I

)600

650

289

mo

Wavelength (nm)

FIG.45. Absorption spectra of E . coli sulfite reductase in the presence of reducing agents. All experiments contained enzyme at a final concentration of 1.54 p M in the sample cell. Spectra were recorded versus a buffer blank as soon as possible after addition of components. A, enzyme in buffer; B, enzyme plus 0.3 mM NADPH (0.1 ml of 23.1 fiM enzyme was added to 1.4 ml of a solution of NADPH which had been bubbled with N, for 30 min); and C, enzyme plus sodium dithionite. From Siege1 et al. (397).

MVH is inhibited by CO, cyanide, arsenite, and sulfide. Carbon monoxide, cyanide, and arsenite react only with the reduced enzyme. Spectral modifications of the heme and other results have indicated that the heme is the site of action of these inhibitors as well as the site a t which sulfite, nitrite, and hydroxylamine are reduced. The Michaelis constants of the enzyme for sulfite and NADPH are both about 4-5 ,AM. Treatment of E . coli sulfite reductase with p-mercuriphenyl sulfonate results in the specific release of F M N from the enzyme (390). FMNdepleted sulfite reductase can be prepared also by photodestruction of FMN. The enzyme-FMN dissociation constant is 10 n M a t 2 5 O , and light irradiation can deplete the enzyme of F M N by destroying the released flavin. These treatments do not lead to removal or destruction of other components of the enzyme. The FMN-depleted enzyme is no longer capable of NADPH-dependent reduction of sulfite, nitrite, hydroxylamine,

290

YOUSSEF HATEFI AND DIANA L. STIGGALL

and diaphorase-type acceptors such as ferricyanide, cytochrome c , and menadione. However, it is fully capable of the reduction of acetylpyridine adenine dinucleotide phosphate by NADPH, and the reduction of sulfite, hydroxylamine, and nitrite by MVH. Further, the remaining flavin (essentially FAD), but not the heme, is still reducible by NADPH as rapidly as the most rapidly reduced flavin of the native enzyme (k = 190 sec-l) . These results and kinetic studies (416) have indicated that FAD is probably the first acceptor of reducing equivalents from NADPH, that FMN is the link between FAD and heme as well as the site of reduction of diaphorase-type acceptors, and finally that the heme is the last component of the enzyme t o be reduced. The FMN-depleted sulfite reductase can be reactivated by added FMN, FAD, and a number of other flavins (413, 415).

By treatment with 5 M urea and chromatography on DEAE-cellulose, it has been possible to dissociate the E. coli NADPH-sulfite reductase into a flavoprotein and a hemoprotein fraction. The flavoprotein fraction has been shown to be an octamer of a single polypeptide of molecular weight 58,000-60,000 and to contain F M N and FAD in equimolar amounts, but no heme, nonheme iron, or labile sulfide. The hemoprotein fraction is a tetramer of a polypeptide of molecular weight 54,000-57,000, and contains heme, nonheme iron, and labile sulfide, but no flavin. Thus NADPH-sulfite reductase is considered to be an enzyme of asp4 subunit composition. The amino acid composition of the whole enzyme and the flavoprotein and hemoprotein fractions have been determined (414). The hemoprotein fraction has no NADPH-dependent activities, but reduces sulfite in the presence of MVH. The flavoprotein fraction catalyzes electron transfer from NADPH to diaphorase-type acceptors and to acetylpyridine adenine dinucleotide phosphate. It does not reduce sulfite, nitrite, or hydroxylarnine with either NADPH or MVH as electron donor. The molecular weight of the flavoprotein is estimated to be 470,000 (two-thirds of the whole enzyme). A similar flavoprotein with a molecular weight of 460,000has been isolated from a S. typhimurium mutant, which requires cysteine for growth. Other genetic data on the S. typhimurium enzyme (994), which appears to be essentially identical to the E. coli sulfite reductase, are in agreement with the above results. Thus mutants lacking the flavoprotein or the hemoprotein component of the enzyme and containing only the appropriate partial activities have been obtained and the respective partial enzymes isolated. The absorption spectra of sulfite reductase preparations from the wild type and from these mutants are shown in Fig. 46, and the proposed structure for the two components of the wild-type enzyme is shown in Fig. 47.Reconstitution of NADPH-sulfite reductase by recombination of the flavoprotein

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

291

FIG.46. Comparison of the absorption spectra of wild-type and mutant (cys G-439 and cys 1-68) sulfite reductases from Salmonella typhimurium. Spectra of S. typhimurium sulfite reductase, cys G-439 NADPH-cytochrome c reductase, and cys 1-68 NADPH-cytochrome c reductase, each dissolved in 0.05 M potassium phosphate buffer, pH 7.7, containing 0.1 mM EDTA, were read against a blank containing only buffer. The spectrum of each enzyme is presented in terms of its millimolar extinction coefficients, assuming 8 moles of flavin per mole of enzyme. Light broken line, calculated difference spectrum between those of wild-type and cys G enzymes when both enzyme solutions contain equal concentrations of flavin. From Siegel et al. (394).

and hemoprotein fractions separated by urea treatment has been achieved (414). Similarly, appropriate partial enzymes isolated from S. typhimuSum mutants have been recombined in vitro to reconstitute NADPHsulfite reductase activity.

cyt c

T

MV

FIG.47. Schematic diagram of the proposed structure and function of the S. typhimurium NADPH-sulfite reductase and its component “subenzymes.” From Siegel et al. (394).

292

YOUSSEF HATEFI AND DIANA L. STIGGALL

co

pCMPS

- -

NADPH. AcPyADP’ , NADP’

FAD

FMN

1

MVH

/

\ *

CN AsO;

,’

Heme

- SO,’-. NOz-. NH,OH

Diaphorase Acceptors. 0 2

FIG.48. Proposed minimum linear scheme of electron flow within the sulfite reductase molecule. The dotted arrow between FMN and heme indicates that the mechanism of electron flow from flavin to heme is not clear. From Siege1 et al. (413).

The above results are summarized in the scheme shown in Fig. 48. Thus, the NADPH-sulfite reductase of enterobacteria appears to be composed of an octameric flavoprotein and a tetrameric hemoprotein, which also contains iron and labile sulfide. The flavoprotein contains 4 moles of FAD and 4 moles of F M N per mole, and appears to bind 1 mole of NADP per mole of FAD. Electron transfer occurs from NADPH to FAD to FMN, and the two flavin sequence is considered to be a device for “stepping down” a two-electron donor, NADPH, to a one-electron acceptor, the heme (413). This is in agreement with the findings that flavin free radical seems to appear after full reduction of the flavins, and that the rate of FH. formation is too slow for the radical to serve as electron donor in the diaphorase reactions (390). The flavoprotein segment catalyzes electron transfer to the hemoprotein, to diaphorase-type acceptors, and to acetylpyridine adenine dinucleotide phosphate. The latter reduction does not require the presence of FMN. The hemoprotein accepts electrons from the flavoprotein or from appropriate dyes and in turn reduces sulfite, nitrite, and hydroxylamine, apparently by direct electron transfer through the heme. The role of iron and labile sulfide is not clear. They might be involved in electronic communication between F M N and the heme. It is also possible that electrons from MVH enter the system a t the level of the iron and labile sulfide. The iron and labile sulfide are likely associated in the form of clusters found in iron-sulfur proteins. However, unlike most iron-sulfur proteins, these clusters appear to be resistant to destruction by mercurials (397). Another interesting point is that it has been suggested that both the heme and the iron-sulfur moieties of NADPH-sulfite reductase have reduction potentials considerably more negative than that of the electron donor, NADPH (415). The NADPH-sulfite reductase of S. cerevisiae (398-401, 420-424) has properties similar to the reductase from enterobacteria. The enzyme has been purified to near homogeneity by Yoshimoto and Sato (400). It contains 1 mole each of FAD and FMW and 5 g-atoms of iron per 350,000

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

293

g protein, and a hemelike chromophore with absorption peaks a t 386 and 587 nm. The oxidized enzyme is greenish yellow, and its spectrum (Fig. 49) is very similar to that of the E . coli sulfite reductase. The yeast enzyme catalyzes the reduction of sulfite, nitrite, and hydroxylamine by NADPH or MVH, and the reduction of diaphorase-type acceptors (Le., quinones and ferric compounds) by NADPH. The NADPH-dependent activities are inhibited by NADP, 2’-AMP, and p-mercuribenzoate. The Michaelis constants for NADPH, sulfite, nitrite, and hydroxylamine are 18-21 p M , 14 p M , 1 mM, and 4.5 mM, respectively. The NADPH-treated enzyme is inhibited by carbon monoxide or cyanide (390,401). The latter treatment results in the formation of a reddish violet color with peaks a t 397 and 411 nm. Cyanide and carbon monoxide are considered to react with the heme moiety and inhibit the enzyme a t the site of reduction of sulfite, nitrite, and hydroxylamine. Yoshimoto and Sat0 (421) have isolated sulfite reductases from four mutants of S. cerevisiae incapable of sulfite assimilation. These enzymes were inactive for sulfite reduction by NADPH, but could utilize MVH as electron donor. All the mutant enzymes contained the chromophore responsible for the 386- and 587-nm peaks, nonheme iron, and labile sulfide. Three of these mutant enzymes contained F M N ; no flavin was detected in the fourth. The sedimentation coefficients of these preparations

1 2a

!i

e

:: 9 1.0

1

300

4 00

500

600

Wavelength (nm)

FIG.49. Absorption spectra of purified NADPH-sulfite reductase from ’Saccharomyces cerevisiae. Curve A : 3.38 mg of enzyme protein in 2.0 ml of 0.3 M potassium phosphate (pH 7.3) containing 1 mM EDTA. Curve B: a mixture containing 3.38 mg of enzyme protein, 0.2 pmole of NADP, 10 pmoles of glucose 6-phosphate, 8 units of glucose-6-phosphate dehydrogenase, 0.3 M potassium phosphate buffer (pH 7.3), and 1 mM EDTA in n final volume of 2.0 ml was incubated anaerobically for 60 min. The reference cell contained all the components except sulfite reductase. From Yoshimoto and Sat0 (400).

294

YOUSSEF HATEFI AND DIANA L. STIGGALL

were 5.1 S for the enzyme lacking FMN, 6.6 S for the three enzymes containing FMN, and 14.8 S for the wild-type enzyme. The authors have concluded, therefore, that the yeast sulfite reductase is composed of a t least three components, one each carrying FAD, FMN, and the heme. The FAD-containing component is the site of NADPH oxidation, and the heme-containing component the site of sulfite (also nitrite and hydroxylamine) reduction, Thus, the mutant enzymes lacking the former component can reduce sulfite only in the presence of an artificial electron donor such as MVH which could reduce both F M N and the heme (Fig. 50). These conclusions regarding the yeast sulfite reductase are essentially in agreement with our current knowledge of the mechanism of sulfite reduction by NADPH and MVH in the E. coli enzyme. Since the reduction of sulfite to sulfide is a six-electron reaction, twoelectron reduction steps may be written as

so2- + (SO,2-)

(sol-)

(15) However, in both the yeast and the E. coli systems, the stoichiometries for NADPH:S2- and S03*-:S2- are 3 : l and 1:1, respectively. These results and the inability to detect 2-electron- and 4-electron-reduced intermediates in these systems have suggested that such intermediates, if present at all, must be firmly held on the surface of the enzyme. It has further been suggested that the presence of multiple flavins and hemes in the enzyme might be a device for achieving a rapid six-electron reduction of sulfite without the release of intermediates (414). This situation is analogous to the four-electron reduction of 0, to 2H,O by cytochrome oxidase and the six-electron reduction of nitrite to ammonia by various assimilatory nitrite reductases. However, unlike cytochrome oxidase,

-

NADPH-reacting site

,4,8

FAD (1)

i

FMN

(n)

+

--f

52-

M$=reacting site

j 587 Chromophore

;

(rn)

Wild-type

Strain 6 , l l and 20

5,,

I

587 Chromphore

cm,

Strain 21

FIG.50. Schematic illustration of a tentative relationship between NADPH-sulfite reductase from the wild-type strain of Saccharomyces cerevisiae and two categories of MVH-sulfite reductases from various mutant strains. From Yoshimoto and Sat0 (421).

4.

METAL-CONTAINING FLAVOPROTEIN DEHYDROGENASES

295

which does not seem to reduce H,O,, several nitrite and sulfite reductases can reduce hydroxylamine to ammonia. This fact indicates that these sulfite and nitrite reductases are capable of catalyzing a two-electron reduction reaction. Indeed, sulfur compounds of oxidation states between sulfite and sulfide have been observed during sulfite reduction by MVHsulfite reductases (395).

B. REDUCED METHYLVIOLOGEN-SULFITE REDUCTASES The methyl viologen-sulfite reductases have been isolated from Aspergillus nidulans (4OZ, .4OoS), Desulfotomaculum nigrificans (392),Desulfovibrio gigas (389, 427), Desulfovibrio vulgaris (396), and from higher plants, such as spinach (359) and Allium odorum (405). These sulfite reductases are incapable of utilizing NADPH or NADH as electron donor. With the possible exception of the sulfite reductase of A . nidulans, they also appear to lack flavin. They all exhibit, however, absorption maxima characteristic of siroheme. Indeed, it has been shown by Murphy and his colleagues (410, 412) that a number of sulfite and nitrite reductases appear to contain the siroheme-type tetrahydroporphyrin. In addition to heme, the sulfite reductase preparation of D.nigrificans also contains nonheme iron, labile sulfide, and zinc. I n general, the methyl viologen-sulfite reductases appear to have lower molecular weights than the NADPH-sulfite reductases. For the enzymes from D . nigrificans, spinach leaves, and D. vulgaris, the reported molecular weights are, respectively, 145,000, 84,000, and 26,800. The physiological electron donor for the MVH-sulfite reductases is not known. However, similar to the ferredoxin-nitrite reductases, certain MVH-sulfite reductases have been shown to use ferredoxin as electron donor (388, 389; see also 373a).

X. Addendum

This additional material is intended to bring to the readers’ attention the recent major developments. For easy identification, the following comments are marked by the same section designations to which they pertain in the text of the chapter.

II,A,3 SDS-Acrylamide gel electrophoresis of the soluble NADH dehydrogenase derived from complex I has shown that this enzyme is composed of two subunits with molecular weights of approximately 28,000 and 56,000 (G. Dooijewaard and E. C. Slater, private communication). 427. H. D. Peck, BBRC 22, 112 (1966).

296

YOUSSEF HATEFI AND DIANA L. STIGGALL

II,A,6 Hatefi et al. (428) have shown that incubation of submitochondrial particles or complex I with 2,3-butanedione, in the presence of borate buffer a t pH 9.0, inhibits the NADPH to NAD transhydrogenase activity with little or no effect on the NADH and NADPH dehydrogenase activities. Presence in the incubation mixture of NAD, NADP, and more effectively NAD NADP, prevented the inhibition of transhydrogenase activity by butanedione. Since butanedione specifically reacts with protein arginyl residues, these findings agreed with the sensitivity of the transhydrogenase activity to trypsin (see Fig. 17) and suggested that the nucleotide-binding site of the transhydrogenase enzyme contains a susceptible arginyl residue. II,A,7 Ohnishi (429) has published revised Em values for the ironsulfur centers of complex I. These values for iron-sulfur centers 1, 2, 3 and 4 a t pH 7.2 are center 1, component a, -380 -+ 20 mV; center 1, component b, -240 -C 20 mV; center 2, -20 20 mV; center 3, -240 20 mV; center 4, -410 f 20 mV. Additional centers ( 5 and 6 with Em value of -260 f 20 mV) are also claimed by Ohnishi to exist in the NADH-ubiquinone segment of the respiratory chain. III,A According to Ohnishi and collaborators (430, 4 3 l ) , succinate dehydrogenase preparations which are capable of electron transfer to the respiratory chain contain 3 iron-sulfur centers, designated centers S-1, S-2 and 5-3. The Em values of these centers a t pH 7.4 have been given as follows: center S-1, O r + 10 mV; center 5-2, -4OOk 15 mV; center 15 mV. Centers S-1 and S-2 are thought to contain one Fe,S2 S-3, +SO cluster each. Center 5-3 is considered to contain one Fe,S, core, and to be the oxygen-sensitive center necessary for electron transfer from succinate dehydrogenase to the respiratory chain. The findings of Ohnishi and co-workers reported here and above have not yet been confirmed by other groups. VI,A A communication from L. Lederer has pointed out that the N-terminus of the intact cytochrome b, chain is Asn, not Glu, (cf. Table XV). Also a note has appeared from the same laboratory (432) on additional similarities between cytochrome b, and liver microsomal cytochrome b, (cf. Section VI,A,2).

+

*

*

*

428. Y. Hatefi, L. Djavadi-Ohaniance, and Y. Galante, in “Electron-Transfer Chains and Oxidative Phosphorylation” (E. Quagliariello, et al., eds.) . NorthHolland Publ., Amsterdam (in press). 429. T. Ohnishi, BBA 387, 475 (1975). 430. T. Ohnishi, D. B. Winter, J. Lim: and T. E. King, BBRC 61, 1017 (1974). 431. T. Ohnishi, J. S. Leigh, D. B. Winter, J. Lim, and T. E. King, BBRC 61, 1026 (1974). 432. B. Guiard, F. Lederer, and C. Jacq, Nature (London) 255, 422 (1975).

4.

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FLAVOPROTEIN DEHYDROGENASES

297

VII According to Husain and Sadana (433), the earlier preparation of Achromobacter fischeri nitrite reductase with a molecular weight of 95,000 2 40,000 (341) was found to be polydisperse. A monodisperse preparation subsequently studied (433) had a molecular weight of 80,000, and was shown to be composed of two subunits of approximate molecular weights of 39,000 with methionine as the sole N-terminal residue. The subunits are stated to be linked together by disulfide bridges. I n a private communication, Henry Kamin has indicated to us that (a) J. Vega, R. H. Garrett and L. Siege1 have demonstrated recently that the nitrite reductase of Neurospora has siroheme as its prosthetic group, and have given us permission to cite this new finding. Kamin has further suggested that we emphasize the fact that enzymic properties and patterns of repression and derepression clearly show that the E. coli nitrite reductase is distinct from the sulfite reductase of this organism, which is also capable of nitrite reduction (Section IX,A). VIII Recent data of Bramlett and Peck (434) indicate that, as predicted in the above review, the adenylyl sulfate reductase of Desulfovibrio vulgaris does contain acid-labile sulfide. The enzyme with a molecular weight of 220,000 has been shown to contain 1 mole of FAD, 12 g-atoms of iron and 12 moles of labile sulfide per mole. I n addition, SDS-gel electrophoresis has revealed the presence of subunits with molecular weights of 20,000 and 72,000. X A new flavoprotein, containing iron and labile sulfide, has been discovered in the mitochondria1 inner membrane independently by Ruzicka and Beinert (435) and Hatefi et al. (436).The protein contains acid-extractable FAD, and 4 g-atoms of iron and 4 moles of labile sulfide per mole of flavin. The molecular properties and the enzymic function of this iron-sulfur flavoprotein are not clear.

ACKNOWLEDGMENTS The authors are grateful to the investigators whose work has been reviewed for kindly providing them with reprints and preprints in advance of publication. They also wish to thank Mrs. C. Schaeggl for typing the manuscript. The work of this laboratory reported in Sections I1 and I11 was supported by USPHS grants AM08126 and CA13609 to Y. H.

M. Husain and J. Sadana, Rur. J. Biochem. 42, 283 (1974). R. N. Bramlett and H. D. Peck, Jr., JBC 250, 2979 (1975). F. J. Ruzirka and H. Beinert, BBRC 66, 622 (1975). Y. Hatefi, Y. M. Galante, D. L. Stiggall, and L. Djavadi-Ohaniance, in “The Structural Basis of Membrane Function” ( Y . Hatefi and L. Djavadi-Ohaniance, edu.), p. 169. Academic Press, New York, 1976. 433. 434. 435. 436.

This Page Intentionally Left Blank

Cytochrome c Oxidase WINSLOW S. CAUGHEY WILLIAM J . WALLACE JOHN A . VOLPE SHINYA YOSHIKAWA 1. Introduction . . . . . . . . . . . . . . . . 299 A . The Role of Cytochrome c Oxidase in Biological Systems . 299

B . History . . . . . . . . . . . . . . . C . The Chemical and Physical Properties of Cytochrome c Oxidase . . . . . . . . . . . . . . D . The Chemistry of Oxygen Reduction . . . . . . I1. Isolation and Characterization . . . . . . . . . . A . Preparation . . . . . . . . . . . . . B . Metal Components . . . . . . . . . . . C . Protein . . . . . . . . . . . . . . . D . Lipids . . . . . . . . . . . . . . . I11. Chemical and Physical Properties . . . . . . . . . A . Models . . . . . . . . . . . . . . . B. Electronic Spectroscopy . . . . . . . . . . C . Ligand Binding Studies . . . . . . . . . . D . Potentiometry . . . . . . . . . . . . . E . Electron Paramagnetic Resonance Studies . . . . F . Interaction of Cytochrome c Oxidase with Cytochrome c G . Kinetic Studies . . . . . . . . . . . . IV . Mechanisms . . . . . . . . . . . . . . . .

.

300

. 301

. . . .

. . .

. . . .

. . . .

302 305 305 307 309 312 313 314 315 319 325 329 334 335 337

I. Introduction

c OXIDASEIN BIOLOGICAL SYSTEMS A. THEROLEOF CYTOCHROME

Cytochrome c oxidase. the terminal oxidase in the respiratory metabolism of all aerobic organisms. plants. animals. yeasts. algae. and some bacteria. is responsible for catalyzing the reduction of dioxygen to water . 299

300

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

The electrons are provided by reduced cytochrome c in the following overall reaction: 02

+ 4 cyt c*+ + 4 H+ -+

2 H20

+ 4 cyt c3+

(1) The free energy developed in oxygen reduction is used to promote oxidative phosphorylation and, in consequence, becomes available, as ATP, to satisfy the energy requirements of the cell. It is not surprising then that this oxidase is found in high concentrations in tissues where the energy requirements are high. Especially high levels have been observed in heart muscle ( I ) , flight muscles of birds (2) and insects (S),red skeletal muscles ( 4 ) , liver mitochondria ( 5 ) , brain gray matter (6),corpora lutea of sheep (Y), the parasitic worm Ascaris (8),and sugar cane roots ( 9 ) .It is estimated that 90% of the energy for heart muscle contraction (1) and 96% of the energy for bird flight muscle contraction (2) is provided through aerobic metabolism via cytochrome c oxidase. Chronic muscular disease gives rise to oxidase depletion (10). A further indication of its importance in the energetics of biological systems is the suggcstion by Malmstrom (11) that 90% of biological oxygen consumption is directed through the oxidase. The locus of this activity is in the inner membrane of mitochondria in eukaryotes and in the plasma membrane of prokaryotes.

B. HISTORY In 1886, MacMunn ( 1 2 ) discovcred the respiratory pigment, myohematin, which was widely distributed in plant and animal tissues. This important observation attracted little attention a t the time of its publication and became cffectively lost in the literature (IS). I n 1925, Keilin (14) 1. D. R. Challoner, N a l w e (London) 217, 78 (1968). 2. A. Tucker, Science 154, 150 (1966). Ser. B 98, 312 (1925). 3. D. Keilin, Proc. R o y . SOC., 4. M. S. Gordon, Science 159, 87 (1968). 5. D. L. Drabkin, Physiol. R e v . 31, 345 (1954). 6. S. Manocha and G. H. Bourne, E x p . Brain Res. 2, 230 (1966). 7. L. Arvy and P. Mauleon, C . R . SOC.Biol. 158, 453 (1964). 8. M. H. Smith, N a l w e (London) 223, 1129 (1969). P . R . 50, 131 (1966). 9. A. G. Alexander, J. Ag?. Unit!. 10. F. W. Booth and J. R. Kelso, Can. J. Physiol. Phurmucol. 51, 679 (1973); V. P. Andrcev, Dokl. Akad. Nauk Beloniss. SSR 17, 470 (1973). 11. B. G. Malmstrom, Quart. Reu. Biophys. 6,389 (1973). 12. C. A. MacMunn, Phil. Trans. R o y . Soc. London 177, 267 (1886). 13. D. Keilin. in “The History of Cell Respiration and Cytochromes” (J. Keilin, ed.), p. 95. Cambridge Univ, Press, London and New York, 1966. 14. D. Keilin, Proc. R o y . SOC.,Ser. B 98, 312 (1925).

5.

CYTOCHROME C OXIDASE

301

rediscovered the MacMunn pigment, proved it to be a mixture‘ of three spectroscopically identifiable components which he named cytochromes a, b, and c, and showed them to be links in the respiratory chain that connected activated substrates to activated dioxygen. Cytochromes a and c showed a special relationship to each other; cytochrome a was the sole physiological oxidizing agent for cytochrome c. Hence, the name cytochrome c oxidase ( 1 5 ) . The ligand binding and autoxidizability studies seemed compatible with the presence of two components, cytochromes a and as, of which only as was considered to be autoxidizable and able to combine with carbon monoxide or cyanide. Keilin and Hartree (16) identified this a3 component spectroscopically with the Atinungsjerment which Warburg had shown on the basis of the photochemical action spectrum of its CO complex to be a heme protein ( 1 7 ) .Further advances in the understanding of this important enzyme were not to come for another 25 years until renewed interest and improved isolation techniques paved the way for further progress. Lemberg (18) and Lemberg and Barrett (19) have summarized the development of this understanding. Less extensive reviews have been provided recently by Wharton (20) (emphasizing the role of copper) , by Nicholls and Chance (21) (emphasizing kinetic measurements) , and by Malmstrom (11) (emphasizing physicochemical measurements). AND PHYSICAL PROPERTIES OF CYTOCHROME c C. THECHEMICAL OXIDASE

The focus of this article will be upon those aspects of the structure and function of cytochrome c oxidase that contribute particularly to an understanding of the chemical events that lead to the reduction of dioxygen to water. This important function is, however, only one aspect of its physiological role. The functioning enzyme is provided with electrons from the electron transport chain by cytochrome c, uses these electrons to reduce dioxygen bound at the active site, communicates the energy released in this reduction to the site of oxidative phosphorylation, 15. 16. 17. 18. 19. 1972. 20. 21.

D. Keilin and E. F. Hartree, Proc. Roy. Soc., Ser. B 121, 173 (1936). D. Keilin and E. F. Hartree, Nature (London) 141, 870 (1938). 0. Warburg and €3. Negelein, Biochem. 2.214,64 (1929). R. Lemberg, Physiol. Rev. 49, 48 (1969). R. Lernberg and J. Barrett, “The Cytochromes.” Academic Press, New York,

D. C. Wharton, Metal Zons Biol. Syst. 3, 157 (1974). P. Nicholls and B. Chance, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 479. Academic Press, New York, 1974.

302

W. S. CAUGHEY, W. J . WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

and is strictly controlled in these functions by respiratory control processes. The structure of the oxidase and its placement in the cell reflect the multiplicity of roles it is required to play. The active site of the enzyme, which contains iron, as heme A, and copper, is the locus for oxygen binding and reduction; thus, it is necessary to provide pathways to get electrons in and energy out of the active site. The multisubunit lipoprotein in which the active site is embedded is itself embedded in the inner mitochondria1 membrane in such a way that it, along with NADH dehydrogenase and possibly cytochrome b, spans the membrane ( 2 2 ) . Thus, a t each site of energy conservation there appears to be an electron carrier which spans the entire membrane. Perhaps this transmembranous configuration provides an extended surface for interaction with electron carriers and may also provide for an important interaction with ATPase. Such an interaction is supported by the observation of Wilson et al. (23,24) that binding of ATP to ATPase influences the redox potentials and ligand binding characteristics of oxidase. Thus, cytochrome c oxidase appears very carefully tuned to a variety of specific tasks. The components are so adjusted that a low energy pathway is available to entering electrons, an efficient nonthermal energy transport pathway is available for energy conservation, and a set of electron donors has been assembled into an array that will permit facile reduction of dioxygen via an efficient low energy pathway that is unprecedented in simple systems. Nonenzymic reduction of dioxygen to water is often slow and usually involves a complex series of steps (25-27). The enzymic reaction is fast and appears to be accomplished in either a single step or in a series of concerted steps; no evidence for intermediate reduction products (i.e., superoxide or peroxide) has been found.

D. THECHEMISTRY OF OXYGEN REDUCTION The chemical inertness of dioxygen a t first seems surprising because the transformation to water is so strongly thermodynamically favorable ( 4 3 0 kcal) (Fig. 1 ) (28, 2 9 ) . However, on the basis of the standard redox potentials, the simplest reduction step, the one-electron step to 22. E. Racker, Hosp. Pract. p. 87 (1974). 23. J. G. Lindsay and D. F. Wilson, Biochemistry 11, 4613 (1972). 24. D. F. Wilson and K. Fairs, A B B 163, 491 (1974). 25. C. T. Mathews and R. G . Robins, Australas. Znst. Mining Met., Proc. C31 242, 47 (1972). 26. D. V. Stynes, H. C. Stynes, J. A. Ibers, and B. R. James, JACS 95, 1142 (1973). 27. I. A. Cohen and W. S. Caughey, Biochemistry 7,636 (1968). 28. M. S. Tsao and W. K. Wilmarth, Advan. Chem. Ser. 36, 113 (1962). 29. K. Sehested, 0. L. Rasmussen, and H. Fricke, J . Phys. Chem. 72,626 (1968).

5.

303

CYTOCHROME C OXIDASE

m

Q-%O$B%

d*%V

B A2OV

+0.82V

FIG.1. Standard oxidation-reduction potentials for the steps involved in the conversion of dioxygen to water at 25” and pH 7. superoxide, is thermodynamically highly disfavored (SO). Hence, reactions involving dioxygen must either have enormous driving energies to go through the superoxide or have access to a two-electron step to peroxide. Although this conclusion depends upon reasoning based upon standard potentials (SO) (all concentrations 1 M and pH 7), i t seems valid since oxygen reduction by a low energy pathway is found to proceed via the two-electron reduction to peroxide as the first recognizable product (31) .

The other property of dioxygen that contributes to the slowness of its reactions is its electronic structure (3.2). I n common with most stable molecules dioxygen has an even number of electrons. Uncommonly, though, the molecule is paramagnetic with two unpaired electrons in the two highest occupied molecular orbitals. Since both peroxide and oxide are completely spin paired, reactions involving dioxygen must involve spin reversal and are therefore spin forbidden and slow. The forbiddenness can be removed if dioxygen can interact with a paramagnetic center to participate in exchange coupling. The transition metal ions frequently have unpaired electrons and turn out to be excellent catalysts for dioxygen reduction. Despite the long history of transition metal ion induced reduction of dioxygen surprisingly little is known about the mechanism of the reaction ( 2 5 ) .Perhaps the most studied of these metal ion oxygenation reactions is that between the nitrogen ligand complexes of cobalt (11) and dioxygen. The traditional method for preparing cobalt (111) ammine complexes was to assemble the desired ligands on cobalt(I1) and oxygenate the solution. Upon long oxygenation a cobalt (111) complex was formed ( 3 3 ) .It is now known that the first step in this reaction is the formation of an unstable Co”02 complex (34) and upon standing, a second cobalt(I1) ion is added to produce the p-peroxo bridged complex [reactions (2) and (3)1. I n the 30. P. M. Wood, FEBS (Fed. Ew. Biochem. Soc.) Lett. 44, 22 (1974) ; P. George, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 3. Wiley, New York, 1965. 31. R. G. Wilkins, Adwnn. Chem. Ser. 100, 111 (1971). 32. H. Taube, J . Gen. Physiol. 49, 29 (1965). 33. A. Werner, 2. Anorg. Chem. 3, 267 (1893). 34. J. Simplicio and R. G. Wilkins, JACS 89,6092 (1967).

304

W. S. CAUGHEY, W. J. WALLACE, J.

A. VOLPE,

AND S. YOSHIKAWA

presence of a second bridging ligand, such as amido or hydroxo, the p-peroxo complex is stabilized and can be isolated (36) [reaction ( 4 ) ] .

con-0,

+ con

-

com-0,

L

corn- 0,0-corn

(3)

0-co"'

coF0NL\ -o,Coul

L = NR, or OR

(4)

I n the absence of the second bridging ligand, further oxidation occurs by a series of, as yet, not completely understood steps (36) and the cobalt (111) product results [reaction ( 5 )1. coin -0,

0-co'"

-

2 Com-OH

(5)

I n a similar way chromium(I1) (37) and copper(1) (38)react with dioxygen by way of a p-peroxo intermediate. And, of special relevance to cytochrome c oxidase, the reactions of hemes (27, 39, 40) as well as simple aquo ferrous iron (26, 41) with dioxygen seem to proceed through two-electron reduction of bridged intermediates. Thus, dipyridine hemes react cleanly with dioxygen to form py-FeII-py-

(7)

+ py-Fen-py-Fem-O,-Fem-py

(8)

py-FenI-O~Fem-pyt++Fdn-O-Fenl Fem-O, FeIV-0.

(6)

+ py

-py-Fen-O,

py-Fen py-FeI1-O0,

+ 0,

py-Fen

0 Fen'

+

-2

+

2 py

+

py

(9)

Fe"-O.

py-Ferl-Feln-O-Fem

(11)

35. M. Mori and J. A. Weil, JACS 89, 3732 (1967). 36. L. G. Stadtherr, R. Prados, and R. B. Martin, Znorg. Chem. 12,1814 (1973). 37. T. B. Joyner and W. K. Wilmarth, JACS 83, 516 (1961). 38. C. DeMarco, S. Dupre, C. Crifo, G. Rotilio, and D. Cavallini, ABB 144, 496 (1971). 39. I. A. Cohen and W. S. Caughey, in "Hemes and Hemoproteins" (B. Chance, R. W. Estabrook, and T. Yonetani, eds.), p. 577. Academic Press, New York, 1966. 40. W. S. Caughey, J. L. Davies, W. H. Fuchsman, and S. McCoy, in "Structure and Function of Cytochromes" (K. Okunuki, M. D. Kamen, and I. Sekuzu, eds.), p. 20. Univ. of Tokyo Press, Tokyo, 1968. 41. P. George, JCS p. 4349 (1954).

5.

305

CYTOCHROME C OXIDASE

p-oxobishemins (42) as shown by reactions (6) through ( 9 ) . Kinetic data fully support reactions ( 6 ) through (8) but have not yet provided information on the steps from the bridged oxygen species (presumably p-peroxobishemin) to p-0x0 dimer. When the solvent medium is able to provide protons, solvolysis to produce H,O, would likely follow formation of the p-peroxobishemins complex and sequence ( 6 ) , ( 7 ) , and (8). However, where solvolysis cannot occur, as in aprotic solvents, formation of the ferry1 (FexVO)intermediate as suggested by reactions (10) and (11) is reasonable (40). But in no case does our certain knowledge about the mechanism of the reduction reaction extend beyond the bridged dimer. Apparently the peroxide formed in the initial reaction is a kinetically inert (fully spin-paired) molecule that does not readily accept additional electrons despite the favorable thermodynamics for reduction to water (Fig. 1). The most commonly suggested mechanism (43) for the subsequent reduction steps in protic media is shown by reactions (12), (13), and (14). Mi--O\ MI- 0, 0-H MI--+

H+

0-MI

+

M;+

+

H,O, -MIOH

MI--,

0- H

+

M:

+ M,OH+

M,-o+

+

0,

+

H+

(13)

(14)

It is anticipated that despite the specially favored environment provided for oxygen reduction by the protein the fundamental principles of chemistry in simple systems will apply to the enzyme. Thus, any proposed mechanism for the enzymic reduction of dioxygen will have to accommodate two electron steps leading sequentially to peroxide and water and provide a means to overcome the characteristic stability of the peroxide intermediate. II. Isolation and Characterization

A. PREPARATION In general cytochrome c oxidase has been isolated from mitochondria or mitochondria1 fragments by initial extraction of proteins with a sur42. N. Sadasivan, H. I. Eberspaecher, W. H. Fuchsman, and W. S. Caughey, Biochemistry 8,534 (1969). 43. M. L. Kremer, Trans. Faraday Soc. 59, 2535 (1963); E. Zidoni and M. L. Kremer, ABB 161, 658 (1974).

SPECIFIC

ACTIVITIESO F CYTOCHROME

C

TABLE I OXIDASE PREPAR.%TIONS

FROM

DIFFERENTISOL.4TION

Conditions of assay

Preparation

Buffer

PH

Yonetani Griffiths and Wharton Okunuki et al. Horie and Morrison Sun and Jacobs Wainio Fowler et al. Kuboyama et al. I-olpe and Caughey

0.050 M phosphate

5.9 7.0 5.95

a

Data not provided.

0

0.075 M phosphate

0 . 1 0 M phosphate 0.070 M phosphate 0.10 M phosphate

pl

Specific activity Initial concn. cytoTemp. chrome c*+ 25 38 25

D

6.0 6.0 6.0 5.7 5.9

4

PROCEDURES

25 25 25 23 22

15 18 15 24 15

pg

protein/ml 6.6 0.55 1.81 1.22

( I

22 20 15

0.5 0.5 0.33

s-l/mg protein/3 ml 4.50 2.70 5.20 4.70 0.08 6.7 14.3 16.0 15.4

Ref.

44 45 46

47 48 43 60 51 5.9

5.

CYTOCHROME C OXIDASE

307

face-active agent followed by removal of contaminating detergent and protein (Table I ) , (44-52). Procedures differ in the nature and amount of detergent used and retained. Recently, a preparation with high s o h bility in detergent-free media was reported ( 5 2 ) . The preparations obtained vary widely in activity (Table I ) , and this suggests that the detailed manipulative history is critical to the “nativeness” of the isolated enzyme. Nevertheless, there is good reason to believe that the oxidases isolated by many procedures are similar, but not identical, in response to chemical and physical probes (18).

B. METALCOMPONENTS

It is now widely agreed that both copper and iron are essential components (52-56). The metal content (11 nmoles/mg protein) and the iron to copper ratio (1.0) are well established for the bovine enzyme, whereas in yeast the reported metal contents are higher and more variable (5-15 nmoles of iron per milligram of protein) and the copper to iron ratio is greater than unity (-1.5) (Table 11) (44-46, 48-52, 57-59). The iron is present as the unusual heme, heme A, with an apparently unique structure (Fig. 2) (60). The coordination environment of copper is far less clear, but the easy reducibility of copper seems to require a ligand envi44. T. Yonetani, JBC 236, 1680 (1961). 45. D. E. Griffiths and D. C. Wharton, JBC 236, 1850 (1961). 46. K. Okunuki, I. Sekuzu, T. Yonetani, and S. Takemori, J . Biochem. (Tokyo) 45, 847 (1958). 47. S. Horie and M. Morrison, JBC 238, 1855 (1963). 48. F. F. Sun and E. E. Jacobs, BBA 143,639 (1967). 49. W. W. Wainio, JBC 239, 1402 (1964). 50. L. R . Fowler, S. W. Richardson, and Y. Hatefi, BBA 64, 170 (1962). 51. M. Kuboyama, F. C. Yong, and T. E. King, JBC 247, 6375 (1972). 52. J. A. Volpe and W. S. Caughey, BBRC 61,502 (1974). 53. D. E. Griffiths and D. C. Wharton, JBC 236,1857 (1961). 54. H. Beinert, in “Biochemistry of Copper” (J. Peisach, P. Aisen, and W. E. Blumberg, eds.), p. 213. Academic Press, New York, 1966. 55. E. C. Slater, B. F. Van Gelder, and K. Minnaert, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 2, p. 667. Wiley, New York, 1965. 56. W. W. Wainio, in “Oxidasas and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 2, p. 622. Wiley, New York, 1965. 57. M. S.Rubin and A. Tzagoloff, JBC 248, 4269 (1973. 58. T. L. Mason, R. 0. Poyton, D. C. Wharton, and G. Schatz, JBC 248, 1346 ( 1973). 59. P. G. Shakespeare and H. R. Mahler, JBC 246,7649 (1971). 60. W. S. Caughey, G. A. Smythe, D. H. O’Keeffe, J. Maskasky, and M. L. Smith, JBC 250, 7602 (1975).

308

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

TABLE I1 COMPOSITION OF CYTOCHROMF: c OXIDASE PREPARATIONS

Preparation

Species

Yonetani Griffiths and Wharton Okunuki et al. Wainio Fowler et al. Sun,and Jacobs Kuboyama et al. Volpe and Caughey Rubin and Tzagoloff Mason et al. Shakespeare and Mahler

Bovine Bovine

0

Bovine Bovine Bovine Bovine Bovine Bovine

Copper (nmole/mg protein)

Iron (nmole/mg protein)

a

7.2 8.2-9.4

9.2-10.6 a

11.0

10.0 11.5 8.4-8.7 8.2 11.1 10.9

Yeast

21.3

15.0

Yeast Yeast

15.8 6.2-11.6

9.4 5.5-7.2

(I

9.4 n

11.8

Phospholipid (%) Ref. 10 24 (I

9 a

22 20 20 3.8

44 46 46 49 60 48 61

68

67

2

68

a

69

Data not provided.

ronment that stabilizes Cu(1) relative to Cu(I1). Thus, sulfur might serve as a ligating atom (as in a disulfide) (61) as may an interaction with the T system of the long side chain of the heme (6‘2). The latter possibility is attractive because it provides a role for the uiiusual side chain by affording a mechanism for electronic coupling between iron and copper (Fig. 3 ) . Whatever the nature of the binding forces, the copper must be well sequestered by the protein because it is not readily moved by the usual complexing agents [EDTA, BCS (62a), and CN-] (53),and there is no evidence that any ligand or any inhibitor binds directly to copper a t the active site (53, 63-65). In addition, it is now clear that there are two quite distinct kinds of copper. One kind is observed by electron paramagnetic resonance (EPR) to be rapidly reduced. The other 61. P. Hemmerich, in ‘LBiocl~emistry of Copper” (J. Peisach, P. Aisen, and W. E. Blumberg, eds.), p. 15.Academic Press, New York, 1966. 62. W. S. Caughey, Adunn. Chem. Ser. 100,248 (1971). 62a. BCS = bathocuproin sulfonate or 2,9-dimethy1-4,7-diphenyl-l,lO-phenanthroline sodium disulfonate. 63. Q.H.Gibson and C. Greenwood, JBC 240,2694 (1965). 64. K.J. H.Van Buuren, P. F. Zuurendonk, B. F. Van Gelder, and A. 0. Muijsers, BBA 256, 243 (1972). 65. J. A. Volpe, M. C. O’Toole, and W. S. Caughey, BBRC 62, 48 (1975).

5.

309

CYTOCHROME C OXIDASE

H2

F\

Y3

7H2 04\OH

04bi

FIG.2. The structure of heme A.

1

FIG.3. Schematic representation of a possible conformation of the 2-alkyl group of heme A.

kind is not detected directly spectroscopically, but is reduced slowly ( 6 6 ) . Similar observations suggest that, despite the fact that the only heme is heme A, two kinds of iron are present ( 6 6 ) .These observations must mean that the minimal functional unit contains two iron atoms and two copper atoms.

C. PROTEIN The 11 nmoles of iron per milligram of protein of bovine preparations corresponds t o an empirical molecular weight of about -90,000. When 20% lipid is added a total of 108,000 is obtained. If the functional unit contains two iron and two copper atoms, the minimum molecular weight then becomes -200,000 with the additional possibility that multiples of 66. C. R. Hartzell, R. E. Hansen, and H. Beinert, Proc. N u t . Acad. Sci. U S . 70, 2477 (1973).

310

W . S. CAUGHEY, W. J . WALLACE, J. A. VOLPE, AND 9. YOSHIKAWA

TABLE I11 AMINO ACID COMPOSITION OF CYTOCHROME c OXIDASE Number of residues Amino acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Cysteine Valine Methionine Isoleucine Leucine Tyrosine" Phenylalanine NH8 Tryptophan Ethanolamine Total residues Molecular weightc Molecular weightd a

Kuboyama el al."

Matsubara rt alSh

28 20 21 52 51 53 52 48 53 55 7 45 13 40 79 29 43 63 27

39 30 31 60 53 A4 60 46 59 62 7 51 35 43 87 33 47 59 30

716 80,054 89,400

827 93,802

From Kuboyama et al. (61).

* From Matsubara rt al. (YO). c

d

Based on amino acid composition; the prosthetic groups have not been included. Based on heme content.

this minimum unit could also be observed. Direct physical measurements of the molecular weight have led to values of 72,000 for an inactive preparation (67) and about 200,000 (68, 69) and 430,000 (51) for active preparations. An empirical molecular weight for protein alone of about 90,000 (51,70) was suggested by amino acid analysis of two preparations 67. R S. Criddle and R. M. Bock, BBRC 1,138 (1959). 68. A. Tzagoloff, P. C. Yang, D. C. Wharton, and J. S. Reiske, BBA 96, 1 (1965); W. W. Wainio, T. Laskawska-Klita, J. Rosmm, and D. Grebner, J. Bioenerg. 4, 455 (1973). 69. B. Love, S. H. P. Chan, and E. Stotz, JBC 245,6664 (1970). 70. H. Matsubara, Y. Orii, and K. Okunuki, BBA 97,61 (1965).

5.

CYTOCHROME C OXIDASE

311

(Table 111).Extraneous protein and variable amounts of lipid can result in differences in determined molecular weight values. However, a molecular weight of about 200,000 is widely accepted for functioning oxidase with 20% lipid. Nevertheless, treatment with alkali (69) or with sodium dodecyl sulfate (71) has been reported to result in a molecular weight of about 100,000 for a monomer (72) which exhibited activity. Such a catalytically active monomer has been difficult to reconcile with different roles for each of two coppers and of two hemes. For this reason, the dimer is still broadly considered the normal form of the native oxidase. Subdivision below the monomer level occurs in the presence of sodium dodecyl sulfate and thiols (mercaptoethanol). The oxidase is thus identified as a multisubunit protein. Both yeast (57, 73) and Neurosporu crussu (74) oxidases were shown to be composed of seven subunits. Bovine heart oxidase, on the other hand, has been reported to have between two (75, 76) and six (57, 76) subunits. The subunits from yeast have molecular weights in the range of I, 40,000; 11, 33,000; 111, 22,000; IV, 14,000; V, 12,700; VI, 12,700; and VII, 4,600 (57, 7 7 ) . The situation for bovine heart is less clear but the six subunits are reported to have molecular weights around I, 40,000; 11, 25,000; 111, 19,000; IV, 14,000; V, 10,000; and VI, 8,000 ( 7 6 ) .When fewer than six subunits are found, their molecular weights invariably correspond to some of the six reported (75-78). The subunit sizes differ for yeast and bovine heart ( 5 7 ) .That the protein compositions differ is also reflected in the failure of antibodies against subunits I1 and VI of yeast oxidase to cross-react with bovine heart oxidase ( 7 9 ) . In both yeast (80) and Neurosporu crassa (74) biosynthetic studies have shown that the four small polypeptides are of cytoplasmic origin while the three large polypeptides are of mitochondria1 origin. Similar studies are unavailable for bovine heart. The polypeptide associations of the copper and heme are still far from clear, but hints that the metals 71. Y. Orii, Y. Matsumura, and K. Okunuki, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 666. Univ. Park Press, Baltimore, Maryland, 1973. 72. S. H. P. Chan, B. Love, and E. Stotz, JBC 245,6669 (1970). 73. R. 0. Poyton and G. Schatz, JRC 250,752 (1975). 74. W. Sebald, W. Machleidt, and J. Otto, Eur. J. Biochem. 38, 311 (1973). 75. H. Komai and R. A. Capaldi, FEBS ( F e d . Eur. Biochem. SOC.) Lett. 30, 273 (1973). 76. T. Yamamoto and Y. Orii, J. Biochem. ( T o k y o ) 75, 1081 (1974). 77. R. A . Capaldi and H. Hayashi, FEBS (Fed. Eur. Biochem. SOC.) Lett. 26, 261 (1972). 78. J. J. Keirns, C. S. Yang, and M . V. Gilmour, BBRC 45,835 (1971). 79. R. 0. Poyton and G. Schatz, JBC 250, 762 (1975). 80. T. L. Mason and G. Schatz, JBC 248, 1355 (1973).

312

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S . YOSHIKAWA

are coupled with the low molecular weight subunits have appeared (76, 78, 81). The picture that emerges, albeit imperfectly, from these studies is of a multisubunit protein synthesized partly in the cytoplasm and partly in the mitochondria (@). The subunits are then assembled in the mitochondria1 membrane, where the functioning enzyme resides, with the times of incorporation of copper, heme, and lipid quite uncertain.

D. LIPIDS The role of the lipid component remains in doubt but is becoming somewhat clearer. The phospholipid content is reported for several preparations (Table IV) (83-86).Somewhat more diphosphatidylglycerol and TABLE I V COMPOSITION OF CYTOCHROME C OXIDASK MITOCHONDRIA FROM BOVINEHEART

PHOSPHOLIPID

AND

Percent of total lipid Phosphatidylcholine

Phosphatidylethanolamine

10 5

37 38

31 30

16 18

83

9 11

26 27 46 32

25 21 26 30

30 31 11 30 73

83 86 86

PhosphatidylPreparation investigators inositol Mitochondria Fleischer et al. Awasthi et al. Cytochrome c oxidase Fleischer et al. Brierley and Merola Yu el al. Awasthi et al. Awasthi et al.@

Diphosphatidylglycerol Ref.

84

84 84

"Lipid-free" cytochrome c oxidase was obtained by treatment of mitochondria with Triton X-100 and Triton X-114. The remaining lipids (27%) were not identified but were possible breakdown products of diphosphatidylglycerol. 0

81. G.Schatz, G.S. P. Groot, T. Mason, W. Rouslin, D. C. Wharton, and J. Saltzgaber, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 31,21 (1972). 82. W. L. Chen and F. C. Charalampous, BBA 294,329 (1973). 83. S. Fleischer, H.Klouwen, G. Brierley, E. Carpenter, and T. Moran, JBC 236, 2936 (1961). 84. Y . C. Awasthi, T. F. Chuang, T. W. Keenan, and F. L. Crane, BBA 226, 42 (1971). 85. G.P. Brierley and A. J. Merola, BBA 64,205 (1962) 86. C. Yu, L.Yu, and T. E. King, JBC 250,1383 (1975).

5.

CYTOCHROME C OXIDASE

313

somewhat less phosphatidylethanolamine are found in the oxidase than in whole mitochondria, but the amount and kind of phopholipid in an enzyme preparation is dependent upon the treatment it has received ( 8 4 ) . Conventional purification procedures preferentially remove phosphatidylinositol, phosphatidylcholine, and phosphatidylethanolamine. These lipids are extracted readily with acetone (84, 8 7 ) , methanol chloroform ( 8 8 ) , or nonionic detergents such as Triton X-100 or X-114 (89, 90) to leave behind a small amount of lipid (-1.6-1.7 pg phosphorus per milligram of protein) that is largely cardiolipin (Table IV) ( 8 4 ) . This residual cardiolipin (about 2 moles/mole of oxidase) (87) is both difficult t o extract (only chloroform-methanol-ammonia proved efficacious) and insensitive to digestion by phospholipase A under conditions where the phospholipids of mitochondria are readily hydrolyzed ( 8 4 ) . As the other phospholipids are removed, the enzyme becomes progressively less active (87), but the activity is restored by adding back purified lipids, mitochondrial lipid, or detergent (such as Emasol) (84, 85, 8 7 ) . However, if the residual cardiolipin is removed, restoration of activity is minimal ( 8 4 ) . Such studies suggest a t least three ways in which phospholipid interacts with and affects the activity of the enzyme: (1) in the incorporation of the oxidase into membranes, with a concomitant increase in the accessibility of the active site (91) ; (2) in the formation of a complex between cytochrome c and the oxidase (84); and ( 3 ) in the stabilization of active conformations ( 9 2 ) . It is evident that particular attention must be paid to both lipid and detergent contents before many of the differences in properties among preparations can be rationalized.

111. Chemical and Physical Properties

The chemical and physical properties of cytochrome c oxidase have been widely studied in intact mitochondria, in mitochondria1 particles, and as the isolated enzyme. Aside from variations in activity mentioned above and the recently observed effect of detergent on intensities of electronic spectra (52), the properties have proved remarkably insensitive 87. W. L. Zahler and S. Fleischer, J . Bioenerg. 2, 209 (1971). 88. Y. C. Awasthi, T. F. Chuang, T. W. Keenan, and F. L. Crane, BBRC 39, 822 (1970). 89. E. E. Jacobs, F. H. Kirkpatrick, Jr., E. C. Andrews, W. Cunningham, and F. L. Crane, BBRC 25, 96 (1966). 90. F. F. Sun, K. S. Preebindowski, F. L. Crane, and E. E. Jacobs, BBA 153, 804 (1968). 91. T . F. Chuang, Y . C. Awasthi, and F. L. Crane, J . Bioenerg. 5, 27 (1973). 92. T. F. Chuang and F. L. Crane, J. Bidenerg. 4, 563 (1973).

314

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

to its physical state. These studies have encompassed a wide range of techniques, and in this section it is intended to review the information available from them with particular emphasis upon the evaluation and interpretation of those data which may be of importance in unraveling the chemkal events that lead to the reduction of dioxygen to water. A. MODELS In the development of ideas about the fundamental mechanisms of dioxygen reduction one expects to be able to carry over into the protein system the basic physical and inorganic chemistry of copper and heme iron. Consequently, it is important to have a clear understanding of the response of copper complexes, heme A, and, perhaps, copper-heme A complexes to those chemical and physical probes that might be used with the oxidase. Although copper (I) and copper (11) complexes have been subjected to extensive investigation and their properties are quite well understood, few probes are effective in following copper in the oxidase and thus so little is known about the copper environment in the enzyme that the transposition of knowledge from simple systems to the enzyme system is difficult. The main probe is EPR spectroscopy where a portion, and only a portion, of the copper(I1) is visible. There is a probable requirement for copper to retain substantially the same coordination environment in the oxidized and reduced forms under rapid turnover conditions and for different environments for the two copper atoms. Fortunately, there are more opportunities to probe the iron and its associated porphyrin. Several physical properties can be examined and the structure-property relationships have been quite extensively worked out with some hemes, namely, heme B (protoheme) and other 2,4-disubstituted deuterohemes (60, 62, 93). However, the heme studies to date are not as relevant to the oxidase as future studies can be for two reasons. One is that heme A, although its structure has recently been elucidated and a few properties examined (60), has still not been as thoroughly studied as heme B. This is important, since heme A, in those few properties that have been studied differs significantly from heme B. A second factor is the paucity of data on electron exchange interactions between any heme system and another heme or a copper or another donor (or acceptor). Several p-oxobishemins, including p-oxobishemin A, have been thoroughly characterized (60, 94) , p-hydrazine-bishemes have been preRared 93. W. S. Caughey, C. H. Barlow, D. H. O’Keeffe, and M. C. O’Toole, Ann. N . Y . Acad. Sci. 206,296 (1973). 94. D. H. O’Keeffe, C. H. Barlow, G. A. Smythe, W. H. Fuchsman, T. H. Moss, H. R. Lilienthal, and W. S. Caughey, Bioinorg. Chem. (in press).

5.

CYTOCHROME C OXIDASE

315

(93),and copper-heme A interactions have been indicated (95); but, nevertheless, in sum, relatively little is known about interactions between hemes, coppers, and other donor-acceptor systems of the types likely to be present in cytoohrome c oxidase. Then, the application of the ideas of inorganic chemistry to the oxidase must be largely inferential but nevertheless can lead to useful generalizations about mechanistic pathways that might be accessible to dioxygen en route to water. Furthermore, model studies cannot be expected to hold all the answers t o the oxidase function because the protein is able to provide environments for electron transfer and an active site geometry that may not be readily duplicated in conventional chemical systems. Simple metal atom chemistry very rarely provides assemblies of metal ions in an ordered yet flexible environment where cooperative interactions can facilitate, in a single step, reactions that require a number of steps by the separate ions.

B. ELECTRONIC SPECTROSCOPY 1. Absorption Spectra

Electronic spectra provide a simple and convenient way to monitor changes induced in the oxidase by various chemical treatments. Indeed, spectral observations were a t the core of the pioneering observations of MacMunn ( l a ) , Keilin (96),and Warburg (97); and more recently many investigators have examined the spectra of isolated oxidase, mitochondrial particles, and electron transport particles. The spectra of the fully oxidized [oxidase ( I V ) ] (97a) and the fully reduced [oxidase ( 0 ) ] oxidase have been well characterized (52) (Table V). In Table VI are spectral parameters for ligand complexes of various oxidation states (98-105). Although the spectra of most of these complexes have been 95. R. A. Bayne, G. A. Smythe, and W. S. Caughey, in “Probes of Structure and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 613. Academic Press, New York, 1971. 96. D. Keilin, Proc. Roy. SOC.,Ser. B 98, 312 (1925). 97. 0. Warburg, Biochem. 2.152,479 (1924). 97a. Hereafter, the oxidation state (number of electrons removed) of cytochrome oxidase will be represented by a roman numeral, 0 to IV, in parentheses. 98. W. H. Vanneste, Biochemistry 5, 838 (1966). 99. C. Greenwood, M. T. Wilson, and M. Brunori, BJ 137,205 (1974). 100. R. Lemberg and J. Stanbury, BBA 143,37 (1967). 101. A. 0. Muijsers, K. J. H. Van Buuren, and B. F. Van Gelder, B B A 333, 430 (1974). 102. R. Wever, A. 0. Muijsers, B. F. Van Gelder, E. P. Bakker, and K. J. H. Van Buuren, BBA 325, 1 (1973). 103. Y. Orii and K. Okunuki, J. Bioehem. ( T o k y o ) 55,37 (1964).

316

W. S. CAUGHEY, W. J. WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

TABLE V ELECTRONIC ABSORPTION SPECTRAL DATAFOR CYTOCHROME c OXIDASEWITH A N D WITHOUT DETERGENT' Fully oxidized: oxidase(1V)b 830 1.2 1.1

Xmax,nm

emx with detergente e , ~without

detergentd

660 2.4 2.0

598 8.7 6.6

545 8.2 6.3

515 8.3 6.3

418 79 59

560 7.7 6.3

517 7.2 5.3

443 100 78

Fully reduced: oxidase(0)~ 603 19.3 14.5

Xmax,nm

with detergent" without detergentd

C ~ M

em^

Volpe and Caughey (68). As isolated. "Detergent, Tween 20. Oxidase(1V) dissolved in 0.1 M phosphate buffer 0.75% Tween 20, p H 7.4. d Oxidase(1V) dissolved in 0.01 M phosphate p H 7.4. Reduced with slight excess of sodium dithionite. a

observed many times (18) and may, on this basis, be considered well established, there remain some troublesome difficulties in their detailed interpretation (11). Assignment of frequency and intensity values to band maxima for heme a, heme a,, and copper components (98) has been accepted rather widely (18). However, the EPR evidence (66) for strong interaction among the metal components in terms of facile electron exchange and magnetic coupling indicates the likelihood that changes induced a t one component (e.g., oxidation or ligand binding) will affect the electronic spectra (and other properties) of the other components. Thus, it is risky indeed to ascribe individuality to the hemes or coppers on the basis of monotonic dependence of the spectra upon the states of the individual TABLE VI WAVELENGTHS OF ABSORPTIONMAXIMAFOR VISIBLEAND SORET OF CYTOCHROME c OXIDASE' SPECTRAOF COMPLEXES Oxidase-ligand complexb Oxidase(0) .CO Oxidase(II1). CO Oxidase(0) . O p Oxidase(II1) .02 Oxidase(1V) .FOxidase(1V). N8Oxidase(1V) .CN-

Ref.

Xrnnx.nrn

603

638 660

603 605 598 598 598

590(sh) 590

551 547 550 550 545 545 545

517

440(sh)

431. 429 428 428 421 421 425

98 99

100 99 101 108 10s

Preparative methods different but all contained detergent. The precise nature of ligand binding has not been established in these complexes.

5.

CYTOCHROME C OXIDASE

317

metal centers (11, 104, and conclusions drawn from experimental observations which depend upon this separation for quantitation and analysis should be viewed with caution. These comments apply both to attempts to synthesize spectra for cytochromes u and u3 on the assumption that the properties of one heme are independent of the oxidation state of, and ligands bound to, the other metals (98),and to the assignment of the 830-nm band to copper (105). Copper may be an active contributor to the 830-nm band since changes in EPR signals resulting from copper (11) have been noted to follow the intensity of the 830-nm band, but present evidence does not show that copper is the unique contributor, or even a contributor a t all, to the 830-nm band intensity (106-108). A comparison of the spectral differences between cytochrome c oxidase and heme A derivatives with those for hemoglobin or myoglobin and heme B derivatives reveals similar effects of protein environment on the heme moieties. The magnitude and direction of wavelength shifts upon going from heme species to proteins are comparable for the three proteins (Tables V, VI, and V I I ) . There is therefore little doubt that the 8-formyl group of heme A remains intact upon incorporation into the apoenzyme. The conversion from deoxy to carbon monoxy to oxy species results in similar spectral shifts (Table VII) (52, 65, 109-112). An exception is the blue shift in the 605-nm band upon reaction with CO, whereas hemoglobin (Hb) and myoglobin (Mb) experience a red shift upon binding CO. Nevertheless, the remaining spectral evidence supports similar terminal CO to Fe binding for the three proteins as do infrared C-0 stretch bands (113-116). Similar binding of 0, among the three proteins can also be assumed; infrared 0-0 stretch band data have shown this to be 104. W. S. Caughey, Annu. Rev. Biochem. 36, 611 (1967). 105. D. C. Wharton and A. Tzagoloff, JBC 239,2036 (1964). 106. W. S. Caughey and S. McCoy, in “Biochemistry of Copper” (J. Peisach, P. Aisen, and W. E. Blumberg, eds.), p. 271. Academic Press, New York, 1966. 107. L. N. Mackey, T. Kuwana, and C. R. Hartzell, FEBS (Fed. Eur. Biochem. Soc.) Lett. 36, 326 (1973).

108. L. E. AndrCasson, B. G. Malmstrom, C. Stromberg, and T. Vanngard, FEBS (Fed. Eur. Biochem. SOC.) Lett. 28,297 (1972). 109. R. Banerjee, Y. Alpert, A. F. Leterrier, and R. J. P. Williams, Biochemistry 8,2862 (1969). 110. Y. Sugita and Y. Yoneyama, JBC 246,389 (1971). 111. A. 0. Muijsers, R. H. Tiesjema, and B. F. Van Gelder, BBA 234, 481 (1971). 112. K. D. Hardrnan, E. H. Eylar, D. K. Ray, L. J. Banaszak, and F. R. N. Curd, JBC 241, 432 (1966). 113. J. 0. Alben and W. S. Caughey, Biochemistry 7, 175 (1968). 114. S. McCoy and W. S. Caughey, in “Probes of Structure and Function of

Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 295. Academic Press, New York, 1971. 115. W. S. Caughey, R . A. Bayne, and S. McCoy, JCS,D p. 950 (1970). 116. W. S. Caughey, Ann. N . Y . Acnd. Sci. 174, 148 (1970).

TABLE VII DATAFOR OXYAND CARBONYL COMPLEXES OF HEMOGLOBIN, VISIBLEA N D SORETSPECTRAL MYOGLOBIN, A N D CYTOCHROME c OXIDASE Complex 53(13.3) 569( 15.0) 576(15.2)

Oxidase(0)b Oxidase(0)COb Oxidase(0)OZc

603(14.5) 603( 14.0) 603(12)

Mba MbCO MbO,

556(12) 579(12.2) m(14.6)

a

539(14.9) 542(14.3) s590(11.5)

560(6.3) 551 (7.4) 550(10) 540( 14.0) 543(13.7)

Here, Hb stands for hemoglobin and Mb for myoglobin. determined in the absence of detergent. srn~ determined in the presence of 1.0 % cholate.

m Ref.

Xmax.nm(ernM-')

Hba HbCO HbOr

?

517(5.3) 517(6.3)

444 (50)

430(145) 420(208) 415(135)

109 109

?

109,110

4

443(77) 431(62) 428(84)

51 66 111

434( 114) 424( 187) 418(128)

111 111 111

F-1 ?

e,,,~

m

5.

CYTOCHROME C OXIDASE

319

true for HbO, (117)and MbOz (118) but not for oxidase(O).O, as yet. The marked difference in the spectra of the oxidase(1V) and p-OXObishemin A derivatives indicates a simple p-oxobishemin A structure is not present. However, interactions of copper with a p-oxobishemin moiety could result in the spectrum found for oxidase(1V) and explain also the generally low sensitivities of the visible spectra of oxidase to changes in ligation and oxidation state. 2. Circular Dichroic Spectra

The circular dichroic (CD) spectra of many cytochrome c oxidase derivatives have been observed (119-123) with reproducible results. Here also the insensitivity of the C D spectra to ligand substitution makes them difficult to interpret. Thus, the general shapes of the C D spectra for cytochromes a and a3 generated by the algebraic addition of spectra obtained from a number of derivatives were sufficiently independent of the specific ligands on the complexes used to generate the spectra that it was concluded that the hemes were acting independently (121, 123). However, detailed examination of the spectra revealed sufficient ligand-dependent differences in band positions and intensities to suggest cooperative interactions of the hemes (12U,122). Thus, directly opposed interpretations of the same data were presented. On the one hand, the traditional concept of identifiable cytochromes a and a3 is maintained, while on the other, the increasingly supportable notion of cooperativity between the hemes is suggested. Here also it has proved difficult to distinguish between the hemes (cytochromes a and a,) in oxidase(0) and oxidase(1V). But these hemes can be distinguished upon reaction with ligands or upon changing the oxidation state. Current evidence favors heme-heme interaction which results in changes induced a t one heme influencing the behavior and properties of the other. C. LIGAND BINDING STUDIES In addition to studies that have involved the determination of the thermodynamic states (potentiometric) and the observable valence states 117. C. H. Barlow, J. C. Maxwell, W. J. Wallace, and W. S. Caughey, BBRC 55, 91 (1973). 118. J. C. Maxwell, J. A. Volpe, C. H. Barlow, and W. S. Caughey, BBRC 58, 166 (1974). 119. D. W. Urry, W. W. Wainio, and D. Grebner, BBRC 27, 625 (1967). 120. D. W. Urry and B. F. Van Gelder, in “Structure and Function of Cytochromes” (K. Okunuki, M. D. Kamen, and I. Sukuru, eds.), p. 210. Univ. of Tokyo Press, Tokyo, 1968. 121. Y. P. Myer, JBC 248, 1241 (1971). 122. R. H. Tiesjema and B. F. Van Gelder, BBA 347,202 (1974). 123. F. C. Yong and T. E. King, BBRC 40, 1445 (1970).

320

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

(EPR)of the electron acceptors in cytochrome c oxidase, there have been other chemical studies directed toward understanding the changes that occur a t the catalytically active center as it goes from oxidized to reduced and back to oxidized again. These studies, as with the potentiometric and EPR studies, must be interpreted with some caution because they must of necessity be conducted under nonturnover conditions. I n these circumstances the steps that emerge from the analysis of the data may not correspond to any transitions that occur in the functioning enzyme. Nevertheless, such work has provided valuable insight into the interrelationships between the components of the active site. The binding of ligands and the consequent alteration of physical and chemical properties have been used to probe for the identification and differentiation of the cytochromes a and as (18).The observational techniques employed have varied from UV-visible and CD spectroscopy (discussed above) through potentiometry (124, 125) and EPR spectroscopy (126)( to be examined later), but there remains a measure of uncertainty in the interpretation of the results. The stoichiometry and stereochemistry of complexes have both been inferred from the changes in electronic spectra that accompany ligand binding and/or changes in the oxidation state (98, 127). However, while such spectral changes are convenient monitors of ligand interaction, they do not measure directly either the nature or the extent of ligand binding. Consequently, the complexes under study are frequently ill-defined despite the extensive examination that they may have undergone. 1. Azide, Fluoride, and Cyanide Fluoride (101, 128), azide (102, 128-150), and cyanide (103,128, 129, 151-156) have been observed spectrophotometrically to bind reversibly to the oxidase. The absorbance changes are small in all cases and the kinetics complex-at least two forms of complex are kinetically observ124. J. S. Leigh, Jr., D. F. Wilson, C. S. Owen, and T. E. King, ABB 160, 476 ( 1974).

125. Y. Fujihara, T. Kuwana, and C. R. Hartzell, BBRC 61, 538 (1974). 126. C. R. Hartzell and H. Beinert, BBA (in press). 127. T. Yonetani, JBC 235, 845 (1960). 128. S. Yoshikawa and Y. Orii, J . Bioch.em. ( T o k y o ) 71, 859 (1972). 129. S. Yoshikawa and Y. Orii, J . Biochem. ( T o k y o ) 71,873 (1972). 130. R. Wever, A. 0. Muijsers, and B. F. Van Gelder, BBA 325,8 (1973). 131. P. W. Camerino and T. E. King, JBC 241,970 (1960). 132. K. J. H. Van Buuren, P. F. Zuurendonk, B. F. Van Gelder, and A. 0. Muijsers, BBA 256, 243 (1972). 133. K. J. H. Van Buuren, P. Nicholls, and B. F. Van Gelder, BBA 256, 258 (1972). 134. S. Yoshikawa and Y. Orii, J . Biochem. ( T o k y o ) 73, 637 (1973). 135. S. Yoshikawa and Y. Orii, J . Biochem. ( T o k y o ) 76, 271 (1974).

5.

CYTOCHROME C OXIDASE

321

able in each case. Where the stoichiometry of the ligand binding reaction was tested (cyanide and azide), only a single (133) 1:l complex (102, 132) was formed. Fluoride was suggested, on the basis of its inhibitory behavior, to bind 2 moles/mole of oxidase although EPR spectra suggest little interaction with the heme iron (128). The site(s) of ligand binding has, then, not been clear but the suggestion has been made (136, 137) that azide inhibits a t a site common to both electron transport and energy conservation. In support of this contention, Wilson (137)has shown that 5-13 (5-chloro-3-t-butyl-2'-chloro-4'-nitrosalicylamide) releases azide inhibition of ATPase and electron transfer and that 1.35 molecules of s-13 per respiratory chain are needed to release inhibition of respiration. I n the presence of azide, the redox potential is altered, (138) and the EPR visible iron undergoes a high- to low-spin transition (139).Infrared observations of oxidase(1V) in the presence of azide revealed that even upon long standing the vNS- is not shifted from its free solution value ( V = 2047 em-', Av% = 28 cm-') (140). However, oxidase(I1) treated with azide followed by reoxidation t o oxidase (IV) exhibited a frequency (2038 cm-', h v , = 14 em-') consistent with iron-bound azide (140, 141). Clearly, azide was bound to the iron in the latter case but not in the former. It is important to discriminate among the possible ligand bonding configurations, but such differences in binding could not be established on the basis of electronic spectra alone. Infrared difference spectroscopy has enjoyed considerable success in the elucidation of the nature of binding of a variety of ligands to hemes and heme proteins (116, 1411, and since the method provides a direct measure of the character of the ligand bonds it promises to be a powerful tool in probing oxidase ligands, even within intact tissue (142), despite the large size of the enzyme.

2. Carbon Monoxide Carbon monoxide binds readily to iron (11), but not iron (111), porphyrins to form complexes that are quite distinctive in terms of the spectral properties both of the heme and of the bound CO. Thus, CO has been widely used as a probe of the active site of heme proteins (113, 136. D. F. Wilson and B. Chance, BBRC 23,751 (1966). 137. D. F. Wilson, Biochemistry 8, 2475 (1969). 138. D. F. Wilson, J. G. Lindsay, and E. S. Brocklehurst, BBA 256, 277 (1972). 139. B. F. Van Gelder and H. Beinert, BBA 189, 1 (1969). 140. W. S. Caughey, C. H. Barlow, J. C. Maxwell, J. A. Volpe, and W. J. Wallace, Ann. N . Y . Acad. Sci. 244, 1 (1975). 141. S. McCoy and W. S. Caughey, Biochemistry 9,2387 (1970). 142. J. C. Maxwell, C. H. Barlow, J. E. Spallholz, and W. S. Caughey, BBRC 61, 230 (1974).

322

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

114). In the past, such investigations with cytochrome c oxidase have been limited to visible UV spectroscopy ; however, more recently, infrared (66, 116), EPR (124), and potentiometric (138) techniques have been employed to investigate the reaction of CO. Carbon monoxide may bind a t iron(I1) (62, 115) or copper(1) (65,115) 0

II

either as a terminal (M-CO) or bridging (M-C-M) ligand. The presence of the C-0 stretch band a t 1963.5 cm-l for the CO complex with oxidase(0) provided firm evidence for binding a t iron(II), and not a t copper(I), in a terminal (end-on) fashion similar to HbACO (v-1951 cm-1) (115) and MbCO (v-1944 cm-1) (114). Furthermore, the extreme narrowness of the band showed a well-ordered (nonrandom) environment about the bound CO well isolated from external medium. A similar band was found for CO bound to the oxidase in intact heart muscle (142). These data are not consistent with suggestions that CO is bound cooperatively by both copper(1) and iron(I1) in CO (143). The stoichiometry of CO binding to the oxidase has been considered to be one CO per two hemes since Keilin's original interpretations of visible spectra (13). Results from many subsequent attempts (98, 144-147) to establish such a stoichiometry though variable, were interpreted as consistent with the Keilin suggestion. However, recent evidence from infrared intensities and exchange of CO from the oxidase to Hb gave clear evidence for one CO for each heme A (66). The very narrow bandwidth ( A V ~ , ~6, cm-I) shows the vc0 values for each heme are either identical or very nearly so since if more than one bond were present the frequencies would necessarily be nearly the same if such a narrow width were to result. Since vco is a very sensitive probe of cis, trans, and medium effects in heme proteins (49, 6 2 ) , the essentially identical vc0 for each heme A provides strong evidence that each heme A is of the same structure including trans ligand (histidine?) and that the environment about each bound CO ligand is the same. The CO ligands in HbACO are found at 1950 and 1952 cm-*, corresponding to the a! and p subunits, respectively, to give a v l I Z of 8 cm-' (148). It is of interest that in the presence of one iron(II1) heme, e.g., oxidase (11).CO, only one CO binds with 143. J. G. Lindsay and D. F. Wilson, FEBS (Fed. E w . Biochem. S o d Lett. 48, 45 (1974). 144. Q.H. Gibson and C. Greenwood, BJ 86,541 (1963). 145. M. Morrison and S. Horie, JBC 240, 1359 (1965). 146. Q. H. Gibson, G. Palmer, and D. C. Wharton, JBC 240, 915 (1965). 147. G. E. Mansley, J. T. Stanbury, and R. Lemberg, BBA 113, 33 (1966). 148. J. A. Volpe, J. C. Maxwell, W. J. Wallace, W. S. Caughey, and S. Charache, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 34, 687 (1975).

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vc0 a t 1965 cm-’ slightly, but only slightly shifted from the value for the fully reduced oxidase CO complex (149). Little is known about the relative affinities of the two heme A sites for CO. The infrared and H b exchange evidence noted above demonstrates that two CO ligands can bind, a t least in certain enzyme preparations. However, it is reasonable to expect the first CO to be bound with greater affinity than the second; therefore, in some preparations, only one CO may bind. I n mono-CO complexes, the heme to which CO binds can be called a3. But, there is no basis for knowing which of the two liemes in the fully reduced oxidase represents the preferred binding site or even whether there is a preferred binding site. Once CO becomes , other heme may in consequence adopt differbound to one heme ( a 3 ) the ent properties and become heme a. A reasonable interpretation of infrared (65) and other (99) data is one in which with either one or two electrons added to the fully oxidized enzyme, one CO binds a t iron. And, as discussed below, potentiometric and EPR evidence that CO binding affects the properties of other metal centers has been obtained (1.24, 137).

3. Dioxygen The infrared spectra show that CO binds to the oxidase (65, 114) in much the same way it binds to H b and Mb (113, 114). Hence, it.might be expected that the enzyme would form oxidase.0, in the same way that H b and M b form HbO, and MbO,. Formation of such an “oxygenated” complex represents a quite logical initial step in the sequence of reactions that lead to the reduction of oxygen to water by oxidase. Consequently, there have been a number of attempts to identify and characterize an “oxygenated” oxidase. In 1958, Okunuki and Sekuzu discovered a new form of cytochrome c oxidase, characterized by a Soret band a t 426-428 nm and designated i t “oxygenated” oxidase (160).The presence of the band a t 428 nm has been taken as a clear indication of the formation of the oxygenated complex (100) and is, in fact, the only evidence for the formation of such a complex. And, despite some uncertainty about its spectral characteristics (151), all discussion of the oxygenated complex has been cast in terms of the variation in both position and intensity of the band at 428 nm. I n the original concept of Okunuki and Sekuzu, dioxygen was thought 149. J. G. Lindsay, ABB 163, 705 (1974). 150. K. Okunrrki, B. Hagihara, I. Sekuzu, and T. Horio, in (‘Proceedings of the International Symposium on Enzyme Chemistry, Tokyo and Kyoto, 1957” (K. Ichihara, ed.), p. 264. Academic Press, New York, 1958. 151. H. Beinert, C. R. Hartaell, and W.. H. Orme-Johnson, in “Probes of Structure

and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S.Mildvan, eds.), Vol. 2, p. 575. Academic Press, New York, 1971.

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W. S. CAUGHEY, W. J . WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

to bind reversibly to oxidase(O), but the only direct evidence for this has been the observation (158) that upon evacuation the dioxygen appears t o be removed and replaced by carbon monoxide. Greenwood et al. (99) have prepared the “oxygenated” oxidase by photolyzing ferricyanide oxidized oxidase(0) .CO in the presence of 0,. The complex is formed readily and is fairly stable. Decay appears to occur through a slow intermolecular transfer reaction (153-155). Clearly, the mixed valence complex oxidase(II1) -0, is not readily formulated except on the familiar HbO, bent end-on model since only one reduced heme iron is available and there is no evidence to suggest that copper contributes to ligand binding. With two hemes present, two different mono dioxygen complexes are possible, but a t present there is no experimental basis upon which to decide whether the two complexes have the same or different properties or whether one is preferred over the other. However, the presumed observation of relatively stable mixed valence oxygenated oxidase complexes suggests that it may be possible to subject intermediate steps in the reduction of dioxygen to scrutiny by more discriminating techniques (such as infrared) and that this may provide the clues that will lead to a detailed understanding of the steps involved in the important enzymic reduction. The ability to observe oxidase(0) .O, a t low temperature and oxidase(II.1) or (11)*02 a t room temperature is quite understandable in chemical terms. Many of the other observations on “oxygenated” oxidase are rather more difficult to rationalize. Both Gilmour et al. (156) and Tiesjema et al. (157) observed that oxidase(0) reacts rapidly (<5 msec) with 0, to produce what is judged from the spectrum to be a mixture of oxidized [oxidase(IV)] and oxygenated oxidase. The proportions of the oxidized and oxygenated components in the reaction mixture depend on the concentration of dioxygen, more 0, gave more oxidase(O)-O,, and the latter complex was only slowly transformed into oxidase(1V) ( t l I z= 1 hr) . Orii and King (158) found three sequentially formed comto~ *IIIt~/~ahr to oxidase (IV) 3 under not plexes [ I h J S n r i n to I I 5 / ~ ~ 6 m 152. A. J. Davison and W. W. Wainio, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 23, 323 (1964). 153. Q. H. Gibson, C. Greenwood, D. C. Wharton, and G. Palmer, JBC 240, 888

(1966). 154. D. C. Wharton and Q. H. Gibson, JBC 243, 702 (1968). 155. B. Chance, C. Saronio, and J. S. Leigh, Jr., Proc. N a t . Acad. Sci:U. S. 72, 1635 (1975). 156. M. V. Gilmour, R. Lemberg, and B. Chance, BBA 172, 37 (1969). 157. R. H. Tiesjema, A. 0. Muijsers, and B. F. Van Gelder, BBA 256,32 (1972). 158. Y. Orii and T. E. King, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 21, 199 (1972).

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very different conditions. These observations are difficult to rationalize unless it makes a difference how oxygen binds to oxidase(0) in the initial step. Perhaps it is possible (indeed, even likely) that oxygen can interact with either of the two hemes [to form oxidase(0) -Or] or with both of the hemes [to form oxidase(0) . (O,),]. This would be analogous to the reaction of oxidase(0) with CO ( 6 5 ) . It is easy t o see the possibility that those molecules that bound one dioxygen could go on to oxidase(1V) but those that bound two dioxygens would be unable (not enough electrons) to complete the cycle and would remain trapped a t the “oxygenated” stage. The slow oxidation observed could then be accounted for by slow loss of one dioxygen and subsequent oxidation. If this speculation is correct, it would point out a requirement that only a single dioxygen binding site be available in the operating enzyme, i.e., one reduced and one oxidized heme iron.

D. POTENTIOMETRY The redox properties of cytochrome c oxidase have been investigated both by anaerobic reductive titrations (159) and by potentiometric titrations (160).Since measurements of the latter kind are, a t least in principle, able to provide absolute potential values, they have been favored in recent studies. The inconsistencies found in the early work (161-163) may have resulted from the lack of equilibrium conditions in some cases, from differences in the preparations, or simply from some incorrect interpretations of data. The importance of establishing that equilibrium conditions are attained has recently been recognized (107, lab, 125), but identical sets of measurements on the various types of preparations have yet to be reported. 1. Electron Economy

There now seems little doubt that complete oxidation or reduction of the isolated oxidase under anaerobic conditions requires four electron equivalents ( 164, 165). When the potentiometric titration is followed spectrophotometrically (at 605-630 nm) , high potential (at -350-375 mV) and low potential (210-230 mV) hemes (Table VIII) are indicated D. C. Wharton and M. A. Cusanovich, BBRC 37, 111 (1969). D. F. Wilson and P. L. Dutton, ABB 136, 583 (1970). K. Minnaert, BBA 110, 42 (1965). A. 0. Muijsers, R. H. Tiesjema, R. W. Henderson, and B. F. Van Gelder, BBA 267, 216 (1972). 163. T. Tsudzuki and D. F. Wilson, ABB 145, 149 (1971). 164. B. F. Van Gelder, BBA 118,36 (1966). 165. W. R. Heineman, T. Kuwana, and C. R. Hartzell, BBRC 49, 1 (1972). 159. 160. 161. 162.

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W. S. CAUGHEY, W. J. WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

TABLE VIII MIDPOINTPOTENTIALS FOR THE METALCENTERS OF CYTOCHROME c PROTEIN A N D I N MITOCHONDRIA OXIDASEI N ISOLATED Heme a

Copper

Investigators Muijsers et al." Tsudsuki and Wilsonb Leigh et al." Wilson et a1.d Mackey et a1.C Wilson et aZ.d*e Wilson et a1.d.J

200-250 335-360 225 375 210 360 220 380 215 f 10 340 k 10 2.50 105 345

225 215 k 10 350 f 10

~~

~

In the absence of cytochrome c the oxidase titrated as a single component with n = 1 and E = 280 mV. Partially purified cytochrome c oxidase; purified oxidase titrated aa a single component with n = 1 and E = 285 mV. Isolated cytochrome c oxidase. Mitochondria. In the presence of a 50 % CO, 50 % argon gas mixture. J In the presence of 20 mM aride. a

(125,162, 166). Wilson (137)has interpreted these results in terms of a, (high potential) and a (low potential) heme components. However, as pointed out so clearly by Malmstrom (11), a highly cooperative system, such as this one must surely be, does not lend itself to a unique interpretation of data obtained in these potentiometric spectrophotometric studies in terms of the assignment of low and high potential character specifically to a and a,. This does not mean that the hemes may not reside in different environments rather it means that the experimental observations do not require they do so.

2. Iron-Copper Coupling and Redox Potentials One of the objectives of these studies has been to gather evidence about the pathway of electrons through the oxidase to oxygen ; thus, there have been a number of attempts to differentiate between the iron and the copper in each of the redox steps. The earliest attempts a t the separation were based upon the assignment of the 60S-nm band to heme iron and the 830-nm band to copper. Although there is no doubt that the 830-nm 166. L. N. Mackey, T. Kuwana, and C . R. Hartzell, FEBS (Fed. Eur. Biochem. Soc.) Lett. 36, 326 (1973).

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band is sensitive to the oxidation state of the metals (53,167), the identification of this band as resulting from copper has been called into question (107). Nevertheless, Gibson and Greenwood ( 1 6 7 ) )on the basis of a kinetic study using the 830-band to monitor the copper oxidation state, placed the copper ( E m 280 mV) between the two hemes (a and a,) in redox potential. However, this value is the mean of the high and low potential heme components and may represent contributions from both components. In a potentiometric study, also based upon the 830-nm band, Tsudzuki and Wilson (163) reported an Em, value of 225 mV and an n value of 1.0 for copper, and when the decay of copper(I1) was followed by EPR as a function of potential, the corresponding value of E m was 250 mV (168). I n a more extensive study, Mackey et al. (107) generated oxidant and reductant coulometrically and followed the potential as a function of the number of electron equivalents added. From a computer analysis of the resultant potential-composition curves, they assigned midpoint potentials to each of the metal atoms in the enzyme as shown in Table VIII. These results confirmed the previous observations that reduction (or oxidation) proceeds through two iron-copper pairs, one at high potential and one at low potential. Although it now seems certain that the iron and copper components of the oxidase titrate as two pairs, one a t high potential and one a t low potential, caution must be applied in using these results to infer the relation between the heme iron and the copper in the fully oxidized (or fully reduced) states. I n such a highly cooperative system, the entry of a single electron may have a considerable perturbing effect upon the relative electron affinities of all the other electroactive centers in the molecule. N

3. Effect of Ligand Binding o n R e d o x Potentials

The potentiometric ligand binding studies of Wilson et al.(l38) give results that are consistent with the idea that in oxidase(I1) there is a reduced iron-copper pair and an oxidized iron-copper pair and that the hemes, at least, remain in interactive contact. In the presence of CO potentiometric titration of the oxidase (0) in pigeon heart mitochondria by ferricyanide indicates the presence of only a single electroactive component (Em, = 250 mV, n = 1.0). This result was interpreted to mean that CO has been bound to cytochrome LU? raising its Em to a value where it is not accessible to reaction with ferricyanide leaving only the low potential component cytochrome a as a reactive species in the titration. Although it would have been more satisfying to have had the experiment 167. Q. H. Gibson and C. Greenwood, JBC 240, 2694 (1965). 168. D. F. Wilson and J. S. Leigh, Jr., ABB 150, 154 (1972).

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W. S. CAUGHEY, W. J . WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

performed on carefully characterized CO complex (es) [e.g., by infrared spectroscopy (65)] , the observations are not out of line with what would be expected for a system in which only a single CO is bound. Even though it has been shown that two CO molecules will bind to oxidase(0) (65), each CO is expected to bind to oxidase(0) with different affinities, and there may well be circumstances under which one CO is bound per two hemes in fully reduced oxidase. Then the rise in potential for the unliganded heme A component would mean that the cytochromes are coupled in such a way that the unliganded cytochrome is sensitive to the changes in electron density produced a t the other cytochrome when it binds with CO. Alternatively, Malmstrom has suggested (11) that the data could be equally well interpreted in terms of the binding of CO to the low potential component so that it became the high potential component (by a Nernst law potential shift) and the cooperativity in the system then causing the original high potential component to become less readily reducible. However, even in these circumstances, the potentiometric measurements would leave some unresolved problems. If the two heme-copper couples are a t different potentials in the fully reduced oxidase, it would be very surprising if the strongly electrophilic ligand CO bound preferentially to the relatively electron deficient high potential component of the oxidase. However, if CO does not bind to the high potential component, it is necessary to suppose that in consequence of binding CO to the low potential component the high potential component has become less reducible (lower redox potential). A mechanism for this kind of charge transfer is difficult to imagine. There are then difficulties in complete rationalization of the experimental results when an initial difference in electron affinity for the two hemes is assumed. The potentiometric observations in the presence of azide are as expected, if azide binds only to the oxidized low potential component (137). Here, too, the ligand appears to bind to the wrong component. If there is an initial difference in reducibility between the two hemes, the high potential (more readily reducible) component would be expected to have the lowest electron density and hence represent the binding site of preference for the cr-bonding ligand azide. If azide does bind to the high potential site, the potentiometric observat,ions are difficult to rationalize. Thus here, too, the assumption of an initial or inherent difference in the electron affinities of the two heme sites leads to difficulty in the completely rational interpretation of the experimental observations. 4. Interpretation and Summary

On balance there seem to be fewer problems in interpretation if the two heme sites of oxidase(1V) or oxidase(0) are placed a t the same in-

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CYTOCHROME C OXIDASE

329

herent electron affinities. The subsequent introduction (or removal) of electrons or ligands will perturb the electron distribution among the electroactive centers so that they are no longer equivalent (in the redox sense). In a closely coupled set of electron acceptors entry of the first electron should be most thermodynamically favored and subsequent electrons, because of the increased electron density within the system, less so. This suggestion of initially equivalent hemes is supported by the infrared studies of the binding of CO ( 6 6 ) .The vc0 values are found to be very sensitive to differences in electron availability a t iron(I1) (93), and these values for the two hemes of oxidase(0) * (CO), must be nearly identical. Indeed, the visible spectra, which have been .widely used to follow the changes in valence state of the hemes as a function of the potential of the system, also fail to suggest any large differences in initial state electron affinities between the hemes. The large difference in electron density a t irons necessary to produce the observed potential differences would be expected to lead to large splitting of the porphyrin spectral bands (169).Attempts to resolve the visible band into components resulting from a and u3 have never been successful with the unperturbed enzyme system. It is, however, not surprising that spectral changes occur upon partial oxidation or reduction of the active centers since porphyrin spectra are sensitive to the electron densities at the metal centers (170). There seems then no reason to propose differences in the inherent reducibility of the hemes in oxidase(1V) on the basis of the evidence currently available. In fact, the intuitively attractive notion of equivalent and symmetrical heme-copper pairs seem adequate to explain the currently available potentiometric data.

E. ELECTRON PARAMAGNETIC RESONANCE STUDIES 1. Copper

The presence in cytochrome c oxidase of two copper atoms and two iron atoms of variable oxidation states would lead one t o believe that EPR studies would provide a rich fund of information about the oxidation state changes that occur in the functioning enzyme. Accordingly, many such investigations have been reported over the past decade. Much of the early attention was focused upon copper because its strong sharp signal in the vicinity of g = 2 was easily visible in both simple complexes 169. B. D. McLees and W. S. Caughey, Biochemistry 7,642 (1968). 170. W. S. Caughey, in “Inorganic Biochemistry” (G. L. Eichhorn, ed.), p. 797. Elsevier, Amsterdam, 1973.

330

W. S. CAUGHEY, W. J . WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

and proteins. Malmstrom and Vanngard (171) examined a number of both simple and complex copper-containing compounds and found characteristic spectra. Later Viinngard (172) tabulated spectra for many more compounds and pointed out that the copper(I1) could be classified in terms of All into Type I (All < 10 mK) and Type 2 (All > 14 mK). It is possible that these two classes correspond to tetrahedrally distorted coordination (Type I) and rhombohedrally distorted coordination (Type 11). However, too little is known about the coordination environment of copper in proteins to take such assignments seriously and the Type I and Type I1 classification should be considered only as a convenient empirical observation. Nevertheless, it is interesting that the inherent copper of oxidase(1V) (911 = 2.17, gm = 2.03, All 5 3 mK) (173) falls distinctly outside the range for Type I Cu2+. Thus, there appears to be a clear distinction between copper that is integrally bound in the oxidase and adventitious copper that appears as a variable contaminant in most preparations. This difference was not clearly understood in the early literature and led to some confusion about the role played by copper (174). The copper(I1) atom&) that give rise to the characteristic oxidase(1V) E P R signal are strongly bound within the enzyme and are not readily removed by treatment with complexing agents (53, l o g ) , but upon denaturation of the protein a more normal copper(I1) E P R signal is seen (175) and the copper then becomes susceptible to removal by complexing agents (53). Griffiths and Wharton (53) were able to show by a chemical method (176) that in oxidase(1V) both copper atoms were in the +2 oxidation state and this was later confirmed by the coulometric titrations of Heineman et al. (165).It was then surprising to find that the intensity of the E P R signal corresponded to only about 40% of the total copper known to be present as copper(I1) in oxidase(1V) (173) or, as now appears likely, 80% of one copper and none of the second copper. The low signal intensity together with the lack of fine structure in the E P R signal suggests that the copper(I1) interacts with other metal centers in such a way as to quench the inherent paramagnetism of divalent copper and produce the observed lowering of the E P R signal intensity. 171. B. Malmstrom and T. Vanngard, J M B 2, 118 (1960). 172. T. Vanngard, in “Biological Applications of Electron Spin Resonance” (H. M. Swartz, J. R. Bolton, and D. C. Borg, eds.), p. 411. Wiley (Interscience), New York, 1972. 173. H. Beinert, D. E. Griffiths, D. C. Wharton, and R. H. Sands, JBC 237, 2337 (1962). 174. A. Ehrenberg and T. Yonetani, Acta Chem. Scnnd. 15, 1071 (1961). 175. H. Beinert and G.Palmer, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), Vol. 2, p. 567. Wiley, New York, 1965 176. G. Felsenfeld, ABB 87, 247 (1960).

5.

331

CYTOCHROME C OXIDASE

2. Iron

The E P R signal resulting from iron(II1) because of its breadth and partial submersion under the copper(I1) signal has proved much more difficult to study quantitatively. Nevertheless, it now seems well established that in oxidase(1V) iron(II1) is represented by signals a t g = 3, 2, and 1.5 with intensities that correspond to about 40% of the heme iron present (126, 139). Although the oxidation state of the iron in oxidase (IV) has never been directly determined, the electron capacity of the fully oxidized enzyme suggests that all the iron must be iron(II1). Then, the low E P R signal intensity is most conveniently interpreted in terms of metal-metal interaction between iron(II1) (low spin) and some other paramagnetic center which partially quenches the signal expected from the S = .2 net spin on the iron. Both heme-heme and heme-copper (11) interactions have been suggested (138). Further elucidation of this problem might be forthcoming on the basis of careful magnetic susceptibility and Mossbauer studies. Unfortunately, the single published accounts of magnetic susceptibility (174) and Mossbauer (17'7) studies are difficult t o interpret. Nevertheless, it is interesting to note on the most naive basis that the magnetic susceptibility results of Ehrenberg and Yonetani (17'4) correspond t o -80% of two unpaired electrons. This is just as expected on the basis of the E P R observations on intrinsic copper and iron. 3. The Effect of Valence State Changes on the EPR Spectrum

Behavior of the signals resulting from E P R visible iron and copper during reduction provides clues that may be important to the understanding of the chemistry involved in oxygen reduction. Experiments carried out on a relatively long time scale show that the first electrons to enter the system give rise to a diminution of the low-spin iron(II1) ( g 3) signal and a corresponding increase in the broad g 6 signal attributed to high-spin iron(II1) (139). The g 6 signal reaches maximum intensity (corresponding to about 40% of the heme A) when two electrons per mole of oxidase have been supplied to the system and then declines leaving a very small rhombohedra1 signal when four electrons have been supplied. The copper (11) signal remains unchanged (perhaps increasing slightly) during entry of the first pair of electrons and then diminishes t o zero during entry of the second pair (139, 178). Leigh et al. .(124) have shown that the high-spin iron (111) signal makes its appearance with a half-reduction potential of 380 mV and disappears with a half-reduc-

-

-

177. G. Lang, S. L. Lippard, and S. RosBn, BBA 336,6 (1974). 178. C. R. Hartzell and H. Beinert, BBA (in press).

-

332

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S . YOSHIKAWA

tion potential of 210 mV. The copper(I1) signal disappears with a halfreduction potential of 250 mV. Similar observations have been made for both isolated oxidase (124, 125) and pigeon heart mitochondria and phosphorylating submitochondrial particles (168). It is apparent then, that the E P R results are in substantial agreement with the potentiometric results in showing that the iron and copper are reduced as two iron-copper pairs. At low temperature (<15OK), the g 6 signal is resolvable into a number of components. The dominant species are a broad rhombic signal and a narrow axial signal but other small signals are seen as well (126), depending upon the nature of the reductant used. I n ETP and mitochondria the splitting of the rhombic component is quite marked (124, 168) but, as purification proceeds, the outer wings (g = 6.8 and 5.3) broaden and the axial component becomes dominant. The two paramagnetic components reduce with slightly different half-reduction potentials and on this basis may be different species (168). However, Hartzell and Beinert (126) pointed out that it is not clear, on the basis of the available evidence, whether the two components of the g 6 signal are inherently associated with the oxidase so that the distortion of the signal is associated with a conformational change or heme-heme interaction induced by partial oxidation or reduction or whether it is simply another heme protein. Wilson and Leigh (168) point out that in purified cytochrome oxidase only the symmetric component had been seen (159) but that the two component signal appears in ETP and mitochondria1 particles and by implication suggest that this may be due to alteration of the enzyme during purification. However, it was shown by Wever et al. (179)and by Leigh and Wilson (180) that this highly asymmetric signal appears in small amount upon photodissociation of the CO complex of oxidase (11) and may be a property of the oxidase although alteration of the protein cannot be completely discarded. The origin of the two species represented by this signal is far from clear. But, if the system is as highly coupled as many of the experimental observations seem to suggest, it may be that the unpaired electron spin can go visiting on both hemes and that, in fact, both hemes are partially magnetically visible. H

H

4. The Effect of Ligand Binding

-

It is clear that the high-spin iron(II1) ( y 6) appears as the !ow-spin iron(II1) (g 3) disappears. It is, however, not so clear whether the two signals originate from the same or different hemes or, as suggested H

179. R. Wever, J. H. Van Drooge, G . Van Ark, and B. F. Van Gelder, BBA 347, 215 (1974). 180. J. S. Leigh, Jr. and D. F. Wilson, BBRC 48, 1266 (1972).

5.

333

CYTOCHROME C OXIDASE Fe"-O-Fem

A0

FeEOH,

HO-Fe=

N;

FeKOH,

&-Fern

FIG.4. Reductive cleavage of a poxobisiron(II1) complex with subsequent binding of aride to the unreduced iron atom. above, a combination of contributions from both hemes. The results do suggest though that it is an interaction between the two hemes in oxidase (IV) that induces the low-spin electronic configuration in the heme system. Upon partial reduction the stabilizing interaction is lost and the iron (11)-iron (111) pair exhibits high-spin (weak ligand) characteristics for the iron(II1) component. The presence of a bonding interaction between the hemes in oxidase(1V) was also indicated by ligand binding studies. Although some aspects of the binding of ligands like fluoride, azide, and cyanide to oxidized oxidase remain unclear, there seems no doubt that these ligands either do not react or react only very slowly with iron under fully oxidized conditions (131). In most hemins and oxidized hemeproteins, the reaction between iron (111) and anionic ligands occurs a t a virtually diffusion-controlled rate (181). The slow reaction with the iron (111)atoms of oxidase(1V) is then surprising. However upon half-reduction of oxidase (IV) (i.e., addition of two electrons) the reaction between the high-spin iron(II1) ahd anionic ligands such as azide occurs at the expected rapid rate (139) to produce a stable low-spin (g = 2.9, 2.2, and 1.67) (138, 168) iron(II1) azide complex. The binding of azide to iron in these circumstances is also demonstrated by infrared studies (182). The interpretation of these observations that is chemically most appealing would implicate a bridging ligand as the protective interaction between the two oxidized heme A components. Such a bridging ligand would need to have a number of rather sharply defined properties. Effectively it would need to be (a) divalent (for charge saturation), (b) less strongly bound to iron(II1) in the uncoupled than in the coupled form (to allow for displacement by azide), and (c) pH sensitive [Wilson observed pH dependence on the high potential component (137) 1. Although there are many potential ligands to iron available in the protein, it is difficult to find a protein constituent that fulfills all the requirements. The one ligand that is available in aqueous solutions and which otherwise meets all the requirements is oxide. If oxide is the bridging, protecting ligand the uncoupling reaction can be represented as in Fig. 4. 5. p-Oxobishemin A as a Possible Component of Oxidase ( I V )

The p-0x0 complexes are the commonly observed products of the oxygen-induced autoxidation of simple hemes, and their properties have been 181. Q. H. Gibson, L. F. Parkhurst, and G. Geraci, JBC 244, 4668 (1969). 182. J. A. Volpe and W. S. Caughey, BBRC (in press).

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W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

well described (60,94). Such Fe”’O-Fe”’ complexes are known, to be strongly antiferromagnetically coupled (183) and not to exhibit an EPR spectrum (94). Thus, the EPR results on oxidase(1V) are not consistent with only a p-oxobishemin A system. However, EPR results for iron could perhaps be accommodated by the introduction of a closely coupled third paramagnetic center with spin one-half. This could be the “EPR silent” copper. The close coupling of the spins of three spin one-half centers (two hemes and one copper) would give rise to resultant spin states of S = Q and 4 of which the S = 4 is expected to be the lowest lying (184). Such a coupled metal triplet could give rise to the set of ESR signals attributed to low-spin iron(II1). The visible copper would then be either independent of the coupled triad or only weakly coupled to it. (Note that both visible copper and visible “iron” represent only ~ 4 0 % of the signal expected if all were seen, but that if one copper were totally silent then 80%, or nearly all, of the other copper is seen.) Scission of a bridging p-0x0 linkage by partial reduction will lead to uncoupling of one iron atom from the triad with replacement of the strong field oxide ligand by a weak field hydroxo ligand as shown in Fig. 4.

F. INTERACTION OF CYTOCHROME c OXIDASEWITH CYTOCHROME c Cytochrome c is the sole physiological source of electrons for dioxygen reduction a t cytochrome c oxidase (11). It is crucial to a discussion of the mechanism of dioxygen reduction to know whether the binding of cytochrome c to the oxidase is to a single site or to more than one site. This will determine whether the reduction reaction can occur symmetrically (e.g., two sites) or asymmetrically (one site). On the basis of kinetic studies (186, 186) a 1:1 complex between cytochrome c and the oxidase was suggested. This seems to correspond to the ratio usually found in mitochondria (187). On the other hand, preparations have contained an isolated complex with one cytochrome c per heme A (188, 189). There is no obvious basis upon which to reconcile 183. T. H. Moss, H. R. Lilienthal, C. Moleski, G. A. Smythe, M. C. McDaniel, and W. S. Caughey, Chem. Commun. p. 263 (1972). 184. J. E Huheey, “Inorganic Chemistry : Principles of Structure and Reactivity,” p. 35. Harper, New York, 1972. 185. T. Yonetani and G. S. Ray, JBC 240, 3392 (1965). 186. E. Mochan and P. Nicholls, BBA 267, 309 (1972). 187. J. N. Williams, Jr., BBA 162, 175 (1968). 188. Y. Orii, I. Sekuzu, and K. Okunuki, J . Biochem. ( T o k y o ) 51, 204 (1962). 189. T. E. King, M. Kuboyama, and S. Takemori, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 2, p. 707. Wiley, New York. 1965.

5. CYTOCHROME

335

C OXIDASE

these differences. Thus, there is no unequivocal stoichiometric basis for ascribing either symmetry or asymmetry to the cytochrome c-oxidase complex. However, the absence of evidence for a second binding site on the oxidase ( 2 1 ) , the mitochondria1 ratio of cytochrome c to the oxidase (1.0),and the EPR evidence for asymmetry in the reduction of the metal components (126,138) are all consistent with a 1 : 1 complex. This suggests that the two copper atoms are different and that it is always the same copper that lies on the initial electron acceptor pathway.

G. KINETICSTUDIES The study of transformations that can be observed kinetically in the system cytochrome c: oxidase :0, has made important contributions to the understanding of the oxidase function, and much of this work has been reviewed previously (18, 21). Cytochrome c oxidase is reducible at different rates by a variety of agents including dithionite (157, 190), NADH ( 1 5 7 ) , ascorbate (191), and dichlorohydroquinone (153). However, the one that is of most interest is the natural substrate cytochrome c which appears to be the sole reducing agent under physiological conditions. Under both aerobic and anaerobic conditions the initial reaction between reduced cytochrome c and oxidase (IV) exhibits first-order dependence on cytochrome c (108, 185). However cytochrome c’+ acts as a competitive inhibitor for the reaction, blocks the electron transport binding site on the oxidase, and causes the kinetics to become complex as i t accumulates in reaction mixtures containing the isolated oxidase (185). I n the absence of oxygen cytochrome c2+rapidly (t,,. = 10 msec) reduced oxidase (IV) to oxidase (11), but subsequent changes are slow even in the presence of excess cytochrome c2+ and do not involve additional electrons (66, 107, 153, 192-194). The initial rapid reduction which was followed spectrophotometrically in the visible region by the disappearance of cytochrome c?+ or in the EPR by the disappearance of the g 2 (copper) and g 3 (low-spin iron) signals (66, 194) was found to be independent of the presence of cyanide in the reaction mixture (192). These observations have been interpreted in terms of the initial reduction of oxi-

-

IV

190. R.Lemberg and M. V. Gilmour, BBA 143,500 (1967). 191. T.Ozawa, Y. Takahashi, A. N. Malviya, and K. Yagi, BBRC 61, 601 (1974). 192. E. Antonini, M. Brunori, C. Greenwood, and M. T. Wilson, Biochem. Sac. Trans. 1, 34 (1973). 193. K.J. H.Van Buuren, B. F. Van Gelder, J. Wilting, and R. Braams, BBA 333, 421 (1974). 194. H. Beinert, R.E. Hansen, and C. R. Hartrell, BBA (in press).

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W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

dase (IV) being accommodated by the magnetically visible iron and copper and the slow step corresponding to an electronic rearrangement in which the electrons are passed to the other iron-copper pair. This interpretation receives its strongest support from the rapid flow EPR studies of Beinert and co-workers (66, 194) in which the copper ( g 2) and lowspin iron ( g 3) signals disappear upon introduction of the first electrons into oxidase(1V). Upon standing many minutes the copper signal reappeared qualitatively and quantitatively unchanged while the g 3 signal was replaced by a g 6 signal [high-spin iron (111)1. This would suggest that the iron and copper of the magnetically visible pair are the primary electron acceptors in oxidase (IV) , that a facile electron transport pathway connects the iron and the copper, and that the initial electron receptor pair is isolated from the other pair (153,192). The oxidase(I1) formed in this initial reaction should then be able to bind CO or 0, a t the reduced iron site. Recent work on the partially reduced oxidase indicates this to be the case (99,155). In fact, under these conditions, as expected, the oxygen adduct appears quite stable and long-lived (99),and the unreduced iron is able to bind cyanide (134). For the reduction of oxidase(1V) by cytochrome cz+ in the presence of oxygen, however, the situation changes dramatically. The initial, rapid two-electron reduction is followed by a slower, but still rapid, further reduction (108, 155, 192), which is sensitive to the concentrations of both oxygen and cyanide in the reaction mixture (108). I n the presence of excess cytochrome c*+ and high concentrations of oxygen the enzyme will turn over rapidly (19.2). The oxidation by oxygen of oxidase(0) formed by photolydng oxidase(O)-CO was found by Greenwood and Gibson (195) to be rapid (7cd 8 X lo7 M-’ sec-l) , with the kinetics controlled by the rate of diffusion of oxygen to the reaction site, but no evidence was adduced for an “oxygenated” intermediate with a lifetime longer than a few microseconds. The overall rate of oxygen reduction has since been confirmed by a number of investigators (166,171, 185),but the nonformation of an oxygenated adduct as the first step in reduction seems no longer tenable. Chance et al. (155)on the basis of the spectrophotometric titration of the oxidase (formed by photolysis of oxidase(O).CO a t low temperature) with oxygen, suggested that the reaction [reaction (15) ] is occurring with K = 320 pM. The “oxygenated” oxidase is formed a t a rate of

-

-

-

-

-

+ 02 % oxidase(O).O*

(15) -3 X lo4 M-’ sec-1 at - 7 8 O (E, = 9.9 kcal/mole) and decays by a firstorder step to a second complex at a rate of 0.45 sec-‘ (8, = 12.5 kcal/ Oxidase(0)

195. C. Greenwood and Q. H. Gibson, JBC 242,1782 (1967).

5.

CYTOCHROME C OXIDASE

337

mole). I n a parallel experiment oxidase(0) *CO was oxidized by ferricyanide [to oxidase(I1) .CO] and photolyzed in the presence of 0, a t low temperature. The spectral changes observed were interpreted in terms of the initial, rapid, reversible formation of oxidase (11)* 0, which then slowly decayed to an unidentified product spectrally distinct from the decay product of oxidase(0) SO,. Although not firmly established, a reasonable first product of the reaction between oxidase(0) and 0, is one in which dioxygen interacts with a single iron atom in the bent end-on fashion characteristic of isolated heme sites as in HbO, (117)and MbO, (118). This is also supported by the similarity in the frequencies of CO bound to oxidase(0) (65,115) and to H b (113). Despite the fact that very few of the kinetic studies have been carried out under anything approaching turnover conditions for the oxidase, these studies have provided important insights into the mechanism of its function. It seems quite clear that electrons enter the system through the cytochrome c binding site with very low activation energy to reside on the nearly independent iron and copper atoms referred to as EPR visible. Oxygen is able to promote both the entry of a pair of additional electrons into the system and the reoxidation of visible copper and iron, presumably through the reduction of oxygen to water. However, the fully reduced oxidase reacted faster with oxygen than did the half-reduced indicating that, although the electron redistribution problem is improved by the interaction of oxygen with the half-reduced oxidase, the most favorable configuration is obtained only upon full reduction. Electrons enter the oxidase singly but must react with oxygen in pairs. This must be related to the relative stabilities of the oxidation states of the reduced oxygen rather than any problems associated with the binding of oxygen to l-electronreduced states.

IV. Mechanisms

The organization of cytochrome c oxidase to carry out the reduction of dioxygen to water must reflect the mechanism by which electrons are supplied, the thermodynamic requirements of the oxidation states accessible to oxygen, and the chemical transformations of oxygen en route to water. From the above discussions it can be seen that widely diverse evidence relevant to each of these areas has been accumulating, and this evidence is now sufficient to suggest new mechanistic insight into the 0, reduction process. Although any such mechanistic schemes must necessarily be speculative, they are worthwhile not only because of their inher-

338

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

ent interest but also because they can stimulate and provide focus for further experimentation to test their validity. The kinetic and EPR evidence for the presence of two, a t most, weakly interacting, copper-heme A pairs in cytochrome c oxidase is quite convincing. Each pair appears to exchange electrons readily between the respective copper and iron components but in one pair the magnetic coupling between copper and iron, a t least in the fully oxidized species, is greater than in the other pair. Diagrammatically this is represented in Fig. 5. The Cu-Fe pair on the left is represented as having weak magnetic coupling and the pair on the right as having strong magnetic coupling. Little is known about the nature of the magnetic coupling in the fully, or partially, reduced states except that the net effect is to reduce the magnetism below that expected on the basis of the number of paramagnetic centers present. The possibility for magnetic coupling between the irons of oxidase(1V) was discussed above, and the idea of one strongly and one weakly coupled heme copper pair seems entirely consistent with the available data. Restriction of the ability to exchange electrons between the two pairs is represented by the dashed double arrow between the pairs. The evidence for O2 binding a t iron, and not a t copper, is strongly supported by the infrared evidence for CO binding a t iron only (14, 6 5 ) . Furthermore, since each heme A iron can bind CO (65) and heme A derivatives can form FeOFe bridges without undue steric restrictions (60, 6 2 ) , it is reasonable to suggest the possibility, in analogy to model heme autoxidations (27, 40,9 4 ) , that a single oxygen molecule bridges between the two heme irons. With nonphysiological reducing agents small enough to gain access to the active site, e.g., dithionite, it is reasonable to expect reduction to occur through direct interaction between one or both irons and the reducing agent. On the other hand, electron entry from cytochrome c appears to be a t copper in a t least one Cu-Fe pair. This could occur symmetrically (two cytochrome c binding sites) as in Fig. 6, or asymmetrically (one cytochrome c binding site) as in Fig. 7. I n the asymmetric process

FIG.5. Schematic representation of the interrelationships between the metal centers at the active site of cytochrome c oxidase showing an electronically coupled iron-copper pair (left) and an electronically and magnetically coupled iron-copper pair (right) which interact weakly with each other (---I.

5.

CYTOCHROME C OXIDASE

[V]

339

- _ _ _ CytcZ+

Cytc2*----.

,

Fa,- = - = -.Fa HlzO

0 2

FIG.6. Schematic representation of the symmetric reduction of cytochrome c oxidase by cytochrome c . I n this case cytochrome c is shown introducing electrons into both iron-copper pairs with dioxygen extracting electrons a t iron and being converted to water.

(Fig. 7) the copper that receives electrons from cytochrome c is designated the proximal copper (Cup) and the iron with which it is paired is the proximal iron (Fe,) . The metals of the other pair are termed the distal copper (CU,) and distal iron (Fe,). There is, of course, no need (or basis) to make a similar distinction between the coppers in the symmetrical process. A symmetrical reduction process would involve separate electron entry into each Cu-Fe pair from two cytochrome c binding sites. The symmetric process is represented in Fig. 6. If this process is in fact followed, it would be advantageous for both heme irons to go reduced virtually simultaneously and this introduces two possible mechanistic problems. Reduction would be slowed by the requirement of two cytochrome c molecules binding t o one oxidase, and with both iron atoms reduced there is the possibility of one dioxygen molecule binding a t each heme site. Both of these circumstances would tend to retard the reaction. If, despite these obstacles, the reaction does proceed with entry of electrons to both Cu-Fe pairs, a reasonable mechanism for 0, reduction becomes that of 0, binding t o one iron(I1) as to oxyHb or oxyMb (117,118),then, to a p-peroxo

0,

H120

Fro. 7. Schematic representation of the asymmetric reduction of cytochrome c oxidase by cytochrome c. In this case cytochrome c is shown introducing electrons into only one of the iron-copper couples (proximal) with reduction of dioxygen to water induced by electrons that originate on only one of the iron atoms.

340

W. 5. CAUGHEY, W. J . WALLACE, J . A. VOLPE, AND 5. YOSHIKAWA

Fe'

ze

z n+

O*

F%

FeLOw0

FeEOH HO-Fr"

Fe"

-

F&O-FE*

FIG. 8. Representative steps in the symmetrical reduction of dioxygen to water.

intermediate, and, finally, to 2 FeIII-OH or Felll-O-Felll as represented in Fig. 8. In this case, the coppers would both serve as components in the electron pathway to 0,. The cleavage of the 0-0 bond may proceed via a homolytic process to give two intermediate Fe1I1-0' species followed by one-electron reduction of each. The driving force for this cleavage would have to lie in the pressure exerted by the cytochrome c electron transfer system since a similar cleavage has not been demonstrated as a component in the nonenaymic decomposition of peroxides in protic solvents. However, present evidence for cytochrome c reductions suggest there is only one site on the oxidase; therefore, the reduction has been termed asymmetric" and can be represented schematically as in Fig. 7. The evidence that supports an asymmetric process to a greater degree than the symmetric process includes that for only one cytochrome c binding site and for one Cu-Fe pair reducing much more readily than the other pair. Asymmetric electron entry imposes interesting requirements on the 0 2 reduction processes. As shown in Fig. 9, 0 2 binding would occur as in the symmetric case. With only one electron added to oxidase(IV), i.e., to give oxidase(III), the 0 2 could bind to the Fe:! in the bent end-on fashion characteristic of oxphemoglobin (116). This bonding results in charge transfer from iron to dioxygen through interaction of the dr-pn* systems of the metal and dioxygen, respectively. Nevertheless, on the basis of reactivity studies of HbOz with nucleophiles (IN), the iron and the oxygen are expected to retain appreciable Fe"-02 character and ion pair. This is repreaented not to attain the status of an Fe1"-02diagrammatically in A of Fig. 9. In these circumstances, it is expected to be difficult (i.e., high activation energy) to bring up a second electron ((

FIG.9. Dioxygen bridging between proximal and distal iron atoms as the initial step in reduction. 196. W. J. Wallace, J. C. Maxwell and W. S. Caughey, BBRC 57, 1104 (1974).

5.

341

CYTOCHROME C OXIDASE

through an already reduced iron because this would involve (in formal valence terms a t least) transient formation of Fe1-02 or FeIT-02-. Of course, the presence of reduced cytochrome c and Cug coupled to the Feb1-02 would represent a forcing condition that could make the transfer of electrons through iron to O2 more favorable. However, even this extra "electron pressure" may not be enough to cause the reaction to proceed without the system having some way t o redistribute the electron density on Feb' onto another center (other than 02).It is proposed here that a suitably positioned distal iron(II1) atom might serve this purpose by interacting with the partially negative terminal oxygen to form a bridge between the irons across which an electron could pass to form a transient Fe~l'-O-O-Fe~ intermediate (B of Fig. 9), which with oxidation of Cug and Fe;' yields a p-peroxo complex with all metals in their oxidized state (C of Fig. 9). It can be seen here th a t without the driving force of the closely coupled electron source (Cui) there would be little tendency for the 0 2 to bridge between iron(I1) and iron(II1) atoms. This process permits ready formation of the thermodynamically stable peroxide intermediate. It may also be noted that attack by the terminal oxygen atom is likely to proceed by displacement of a ligand, presumably hydroxyl or water, from Fe;'. The next reasonable step involves cleavage of the 0-0 bond. This is envisioned as proceeding in the manner shown in Fig. 10. Here cleavage of the 0-0 bond of the p-peroxobisiron(II1) complex A must occur without the entry of electrons into the distal Cu-Fe pair (no low energy pathway across the p-peroxo bridge) , and consequently must be achieved by an heterolytic process. This could occur uninfluenced by electron pressure from cytochrome c (A + B -+ C) or under the influence of electron pressure (A + D + E + C). T o go from A to B can be thought of

cuf

1.

FeFOH

%"*

D

HO-Fet

C

E

FIG.10. The alternate routes to asymmetric cleavage of the p-peroxobisiron(II1) complex formed in the first stage of dioxygen reduction (see Fig. 9).

342

W. S. CAUGHEY, W. J. WALLACE, J. A. VOLPE, AND S. YOSHIKAWA

cut 0

-

.CUD

QI0

QO-FZ 0-Fe, FIG.11. Delocalization of the negative charge from oxygen over both iron and copper atoms in close magnetic and electronic contact.

as a splitting t o give one positive oxygen (two-electron deficiency, Fe;" 0+)and one negative oxygen (electron saturated, Fe': 0-). The driving force for this cleavage is expected to be the stabilization of the hydroxyl anion bound to iron(II1) promoted by the presence of distal copper(I1) as illustrated in Fig. 11. The hydroxyl anion will be stabilized by charge delocalization from oxygen over iron and copper atoms in a way that is rather like that for the carbanion resulting from removal of a proton from a carbon adjacent to a carbonyl group (Fig. 12). The key to both the stereochemistry and energy of the heterolytic cleavage of dioxygen is the polarization of the 0-0 bond induced by the bridging interaction between the two iron centers. Thus, with a n additional electron in the proximal copper-iron pair, the 0-0 bond is more polarized in D than in A. I n t,his way both the energy and polarity of the 0-0 bond cleavage can be modified to allow the reaction to follow a low energy pathway of unique polarity. Then E will form from D more readily than B from A. Coupling D to cytochrome c2+ will provide still stronger electron pressure for facile cleavage of the 0-0 bond. The mechanism can then be considered a sort of push-pull arrangement. The build up (from cytochrome c2+) of electron density on the proximal Cu-Fe pair results in an ability to place greater electron density on the proximal oxygen atom to cause repulsion of the distal oxygen atom as oxide thereby rupturing the 0-0 bond. The delocalization (i-e., stabilization) of the negative charge on the oxide formed from the distal oxygen atom serves to "pull" away the distal oxygen. The product C of Fig. 10 could represent the normal form of the fully oxidized oxidase, oxidase (IV). However, experience with hemins suggests that the p-oxobishemin would form readily-unless the protein provides special conditions that make its formation unfavorable. Clearly, if an R

0

II H

-C-F;-

I

I V

0 -+-

FIG.12. Schematic representation of stabilization of a carbanion adjacent to carbonyl group by electron delocalization.

IL

5.

343

CYTOCHROME C OXIDASE

FeOFe linkage forms, interactions with coppers (and with protein) are required in explanation of the E P R and electronic spectral differences between oxidase(1V) and p-oxobishemin A derivatives (60). Of course, it is possible that resting oxidase(1V) could have an FeOFe linkage which might never form under turnover conditions. It is also of interest that the asymmetric mechanism presented does not require that the distal Cu”-Fe“’ pair ever become formally reduced during enzyme function. The mechanism also provides for the receipt of electrons readily one a t a time from cytochrome c and delivers them to bound 0, under thermodynamically acceptable conditions. The nature of coupling betweeen copper and hemes in the proximal and distal pairs has been the subject of interesting speculations (60, 69). Although, as noted above, little firm evidence of the environments about the coppers have been reported, possible ways in which copper can interact with heme A have been presented (60). One intriguing possibility has copper interacting with the terminal double bond of the farnesylethyl side chain (Fig. 2) which can assume a conformation (Fig. 3) with n-electron overlap between its three double bonds and porphyrin. If cytochrome c?+ can pass an electron to the copper, then, in this way, there could result a fast, low activation energy pathway for the electron from copper to heme iron and 0, (Fig. 13). This coupling could reasonably provide for facile electron transfer without strong magnetic coupling, a necessary criterion for the proximal Cu-Fe pair. Involvement of the farnesylethyl group in this way also permits conformational control of the electron transfer in that small changes in conformation could turn electron transfer on or off by altering the effectiveness of w-orbital overlap. Such conformational changes could result from interactions with ATP and/or ADP and thus provide a mechanism for respiratory control processes. Other sites for copper interaction with heme A suggested include binding of copper a t the 2-1’ hydroxyl and to porphyrin ring and side chain unsaturation (60). These might serve in the tight coupling found for the distal Cu-Fe pair. I n any case, the differences in copper-heme A interac-

f.-[

-cu-l&Jqe~o Cyt c

0

FIG. 13. Schematic representation of a possible mechanism for electron transfer from cytochrome c to the active site of cytochrome c oxidase. The pathway is represented as involving copper as a mediator between the heme edge of cytochrome c and the “stacked” 2-alkyl chain of heme A followed by transmittal of the electron through the T system of heme A to dioxygen bound a t iron.

344

W. 6. CAUGHEY, W. J . WALLACE, J . A. VOLPE, AND S. YOSHIKAWA

tions between the two hemes is not such as to have provided evidence thus far that the two hemes are detectably different in fully reduced or fully oxidized forms. Copper could also serve in electron transfer between the two hemes, but present evidence appears compatible with bridging oxygen serving this role, if needed, in the functioning enzyme.

ACKNOWLEDGMENT Dr. Helmut Beinert and Dr.Britton Chance kindly provided information in advance of publication. Support from U.S.P.H.S. grant No. HL-19580 from the National Heart and Lung Institute aided the preparation of this chapter.

Cvtochrome c Peroxidase TAKASHI YONETANI

I. 11. 111. IV. V. VI. VII.

Introduction . . . . . . . Preparation and Molecular Properties Structural Aspects. . . . . . Enzymic Activity . . . . . . Reaction Mechanism . . . . . Interaction with Cytochrome c . . General Comments . . . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . . . . . . .

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

345 347 348 352 353 356 360

1. Introduction

Yeast cytochrome c peroxidase ( ferrocytochrome c :hydrogen-peroxide oxidoreductase, E C 1.11.1.5) which catalyzes the oxidation of ferrocytochrome c to ferricytochrome c in the presence of hydroperoxide, was disH,Or

+ 2 ferrocytochrome c -+

2 ferricytochrome c

+ 2 OH-

(1)

covered in brewer’s yeast by Altschul et al. ( 1 ) in 1940. These investigators preliminarily identified this enzyme as a soluble cytochrome oxidase from yeast, because dithionite-reduced ferrocytochrome c was apparently oxidized by this enzyme without addition of hydrogen peroxide. Subsequently, these investigators (2) obtained a highly purified preparation of this enzyme and demonstrated that this enzyme contains protoheme and reacts with a stoichiometric quantity of hydrogen peroxide to form a red compound, the absorption spectrum of which is similar 1. A. M. Altschul, R. Abrams, and T. R. Hogness, JBC 136, 777 (1940). 2. R. Abrams, A. M. Altschul, and T. R. Hogness, JBC 142, 303 (1942).

345

346

TAKASHI YONETANI

to that of Compound I1 of horseradish peroxidase (3), and correctly identified this enzyme as cytochrome c peroxidase. The purification method they devised for this enzyme (1, 2) was somewhat tedious and gave a rather poor yield; thus, subsequent investigators failed to obtain this enzyme in such a high purity as the original investigators did. Consequently, the majority of subsequent studies on cytochrome c peroxidase (4-11) were performed with less pure preparations. It was considered from these investigations that the functional properties of cytochrome c peroxidase were essentially identical to those of other protoheme-containing peroxidases, such as horseradish peroxidase (3-9, 11, 12) and turnip peroxidase ( I S ) , except for differences in their substrate specificity. In 1965, Yonetani and Ray (14) obtained a highly purified preparation of cytochrome c peroxidase in an excellent yield using DEAE-cellulose ion exchange chromatography. Shortly thereafter, Yonetani et al. (16) crystallized this enzyme by isoelectric dialysis. Subsequently, Yonetani and co-workers (16-40) carried out a series of extensive investigations 3. D. Keilin and T. Mann, Proc. Roy. SOC.,Ser. B 122, 119 (1937). 4. B. Chance, in “Enzymes and Enzyme Systems” (J. T. Edsall, ed.), p. 93. Harvard Univ. Press,Cambridge, Massachusetts, 1951. 5. B. Chance, Advan. Enzymol. 12, 153 (1951). 6. B. Chance, ABB 21, 416 (1949) ; 22, 224 (1949) ; 24, 389 (1949). 7. B. Chance, ABB 41, 416 (1952). 8. P. George, BJ 54, 267 (1953) ; 55,220 (1953). 9. P. George, JBC 201, 413 (1953). 10. J. Beetlestone, ABB 89, 35 (1960). 11. P. Nicholls, ABB 106, 25 (1964). 12. D. Keilin and E. F. Hartree, BJ 48, 88 (1951). 13. T. Hosoya, J. Biochem. ( T o k y o ) 47, 369 (1960). 14. T. Yonetani and G. S. Ray, JBC 240,4503 (1965). 15. T. Yonetani, B. Chance, and S. Kajiwara, JBC 241,2981 (1966). 16. T. Yonetani, JBC 240, 4509 (1965). 17. T. Yonetani and G. S. Ray, JBC 241, 700 (1966). 18. T. Yonetani, JBG 241, 2562 (1966). 19. T. Yonetani and T. Ohnishi, JBC 241, 2983 (1966). 20. T. Yonetani, H. Schleyer, and A. Ehrenberg, JBC 241, 3240 (1966). 21. T. Yonetani, D. F. Wilson, and B. Seamonds, JBC 241, 5347 (1966). 22. T. Yonetani, H. Schleyer, B. Chance, and A. Ehrenberg, in “Hemes and Hemoproteins” (B. Chance, R. W. Estabrook, and T. Yonetani, eds.), p. 293. Academic Press, New York, 1966. 23. T. Yonetani and H. Schleyer, JBC 242, 1974 (1967). 24. T. Yonetani and H. Schleyer, JBC 242, 3919 (1967). 25. T. Yonetani and H. Schleyer, JBC 242, 3926 (1967). 26. T. Yonetani, JBC 242, 5008 (1967). 27. T. Yonetani and T. Asakura, JBC 243, 3996 (1968). 28. T. Yonetani and T. Asakura, JBG 243, 4716 (1968). 29. T. Asakura and T. Yonetani, JBC 244 537 (1969).

6.

CYTOCHROME

c

PEROXIDASE

347

on its functional properties. Meanwhile, Ellfolk (41) independently obtained a crystalline preparation of cytochrome c peroxidase and investigated its molecular properties (42, 4s).

II. Preparation and Molecular Properties

Cytochrome c peroxidase is found exclusively in aerobically grown yeast (1, 2, 19, 44). Chantrenne (43) demonstrated that the synthesis of this enzyme in anaerobically grown yeast is induced by exposure to oxygen. Sels and Cocriamont (44) showed that apocytochrome c peroxidase is synthesized in anaerobically grown yeast and that oxygen is required for the in vivo conversion from the apoenzyme to the holoenzyme. Commercially available yeast, such as baker’s and brewer’s yeasts, are the most convenient sources for large-scale preparations of cytochrome c peroxidase. However, the content of this enzyme in yeast varies considerably depending on the type of strain and culture conditions ; thus, preliminary checks to find the brand of commercial yeast, which is rich in cytochrome c peroxidase, is advised before starting a large-scale preparation. The detailed method of chromatographic purification of cytochrome c peroxidase has been described elsewhere (16, 15, 45). The enzyme can be readily extracted with aqueous solvents from yeast that has been lysed in the presence of organic solvents such as toluene and ethylacetate. The extracted enzyme is concentrated by salting out with saturated ammonium sulfate and purified by two cycles of DEAE-cellulose chromatography to yield a preparation with a purity index (the ratio of absorbance at 408 nm to that a t 280 nm) of 0.7to 1.1. Repeated chro30. T. Asakura and T. Yonetani, JBC 244, 4570 (1969). 31. T. Yonetani and T. Asakura, JBC 244,4580 (1969). 32. T. Asakura, H. R. Drott, and T. Yonetani, JBC 244,6626 (1969). 33. T. Yonetani, Aduun. Enzymol. 33, 309 (1970). 34. A. F. W. Coulson, J. E. Erman, and T. Yonetani, JBC 246, 917 (1971). 35. T. Iizuka, M. Kotani, and T. Yonetani, JBC 246,4731 (1971). 36. A. F. W. Coulson and T. Yonetani, BBRC 48, 391 (1972). 37. T. Asakura and T. Yonetani, JBC 247,2278 (1972). 38. R. K. Gupta and T. Yonetani, BBA 29.2, 502 (1973). 39. J. Leonard and T. Yonetani, Biochemistry 13, 1465 (1974). 40. N. Ellfolk, Acta Chem. Scand. 21, 175 (1967). 41. N. Ellfolk, Actu Chem. Scand. 21, 1921 (1967). 42. N. Ellfolk, Actu Chem. Scand. 21, 2736 (1967). 43. H. Chantrenne, BBA 18, 68 (1955). 44. A. A. Sels and C. Cocriamont, BBRC 32, 192 (1968). 45. T. Yonetani, “Methods in Enzymology,” Vol 1 p. 336, 1967.

348

TAKASHI YONETANI

matography does not increase the purity. Cytochrome c peroxidase is readily crystallized from the enzyme preparation have a purity index of more than 0.7 by exhaustive dialysis against distilled water. Cytochrome c peroxidase with a purity index of 1.25 to 1.30 can be prepared by two or three cycles of this isoelectric crystallization. The molecular weight of cytochrome c peroxidase has been determined to be 34,100on the basis of a sedimentation constant of 3.55 S, a diffusion constant of 9.44 F, and a partial specific volume of 0.733 ml/g (49). The enzyme exists as a monodisperse monomer containing one ferric protoporphyrin IX, which is noncovalently bound ( 1 , 2, 1 4 ) . No other transition metal is detected in crystalline preparations of the enzyme ( 2 2 ) .The apoenzyme moiety is an acidic protein with an isoelectric point a t p H 5.0 to 5.2,which is made of 272 amino acid residues: Asp,,, Thr,,, Ser,,, G~uz,,Prol5, Glyz3, A h , Vallz9 Mets, h,Leuz3, Tryl2, Phe16, Lys21, His,, Arg,, Cys,, and Try, (37, 4 3 ) . The N- and C-terminal residues have been identified as threonine and leucine, respectively. The sulfhydry1 group of the cysteine residue reacts with sulfhydryl reagents more rapidly in the apoenzyme than in the holoenzyme. This site can be blocked by p-mercuribenzoate without affecting its enzymic activity. It should be noted that cytochrome c peroxidase contains no carbohydrate (41) , in contrast to other protoheme-containing peroxidases. Crystalline cytochrome c peroxidase has unit cell dimensions of 108.0X 77.8 X 51.4A, a space group of P2,2,2,, and a 2 value of 4 (molecules per unit cell) (46).Several heavy atom derivatives, whose crystalline structure are isomorphous with that of the native enzyme, have been prepared for the X-ray diffraction analysis of crystalline cytochrome c peroxidase. Cytochrome c peroxidase has been reversibly resolved into protoheme and apoenzyme moieties (26) by a modification of Teale's acid butanone technique (47).The apoenzyme may be combined with porphyrins and metalloporphyrins to form synthetic holoenzymes containing unnatural prosthetic groups (9793, 38-40). The apoenzyme (96') and some of the synthetic holoenzymes have been crystallized. 111. Structural Aspects

Since the primary and three-dimensional structures of cytochrome c peroxidase are yet to be established, any discussion on its molecular 46. L. 0. Larsson, L. 0. Hagman, P. Kierkegaard, and T. Yonetani, JBC 245, 902 (1970). 47. F. W. J. Teale, BBA 35, 543 (1959).

6.

CYTOCHROME

c

PEROXIDASE

349

structure becomes somewhat speculative a t the present time. However, the indirect experimental data available today are sufficiently numerous to discuss a definitive molecular structure for this enzyme, particularly a t the heme region. Unlike the globins from myoglobin and hemoglobin, apocytochrome c peroxidase is resistant to freezing-induced denaturation and thus can be stored frozen for an indefinite period in the heme-free state. Apocytochrome c peroxidase has been crystallized with a habit of crystallization apparently identical with that of its holoenzyme (26).The crystal of the protoporphyrin IX-apoenzyme complex has been demonstrated to be isomorphous with the crystal of the native holoenzyme ( 4 6 ) . These observations suggest that the gross conformation of apocytochrome c peroxidase is not significantly affected by the incorporation of the prosthetic group. An anionic aromatic dye, anilino-naphthalene sulfonic acid, interacts with apocytochrome c peroxidase to form a stoichiometric complex in competition with heme ( 4 8 ) . This interaction results in a significant enhancement of the fluorescence emission of the dye, indicating a hydrophobic nature of the dye- or heme-binding site of this enzyme in a similar manner to those of myoglobin and hemoglobin ( 3 9 ) .The chemical modification of heme side chains a t positions 2 and 4 has no appreciable effect on either the enzymic activity or the affinity of the apoenzyme for the prosthetic group (28).On the other hand, both the enzymic activity and the affinity for the prosthetic group are greatly reduced by modification of one or two propionic acid chains a t positions 6 and/or 7 (3U,32). On the contrary, the modification of heme side chains a t positions 6 an 7 has no appreciable effect on the oxygen affinity of myoglobin and hemoglobin as well as the subunit cooperativity in the latter, whereas the modifications a t positions 2 and 4 influence both the oxygen affinity and the subunit cooperativity (49, 50). Since two propionic acid side chains of protoheme are exposed to the exterior of the molecule in myoglobin and hemoglobin (51,52),it is tempting to speculate that the heme group of cytochrome c peroxidase may be embedded in a direction opposite t o that of myoglobin and hemoglobin. I n other words, the heme side chains a t positions 2 and 4 are exposed to the exterior, whereas propionic acid side chains a t positions 6 and 7 face the interior of the molecule in cytochrome c peroxidase (Fig. 1). 48. L. Stryer, J M B 13, 482 (1965). 49. A. Rossi-Fanelli, E. Antonini, and A. Caputo, Advan. Protein Chem. 19, 73 (1964). 50. Y. Sugita and Y. Yoneyama, JBC 246,389 (1971). 51. J. C. Kendrew, Brookhaven Symp. Biol. 15, 216 (1962). 52. M. F. Perutz, J M B 13, 646 (1965).

350

TAKASHI YONETANI

FIQ.1. A proposed orientation of the protoheme prosthetic group in cytochrome c peroxid-.

The nitric oxide complex of ferrocytochrome c peroxidase exhibits an electron paramagnetic resonance (EPR) spectrum of Thombic symmetry (53). Its hyperfine structure has been interpreted as derived from the interaction of the heme iron with two types of axially bound nitrogen atoms, one obviously from nitric oxide and the other from a protein group. The magnitude of the latter hyperfine coupling is consistent with the proposed assignment of a histidyl nitrogen as the fifth ligand of the prosthetic group (Fig. 1). Rose Bengal, an anionic aromatic dye, like anilinonaphthalene sulfonic acid, stoichiometrically reacts with apocytochrome c peroxidase in competition with protoheme ( 5 4 ) .The bound rose bengal acts as an effective photooxidation sensitizer. Photooxidation of the Rose Bengal-apoenzyme complex results in specific modifications of one tryptophan and two histidine residues per mole of protein, only one of the latter being essential to the enzymic activity. Apparently these residues are located adjacent to the bound Rose Bengal in the heme crevice. One of the destroyed histidine residues may be the fifth ligand. The sixth coordination position of the heme iron in cytochrome c peroxidase, which is normally occupied by H,O, is available for reactions with extraneous ligands such as fluoride, cyanide, and azide, as well as substrates and substrate analogs. The acidic-alkaline transition of a hemoprotein, which is caused by the ionization of the bound water ligand, is usually accompanied by significant spectral changes. However, the visible absorption spectrum of cytochrome c peroxidase is not appreciably 53. T. Yonetani, H. Yamamoto, J. E. Erman, J. S. Leigh, and G. Reed, JBC 247, 2447 (1972). 54. A. F. W. Coulson and T. Yonetani, Etrr. J. Biochem. 26, 125 (1972).

6.

CYTOCHROME

CHEMICAL

351

c PEROXIDASE

AND

TABLE I ELECTRONIC EQUILIBRIA OF CYTOCHROME

C

PEROXIDASE

Chemical compound Electronic state “Thermally excited”

I

Activation energy (.I Transition temperature (T,) “Ground”

“Acidic” CCP(FelI1) H 2 0 Low- spin (S

=

1 c = 1230 cm-*

8)

T , = 274°K ( + l ” C )

1

High- spin (S

=

9)

“Alkaline” CCP(FelI1) OH-

-pK 6.3

High- spin (S

=

1

8)

E

=

1830cm-l

T,

=

232°K (-41°C)

1

Low- spin (S

=

8)

affected by changes in pH at room temperature. This anomaly has been resolved by spectrophotometric ( 2 1 ) , EPR ( 2 2 ) ,and magnetic susceptibility (55) measurements of this enzyme a t cryogenic temperatures. The above-mentioned pH-insensitive absorption spectrum of this enzyme can be adequately interpreted by the assumption that cytochrome c peroxidase is a mixture of two chemical compounds (“acidic” and “alkaline” forms), each of which is an electronic mixture of thermally excited and ground states a t acidic and neutral pH regions and room temperature, as indicated in Table I ( 5 6 ) . Since both [‘acidic” and “alkaline” forms are mixtures of respective high- and low-spin electronic states and thus exhibit more or less similar absorption spectra a t room temperature, the rracidic”-L‘alkaline”transition of this enzyme cannot be readily monitored by spectrophotometry a t ambient temperatures. Only when both “acidic” and “alkaline” forms are brought to their respective lowest ground states by lowering the temperature below -lOO°C, do they become pure “high-spin” and “low-spin” states with distinctive spectral characteristics (5‘7). Thus, its “acidic”-“alkaline” transition may be readily followed by cryogenic spectrophotometry (21). Cytochrome c peroxidase becomes unstable and irreversibly modified on standing above pH 8. Thus, it is difficult to follow the transition quantitatively especially a t the alkaline end. Nevertheless, the pK value of its transition is estimated to be about 6.3, which is significantly lower than those of myoglobin and hemoglobin. 55. T. Iizuka, M. Kotani, and T. Yonetani, RBA 167, 257 (1968). 56. T . Iizuka and T. Yonetani, Advan. Biophys. 1, 155 (1970). 57. A. S. Brill and R. J. P. Williams, BJ 78,253 (1961).

352

TAKASHI YONETANI

IV. Enzymic Activity

The reaction of cytochr me c p roxid se follows a general form of compulsory order mechanism for two-substrate enzymes as indicated by Eqs. ( 2 ) , ( 3 ) , and (4) where E is the enzyme, S, is hydroperoxide, Szis ferro-

ka + Sz r ESISZ kr ESzSz E + P

ESi

(3) (4)

+

cytochrome c, and P is ferricytochrome c. Since S, and S, are two-equivalent oxidant and one-equivalent reductant, respectively, the overall reaction requires a stoichiometry of Eq. (1). Furthermore, the reaction is practically irreversible. I n other words, the reverse rates, if they exist, are negligibly small in comparison with the corresponding forward rates. Within these restrictions, some of these rate constants have been estimated from initial steady-state rate measurements, as summarized in Table I1 ( 1 7 ) .The association rates (k,) of the enzyme and hydroperoxides are an order of magnitude faster than the corresponding rates reported for horseradish peroxidase ( 6 ) . The association rates of the ES1 intermediate and ferrocytochrome c (S,) of lo8 M-'sec-' are one of the most rapid rates recorded for protein-protein interactions and closely approach the theoretical limit set of the collision rate of these two proteins (92). TABLE I1 KINETIC CONSTANTS OF PEROXIDATIC OXIDATIONOF FERROCYTOCHROME c CATALYZED B Y CYTOCHROME c PEROXIDASE~ Kinetic constant

Substrate Heart ferrocytochrome c HzOz C,HsOOH Yeast ferrocytochrome c HzOz C zH 6 0 0 H a

ki (M-1 sec-1)

kz

ka

(M-1 sec-1)

(sec-1)

K, (M)

1 . 4 X lo8 2 . 5 X lo7

5 . 9 X lo8 5 . 0 X lo8

2 . 6 X loa 2 . 0 X 10'

4 . 5 X lo-' 4 . 1 X lo-'

1 . 2 X lo8 2 . 2 x 107

5 . 6 X lo8 5 . 2 x 108

1 . 4 X 10' 1 . 2 x 104

2.5 X 2 . 3 x 10-6

Assays were made in sodium acetate buffer, pH 6.0, at 23°C.

6.

CYTOCHROME c PEROXIDASE

353

Cytochrome c peroxidase is reversibly inhibited by ligands that can combine with ferric heme-such as fluoride, azide, and cyanide-but it is not inhibited by carbon monoxide, which indicates that the formation of the ferrous form of the enzyme is not involved in its reaction cycle. The activities of cytochrome c oxidase and cytochrome c peroxidase in tissue may be readily differentiated by the use of carbon monoxide since only the former enzyme is selectively inhibited by carbon monoxide (19). Cytochrome c peroxidase can oxidize a number of reducing agents such as ascorbate, pyrogallol, guaiacol, hydroquinone, and ferrocyanide as well as mammalian and yeast ferrocytochrome c (1, 4, 1 4 ) . Ferrocytochromes b, b,, and c, from mammalian sources and a majority of bacterial ferrocytochrome c are not oxidized. The molecular activity of cytochrome c peroxidase for mammalian and yeast ferrocytochrome c (k,= lo4 sec-’) is a t least two orders of magnitude larger than those for other reductants. Thus, this enzyme has a high substrate specificity toward ferrocytochrome c. Alkylhydroperoxides, such as methyl-, ethyl-, and propylhydroperoxides, can be effectively utilized as S, by this enzyme.

V. Reaction Mechanism

Altschul et al. (1, 2 ) originally discovered that cytochrome c peroxidase reacts with a stoichiometric amount of hydroperoxide to form a red “peroxide compound,” which will be referred t o hereafter as Compound ES. It has a distinct absorption spectrum, as shown in Fig. 2. The formation of Compound ES from the enzyme and hydroperoxides is very rapid (k, > lo7 los M-’ sec-I). No intermediate, which precedes Compound ES, has been thus far detected. In the absence of reductants, or S,, Compound ES is highly stable. The rate constant of its spontaneous decay is of the order of sec-l ( 2 2 ) . The primary peroxide compound (Compound I) of horseradish peroxidase decays much faster a t a rate of sec-’ (6). This unusual stability of Compound ES allows one to determine various physical and chemical parameters quantitatively and reliably. Titrations of Compound ES with reductants such as ferrocytochrome c (16, 20) and ferrocyanide (18, 34) have established that Compound ES is two oxidizing equivalents above the original ferric .enzyme. The absorption spectrum of Compound ES is essentially identical to that of Compound I1 of horseradish peroxidase which contains one oxidizing equivalent per mole in the form of Fe(1V). I n addition, EPR examinations have revealed that Compound ES contains a stable free radical, the spin concentration of which is approximately one equivalent per mole (Fig. 3 ) . Therefore, it is reasonable to conclude that two oxidizIU

354

TAKASHI YONETANI

Wavelength (nm 1

FIQ.2. Absolute and difference light absorption spectra of cytochrome c peroxidaee and Compound ES at pH 7 and 20": (-1 enzymes and (---I CJLOOH.

+

ing equivalents in Compound ES are maintained in the form of Fe(1V) and a free radical of a protein group (R"). Compound ES formed from alkyl-hydroperoxide is indistinguishable from that derived from hydrogen peroxide. Furthermore, the formation of Compound ES upon reaction with ethyl hydroperoxide is accompanied by the release of 1 mole of ethyl alcohol per mole. These observations indicates that the 0-0 bond in S, has been broken upon the formation of Compound ES and that Compound ES is not a so-called reversible enzyme-substrate complex as ES, in Eq. (2) implies, but an enzyme intermediate carrying two oxidizing equivalents per mole in a form other than the original substrate (S,). Reduction of Compound ES with 2 moles of ferrocytochrome c generates the original enzyme rapidly. It has not been possible to detect the formation of the one-equivalent, ferrocytochrome c-reduced intermediate of Compound ES and to determine the rate constants of reactions of

6.

CYTOCHROME c PEROXIDASE

355

i I

0

12.004

6.00

I000

-

2000

3000

4000

51 0

T 1

H

._

( ocrstcmdr)

FIG.3. Electron paramagnetic resonance absorption spectra of cytochrome c peroxidase and Compound ES at pH 7 and at 77°K: (-) enzyme and (---) Compound ES.

Compound ES with first, and second moles of ferrocytochrome c individually. However, using ferrocyanide as a reductant, it has been possible to examine the mechanism of reduction of Compound ES to the original enzyme in detail ( 3 4 ) . Comparison of optical and EPR titrations shows that the reaction of Compound ES with ferrocyanide in a range from pH 5 to 8 is biphasic and strongly supports a mechanism in which two oneequivalent intermediates are a t rapid equilibrium, as shown in Eq. (5) Fe(1V) Fe(II1)-R* (5) where Fe(1V) and Fe(II1) are ferry1 and ferric heme irons and R" is a protein free radical. Optical and EPR parameters thus far available are not sufficient to identify the chemical nature of the protein group responsible for R". However, the spontaneous decay of Compound ES results in destruction of several amino acid residues (36).At pH 4 and 8, tyrosine and tryptophan are the residues principally affected. Thus, it is possible that one of these residues may be responsible for the forma-

356

TAKASHI YONETANI Horseradish peroxidase

2-eq. Oxidation level

1-eq. Oxidation level

Original ferrlc level

Comp. ES [ Fe(IV)-R*l (red)

Comp. I[Fe(IV)-P'] (green)

RoH4

\-/;mp.

ROOH

Cytochrorne c peroxidase

IIlFe(IV)J (red)

ROOH

HRP [Fe(UI)] (brown)

CCP[ Fe(iII)l (brown)

FIG.4. Reaction cycles of horseradish and cytochrome c peroxidases.

tion of the free radical, R", in Compound ES [Fe(IV)-R*] and its oneequivalent reduced form [ Fe (111)-R"] . Low-temperature magnetic susceptibility (55) and Mossbauer spectroscopic (58) data are consistent with assumption that Compound ES contains Fe(1V). On reaction with a stoichiometric amount of hydroperoxide, catalase and horseradish peroxidase are converted to a green colored intermediate, Compound I ( 5 ) . The chemical nature of Compound I has been extensively debated since its discovery by Theorell (59). Recently, Dolphin et al. (60) have demonstrated that upon one-equivalent oxidation several metalloporphyrins are converted to stable porphyrin *-cation radicals, the absorption spectra of which possess the spectral characteristics of Compound I, namely, a decreased Soret n-x" transition and an appearance of the 620-670-nm absorption bands. Since Moss et al. (61) proposed the presence of Fe(1V) in Compound I of horseradish peroxidase from Mossbauer spectroscopic measurements, it is attractive to describe Compound I as Fe (IV) -P", where P" is a porphyrin =-cation radical. Then, Compound I and Compound ES become isoelectronic. Both contain Fe(1V) and a radical: the former as a porphyrin radical (P") and the latter as a protein radical (R"). Then the reaction cycles of horseradish and cytochrome c peroxidases may be compared as shown in Fig. 4.

VI. Interaction with Cytochrome c

The elucidation of the mode of interaction between cytochrome c peroxidase and cytochrome c is not only essential in our understanding of the reaction mechanism of this enzyme but also provides important clues for formulating a general mechanism of electron transfer in biological sys58. G. Lang and T. Yonetani, unpublished results. 59. H. Theorell, Enzymologia 10,250 (1941)t

60. D. Dolphin, R. H. Felton, D. C. Borg, and J. Fajer, JACS 92, 743 (1970). 61. T.H. Moss, A. Ehrenberg, and A. J. Bearhen, Biochemistry 8,4159 (1969).

6.

CYTOCHROME

c

PEROXIDASE

357

tems such as mitochondria1 and microsomal electron transfer systems. The question of whether or not the direct contact between two prosthetic groups is a prerequisite for the electron transfer processes in biological reactions is long-standing and yet to be answered. This problem becomes experimentally approachable by the use of the cytochrome c peroxidase-cytochrome c couple. The formation of a reversible Michaelis-Menten-type complex of the enzyme and ferrocytochrome c [ES,S, in Eq. (3) ] can be postulated from initial steady-state kinetics of the cytochrome c peroxidase reaction ( 1 7 ) . Since cytochrome c peroxidase and cytochrome c are acidic and basic proteins, respectively, their interaction may be governed principally by electrostatic attraction. This assumption is further supported by the fact that several polycations which reversibly and irreversibly bind cytochrome c peroxidase inhibit its enzymic activity in competition with ferrocytochrome c ( l 7 , 6 2 ) . The 220-MHz proton-NMR spectrum of horse heart ferricytochrome c is well characterized by the presence of two low-field methyl resonances a t -35 and -32 ppm and two other relatively sharp high-field methyl resonances a t +2.4 and +2.7 ppm relative to the standard 2,2-dimethyl2-silopentane-5-sulfonate reference at 25O ( 6 3 ) . The two low-field resonances a t -35 and -32 ppm (Fig. 5 ) have been assigned to two methyl side chains of the heme group a t positions 8 and 3, respectively (6'4) (see Fig. 6 ) . I n the presence of equimolar amounts of cytochrome c peroxidase, the linewidths of these two low-field resonances broaden from 20 to -100 Hz. This is accompanied by a decrease in the separation between these two resonances from 3.1 to 2.25 ppm, as the consequence of mutual shifts of the -35 and -32 ppm resonance to up- and downfield directions, respectively. Nuclear magnetic resonance (NMR) titrations of ferricytochrome c with cytochrome c peroxidase as a function of either the resonance linewidths or AV give a stoichiometry of 1 : l to confirm the formation of a stoichiometric complex between these two macromolecules (38).The observed approximately fourfold broadening of the resonance linewidths is consistent with the expected change in the tumbling correlation time of the whole molecule upon a 1 : 1 complexation. When cytochrome c is present in an excess over cytochrome c peroxidase, a time-averaged NMR spectrum of free and complexed ferricytochrome c rather than a simple superposition of two distinction spectra is observed. This indicates that the association and dissociation rates for the 62. E. Mochan and P. Nicholls, BBA 216,SO (1970). 63. K. Wiithrich, Proc. Nut. Acad. Sci. U . S . 63, 1071 (1961). 64. A. G. Redfield and R. K. Gupta, Cold Spring Harbor Symp. Quant. Biol. 36, 405 (1971).

358

TAKASHI YONETANI

1

I

I

36

34

32

Field shifts in PPM

FIQ.5. ‘H-NMR spectra of ferricytochrome c in the presence (b) and absence (a) of an equimolar cytochrome c peroxidase.

FIG.6.A proposed peroxidaae binding site in cytochrome c.

6.

CYTOCHROME

c

PEROXIDASE

359

peroxidase-cytochrome c complex must be much greater than the inverse of the frequency separation between complexed and free states. It is possible to set a lower limit of 200 sec-I on the dissociation rate, which is approximately equivalent to k-, of Eq. (3). The complexation induced shifts of the -35 and -32 ppm resonances to -35.2 and -33 ppm, respectively (Fig. 5), indicate that the spin densities on methyl groups a t positions 8 and 3 are decreased and increased by approximately 276,respectively. This could be merely the electrostatic effect of negatively charged cytochrome c peroxidase which repels the unpaired electron cloud in the complex away from it implying that cytochrome c peroxidase interacts with ferricytochrome c in the area adjacent to the methyl side chain a t position 8. I n other words, the enzyme approaches cytochrome c in a general direction of 7 o'clock from the back side, as indicated by the arrow in Fig. 6. I n this mutual orientation the direct contact between the prosthetic groups of these two hemoproteins will be prevented by the backbone peptide chain of cytochrome c. It is also possible that the binding of the peroxidase produces a conformational change which alters the resonance positions. It is further observed that the linewidths of the resonances in the cytochrome c peroxidase-cytochrome c complex are not sensitive to changes in the electronic structure (high-spin or low-spin) and consequently also the electronic relaxation time of the heme iron of cytochrome c peroxidase upon complexing with fluoride and cyanide. These observations indicate that the heme group of cytochrome c is a considerable distance from the heme iron of the enzyme in the complex (>25 A, assuming an electronic relaxation time of the heme iron of cytochrome c peroxidase of approximately 10-losec). The fluorescence studies of the interaction of cytochrome c with the anilinonaphthalene sulfonate-apoenzyme and protoporphyrin-apoenzyme complexes provide another line of evidence (39) in support of the abovementioned conclusion. Both fluorescence steady-state and lifetime titrations of these fluorescence-labeled apoenzymes with ferro- and ferricytochrome c indicates the formation of a 1:l complex, the affinity for ferricytochrome c being less than that for ferrocytochrome c. From the phosphorescence and fluorescence quenching, the distance between the emitter (a fluorescence label) and the quencher (the heme of cytochrome c ) can be calculated by assuming that no direct electronic interaction exists between them, and that the quenching is derived from the Forster-type energy transfer (65) to the heme. The distance from the apoenzymebound emitter to the heme group of cytochrome c is estimated to be 19 A and 14 W for the anilinonaphthalene sulfonate and protoporphyrin 65.

T.Forster, Discuss. Faraday SOC. 27, 7 (1959).

360

TAKASHI YONETANI

emitter, respectively. The latter should correspond to the heme-heme distance in the complex of the holoenayme and cytochrome c. These NMR and fluorescence data preclude the electron transfer mechanism through a direct contact of two prosthetic groups. The possibility of electron transfer via the polypeptide chains or quantum mechanical tunneling must be seriously considered. VII. General Comments

The molecular structures of oxygen-carrying myoglobin and hemoglobin as well as electron-transferring cytochromes, which are made of relatively small apoproteins (a molecular weight of less than 20,000 per heme-site), have been determined a t atomic levels. However, those of heme enzymes such as peroxidases, catalases, and oxidases, which contain larger apoproteins (a molecular weight of more than 30,000 per hemesite), have not been established yet. Cytochrome c peroxidase provides the opportunity not only to solve the molecular structure of a heme enzyme but also to elucidate the mode of interaction of two hemoproteins by X-ray crystallography, although cocrystallization of the peroxidase with cytochrome c has not yet been accomplished. Various evidences thus far available point to the substantial difference in the molecular structure, especially in the heme region, between these two categories of hemoproteins, as discussed in Section I11 and Fig. 1. Recent metal substitution studies have shown that the chemical substitution of heme iron with manganese produces enzymically active artificial peroxidases (27, 31) but not oxygen-carrying artificial myoglobin and hemoglobin (31, 66, 67) and that the substitution of heme iron with cobalt generates oxygen-carrying artificial myoglobin and hemoglobin (68, 69) but not enaymically active artificial peroxidases (70). These observations indicate that the chemical nature of apoprotein and the mode of its interaction strongly control the physical and chemical characteristics of the prosthetic group in holohemoproteins. The detailed chemical mechanism of the interaction between cytochrome c peroxidase and hydroperoxides to form Compound ES must be further elucidated. The use of substrate analogs such as various oxidia66. T. Yonetani, H. R. Drott, J. S., Jr. Leigh, G . Reed, T. Asakura, and M. R. Waterman, JBC 245, 2998 (1970). 67. M. R. Waterman and T. Yonetani, JBC 245,5847 (1970). 68. B. M. Hoffman and D. H. Petering, Proc. N a t . Acad. Sci. U . S . 67, 637 (1970). 69. T. Yonetani, H. Yamamoto, and G . V. Woodrow, 111, JBC 249, 691 (1974). 70. T. Yonetani, T. Iizuka, and H. Yamamoto, JBC 249,2168 (1974).

6.

CYTOCHROME

c PEROXIDASE

361

ing agents (8, 9 ) , aromatic peracids (71) as well as isotope-substituted substrates may offer the opportunity to effectively shed a light on this problem. The true physiological role of cytochrome c peroxidase in yeast is yet to be established. It may serve as a part of the systems which prevent intracellular accumulation of harmful hydrogen peroxide. It would be of interest to know if cytochrome c peroxidase is synthesized concurrently with or in competition with the production of other peroxide-decomposing systems such as catalase. Although cytochrome c peroxidase is present in mitochondria of aerobically grown yeast in a concentration comparable to that of cytochrome oxidase (19) and possesses an extremely high molecular activity (k, = lo4 sec-') toward yeast ferrocytochrome c ( 1 7 ) , it has not been unequivocally shown that ferrocytochrome c is a true substrate of this enzyme. ACKNOWLEDGMENTS This investigation has been supported by NSF grant (BMS73-00970), NHLI grant (HL-14508), and NIAAA grant (AA-00292).

71. G. R Schonbaum and S. Lo, JBC 247,3353 (1972).

This Page Intentionally Left Blank

Catalase GREGORY R. SCHONBAUM

BRITTON CHANCE

I. Introduction . . . . . . . . . . . . . . . . 11. General Enzyme Properties . . . . . . . . . . . . 111. The Nature of the Active Site . . . . . . . . . . . A. Identity of Ligands at the Fifth and Sixth Coordination Positions of the Prosthetic Group . . . . . . . B. Identity of a Distal Ligand: Selective Modifications of the Apoprotein . . . . . . . . . . . . . . C. Ligand Exchange Reactions . . . . . . . . . . IV. Catalase-Mediated Redox Reactions . . . . . . . . . A. The Nature of Compound I . . . . . . . . . . B. The Catalase Reaction Mechanism. . . . . . . .

363 366 369 369 376 385 388 389 390

1. Introduction

A quarter of a century has passed since the first contribution on catalase to “The Enzymes” : lLEnzyme substrate compounds: Mechanism of action of hydroperoxidases” ( 1 ) . I n this perspective, we can identify a sequence of steps in the development of ideas on the mechanism of enzymic action and the nature of enzyme-substrate compounds. The identification of these compounds and the approach to enzymic reactions a t concentrations stoichiometric with the substrate caused a principal transition of viewpoint on hemoprotein catalysis from free radical mechanisms ( 2 ) unrelated to an active center toward the acceptance of catalysis occurring at the iron atom of the porphyrin (3-5). The latter concept followed natu1. B. Chance, “The Enzymes,” Vol. 2, Part 1, p. 428, 1951. 2. C. Oppenheimer and K. G. Stern, “Biological Oxidation.” Junk, The Hague, 1939. 3. 0.Warburg, “Heavy Metal Prosthetic Groups and Enzyme Action” (A. Lawson, transl.). Oxford Univ. Press (Clarendon), London and New York, 1949. 363

364

GREGORY R. SCHONBAUM AND BRITTON CHANCE

rally from the impetus of Otto Warburg’s emphasis on iron itself (S), together with enlargements of this idea to include the heme caused by the broader views of David Keilin ( 4 ) . Their ideas on the iron atom as the active center of the heme, which serves as a “vise” to hold the iron in place, were focal points in the structural approach to the nature of enzyme-substrate compounds; the protein was held to play a secondary role. Thus, in the 1930’s and 1940’s, the experimental approaches-particularly those of Theorell (6) and Pauling (?)-were focused upon the valence state of the iron, the magnetic properties of the enzyme-substrate compounds, and the ionic or covalent nature of the intermediates (6,7).It remained for X-ray structure studies of the heme region of myoglobin (8) to bring appropriate attention to the active site as a special environment necessary for enzymic action. The present review emphasizes the nature of this site, the environment it affords for the heme and for the oxidizing or the reducing substrate, and the nature of the energy barriers through which the substrates must pass in order to react a t the iron atom. Other studies concerned the chemical nature of the enzyme-substrate intermediate. It was early recognized that “the precise formula (for the enzyme-substrate complex) may differ from that of a simple iron peroxidc complex” ( 5 ). The development of ideas on electron delocalization and, indeed, electron transfer in oxygen and peroxide compounds of hemoproteins has followed over this interval, slowly a t first, beginning with the work of George on myoglobin and peroxidase compounds (9) and reaching a much broader based generality with infrared studies of ferrous iron-oxygen compounds (10). These approaches, together with belated X-ray and nuclear magnetic resonance studies of the structure of the protein (11) and of the active site of the hydroperoxidases ( l a ) , together 4. D. Keilin, in “The History of Cell Respiration and Cytochrome” (J. Keilin, ed.). Cambridge Univ. Press, London and New York, 1966. 5. B. Chance, in “Biological Antioxidants” (C.G . MacKenzie, ed.), p. 54. Josiah Macy, Jr. Found., New York, 1950. 6. H. Theorell, Advan. Enzymol. 7 , 265 (1947). 7. L. Pauling, “Nature of the Chemical Bond and the Structure of Molecules and Crystals: An Introduction to Modern Structural Chemistry.” Cornell Univ. Press, Ithaca, New York, 1960. 8. L. Stryer, J. C. Kendrew, and H. C. Watson, J M B 8,96 (1961). 9. P. George, in “Currents in Biochemical Research” (D. E: Green, ed.), Vol. 11, p. 358. Wiley (Interscience), New York, 1956. 10. W. Caughey, Chapter 5, this volume. 11. L. L. Larsson, L. 0. Hagman, P. Kierkegaard, and T. Yonetani, JBC 245, 902 (1970). 12. R. Hershberg and B. Chance. Biochemistry 14,3885 (1975).

7. CATALASE

365

with the multiple methods of electron paramagnetic resonance (13) and infrared spectroscopy (14) of the enzyme-substrate analogues, ensure that the present review will be only a milestone on the way toward a deeper understanding of iron porphyrin proteins in biological catalysis. Thus this review, which emphasizes the chemistry of the catalase reaction, leads naturally to a second (15) on the “bifunctional” peroxidaticcatalatic nature of catalase activity and its relationship to the physiological function of catalase. The chemistry of the colorful, perplexing, and challenging problem of catalase mechanism has been set forth in numerous reviews. Those by Brill (16), Nicholls and Schonbaum ( I $ ) , and most recently, Deisseroth and Dounce (18) summarize the fundamental properties of catalase and its reactions, and their physiological implications. Further, Feinstein (19) and Aebi (20-22) have presented detailed evaluations of acatalasemia, and de Duve (23) and others have discussed catalase biosynthesis (23-26), its intracellular location (23-25) and its turnover (24, 26-28). These facets of the catalase problem will not be reiterated. Instead, a brief synopsis of the enzyme characteristics will be followed by a discussion on the nature of the active site and the chemistry of the catalase reaction mechanism. 13. J. S. Leigh, M. Maltempo, P. I. Ohlsson, and K. G. Paul, FEBS (Fed. Eur. Biocnem. Soc.) Lett. 51, 304 (1975). 14. C. H. Barlow, P. I. Ohlsson, and K. G. Paul, Abstr., 170th ACS Meet. (1975). 15. B. Chance, N. Oshino, and H. Sies, in preparation. 16. A. S. Brill, Compr. Biochem. 14, 447 (1966). 17. P. Nicholls and G . R. Schonbaum, “The Enzymes,” 2nd ed., Vol. 8, p. 147, 1963. 18. 19. 20. 21.

A. Deisseroth and A. L. Dounce, Physiol. R e v . 50, 319 (1970). R. N. Feinstein, Biochem. Genet. 4, 135 (1970). H. Aebi and H. Suter, Advan. Hum. Genet. 2, 143 (1971). H. Aebi, E. Bossi, M. Cants, S. Matsubara, and H. Suter, in “Hereditary Disorders of Erythrocyte Metabolism” (E. Beutler, ed.), p. 41. Grune & Stratton, New York, 1968. 22. H. Aebi, in “Isozymes” (C. L. Markert, ed.), Vol. I, p. 227. Academic Press, New York, 1975, 23. C. de Duve, in “Alcohol and Aldehyde Metabolizing Systems” (R. G. Thurman et al., eds.), p. 161. Academic Press, New York, 1974. 24. C. M. Redman, D. J. Grab, and R. Irukulla, ABB 152, 496 (1972). 25. R. S. Holmes and C. J. Masters, ABB 148, 217 (1972). 26. G. L. Jones and C. J. Masters, ABB 161,601 (1974). 27. M. Rechcigl, in “Enzyme Synthesis and Degradation in Mammalian Systems” (M. Rechcigl, Jr., ed.), p. 273. Univ. Park Press, Baltimore, Maryland, 1971. 28. B. Poole, in “Enzyme Synthesis and Degradation in Mammalian Systems” (M. Recheigl, Jr., ed.), p. 375. Univ. Park Press, Baltimore, Maryland, 1971.

366

GREGORY R. SCHONBAUM AND BRITTON CHANCE

II. General Enzyme Properties

Catalase (H,O,:H,O, oxidoreductase; EC 1.11.1.6) was one of the first enzymes to be isolated in a high state of purity, and its crystallization (29) from beef liver extracts ranked among the early triumphs of biochemistry. Extensive physicochemical studies which followed (30-41) led to an impressive elucidation of its properties, but as yet not to a definition of the apoprotein function in the enzyme catalysis. Typically, all carefully characterized catalases are oligomers (isoelectric point -5.5) containing four (42-46) tetrahedrally arranged (47), 60,000-dalton subunits (48-60). Each subunit consists of a single polypeptide chain (41) that associates with a single prosthetic group, ferric protoporphyrin IX (61). The subunits apparently function independently of one another (66,6 3 ) . I n view of the structural complexity of catalase, it is not surprising 29. J. B. Sumner and A. L. Dounce, JBC 121,417 (1937). 30. D. Keilin and E. F. Hartree, Proc. R o y . SOC.,Ser. B 121, 173 (1937). 31. J. B. Sumner and N. Gralen, JBC 125,33 (1938). 32. J. B. Sumner and A. L. Dounce, JBC 127,439 (1939). 33. J. B. Sumner, A. L. Dounce, and V. L. Frampton, JBC 136, 343 (1940). 34. H. Theorell and K. Agner, Ark. Kemi 16, 1 (1943). 35. B. Chance, Acta Chem. Scand. 1,236 (1947). 36. B. Chance, JBC 194, 471 (1952). 37. B. Chance, JBC 194, 483 (1952). 38. J. Yang and T. Samejima, JBC 238,3262 (1963). 39. M. J. Stansell and H. F. Deutsch, JBC 240, 4299 (1965). 40. A. Tasaki, J. Otsuka, and M. Kotani, BBA 140, 284 (1967). 41. W. A. Schroeder, J. R. Shelton, J. B. Shelton, B. Robberson, and G. Appell, ABB 131, 653 (1969). 42. R. E. Greenfield and V. E. Price, JBC 220,607 (1956). 43. A. W. Galston, R. K. Bonnichsen, and D. I. Arnon, Acta Chem. Scand. 5, 781 (1951). 44. H. F. Deutsch, Acta Chem. Scand. 8, 1516 (1952). 45. D. Herbert and J. Pinsent, BJ 43, 193 (1948). 46. D. Herbert and J. Pinsent, BJ 43, 203 (1948). 47. B. K. Vainshtain, S. Ya. Karpukhina, N. I. Sosfenov, and L. A. Feigin, Dokl. Akad. Nauk SSSR 207,1336 (1972). 48. H. Sund, K. Weber, and E. Molbert, Eur. J. Biochem. 1, 400 (1967). 49. K. Weber and M. Osborn, JBC 244, 4406 (1969). 50. H. Furuta, A. Hachimori, Y. Ohta, and T. Samejima, J. Biochem. ( T o k y o ) 76, 481 (1974). 51. K. G. Stern, JBC 112, 661 (1936). 52. B. Chance, JBC 179, 1299 (1949). 53. M. L. Kremer, J. Theor. Biol. 29, 387 (1970).

7.

CATALASE

367

that its biosynthesis occurs stepwise. The synthesis is governed by a single autosomal determinant and proceeds, in the case of (rat) liver catalase, in three distinct stages (23): (a) synthesis of approximately 60,000-dalton apocatalase subunits, (b) intercalation of heme, and (c) formation of tetramers. The resulting oligomer is relatively stable, a property recognized early in the course of its isolation from liver (54, 55) blood erythrocytes, and bacteria (45,46, 56, 5 7 ) . Indeed, dissociation of catalase generally entails its irreversible denaturation [but, see Samejima and Yang ( 5 8 ) ] and occurs most commonly under rather drastic conditions (48): below p H 3 (58, 5 9 ) , above p H 10 (48, 60, 6 1 ) , in the presence of detergents ( 4 8 ) , or following an extensive chemical modification of the apoprotein (50, 6 2 ) . Gentler modifications such as controlled acetylation with N-acetyl imidazole (50) do not result in tetramer dissociation although the subunits do partly unfold. These changes are not accompanied by a pronounced loss of enzymic activity, and the modified derivative, like the native catalase ( S O ) , is nonreducible by dithionite. Despite the size of the protein subunits, their integrity does not depend on cross-linking via disulfide bonds (63) and no disulfide bridges have been identified within the partially completed amino acid sequence (41, 6Sa). Nor is there any evidence that association of subunits depends on covalent bonding; rather, it appears to involve mainly hydrophobic interactions ( 5 0 ) .Of particular interest in this context is the observation that some forms of acatalasemia are attributable to the formation of a catalase variant, of approximately normal specific activity, but with a tendency t o dissociate into subunits ( 6 4 ) . These more recent studies (50, 58) as well as others involving photooxidation (65) of the enzyme, its carboxymethylation ( 6 6 ) , and the deA. L. Dounce, JRC 143, 497 (1942). A. J,. Dounce and V. Frampton, Science 89, 300 (1930). P. Waentig and W. Gierisch, Fermentforschung 1, 165 (1914-1916). M. Tsuchihashi, Biochem. 2 140, 63 (1923). T. Samejima and J. Yang, JBC 238, 3256 (1963). C. Tanford and R. Lovrien, JACS 84, 1892 (1962). 60. T. Samejima, J. Biochem. (Tokyo) 46, 1101 (1959). 61. M. Nagahisa, J. Biochem. (Tokyo) 51, 216 (1962). 62. K. Abe, M. Hiraga, and F. Anan, J. Biochem. (Tokyo) 74,889 (1973). 63. S. Morikofer-Zwea, M. Cantz, H. Kaufmann, J. P. von Wartburg, and H . Aebi, Eur. J. Biochem. 11,49 (1969). 63a. W. A. Schroeder, personal communication. 64. H. Aebi, S. R. Wyss, B. Scherz, and F. Skvaril, Eur. J. Biochem. 48, 137 (1974). 65. M. Nakatani, J. Biochem. (Tokyo) 49,98 (1961). 66. M. Nakatani, J. Biochem. (Tokyo) 48, 476 (1961). 54. 55. 56. 57. 58. 59.

368

GREGORY R. SCHONBAUM AND BRITTON CHANCE

rivatization of its sulfhydryl group (67493, tentatively suggest that (a) the prosthetic group is deeply intercalated but not necessarily strongly bound within the protein matrix; (b) the functional groups a t the active site are not readily accessible to modification; (c) sulfhydryl groups are not essential to enzymic activity; (d) in peroxisomes which are characterized by an extraordinarily high catalase content (69a), dissociation of the enzyme (69b) cannot be central to the modulation of its activity (69c) ; and (e) the perturbations of the protein mantle are not intimately reflected in the catalytic properties of the active site. It seems, therefore, rather unlikely that different conformations of catalase in the high-spin ferric state-inferred from the EPR data (70)-play a key role in the expression of enzymic activity, specifically in (1) the decomposition of hydrogen peroxide into oxygen and water and (2) the peroxide-dependent oxidation of various substrates. The set of possible redox transformations previously observed in catalase-mediated reactions is summarized in Eq. ( 1 ) Ferricatalase

111

pathways

where Compounds I, 11, and I11 are enzyme-peroxide derivatives in formal oxidation states Fe(V), Fe(IV), and Fe(VI), respectively; XHOH, two-electron equivalent donor (reductant) ; AH, one-electron equivalent donor (reductant) ; and ROOH, a hydroperoxide (R = H, alkyl or acyl) . Interpretations of the reactions outlined in Eq. (1) cannot be divorced 67. A. Pihl, R. Lange, and A. Evang, Acta Chem. Scand. 15,1271 (1961). 68. H. R. Schutte and H. Nurnberger, Hoppe-Seyler’s Z. Physiol. Chem. 315, 13 (1959). 69. K. Abe, M. Hiraga, and F. Anan, Bull. Tokyo M e d . Dent. Univ. 14, 309 (1967). 69a. C. de Duve and P. Baudhuin, Physiol. R e v . 46,323 (1966). 69b. P. Jones and R. H. Pain, and A. Suggett, Nature (London) 217, 1050 (1968). 69c. G. H. Barlow and E. Margoliash, BBA 188, 159 (1969). 70. W. E. Blumberg and J. Peisach, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), 2nd ed., Vol. 1, p. 299. Univ. Park Press, Baltimore, Maryland, 1973.

7.

369

CATALASE

from the discussion of the prosthetic group-apoprotein interactions-specifically, those bearing on the nature and labilities of ligands in the inner coordination sphere of the protoheme complex [L, and L,, Eq. ( 2 ) ] , the role of a distal (outer sphere) group YH, and ligand exchange mechanisms. 4-Fe-h

-YH

111. The Nature of the Active Site

A. IDENTITY OF LIGANDS AT THE FIFTHA N D SIXTH COORDINATION POSITIONS OF THE PROSTHETIC GROUP Protoheinin is the major structural component of the active site, and, indeed, is the principal determinant of activity and specificity in hemoproteins. While hemoproteins are generally categorized into the three classes of oxidases, peroxidases, and electron transport components, the first two groups may have a number of features in common in that the oxidase activity may involve certain steps in which peroxide compounds are involved, and the peroxidases may participate in oxidase activities under appropriate conditions. Nevertheless, it is the purpose of this contribution to explore how the apoenzyme determines the fine structure of the catalytic activities of the hemoprotein ( 7 1 ) . In this context, two hypotheses merit consideration: ( 1 ) that constraints imposed by the protein modulate hemin properties (1, 72-74) and (2) that the protein participates intimately in the catalytic function (s) of the enzyme ( 1 7 , 7 5 ) . Both notions are necessarily deductive since the configuration of the apoprotein near the prosthetic group is unknown ; also, because the characteristics of catalase or its derivatives do not lend themselves to singular assignments (74, 76-80) of the proximal ligand (L5,L6; cf. Table I ) and 71. H. Theorell, Ark. Kemi 16A, 1 (1942). 72. P. Nicholls, BBA 60, 217 (1962). 73. P. George and R. J. L. Lyster, Proc. Nat. Acad. Sci. U . S . 44, 1013 (1958). 74. A. S. Brill and H. E. Sandberg, Biophys. J. 8, 669 (1968). 75. P. Jones and A. Suggett, BJ 110, 621 (1968). 76. D. W. Smith and R. J. P. Williams, Struct. Bonding (Berlin) 7, 2 (1970). 77. W. E. Blumberg and J. Peisach, Bioorg. Chem. 1, 271 (1971). 78. W. E. Blumberg and J. Peisach, in “Probes of Structure and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 215. Academic Press, New York, 1971. 79. A. S. Brill and H. E. Sandberg, Proc. Nat. Acad. Sci. U . S . 57, 136 (1967). 80. T. Yonetani and H. Yamamoto, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), 2nd ed., Vol. 1, p. 279. Univ. Park Press, Baltimore, Maryland, 1973.

370

GREGORY R. SCHONBAUM AND BRITTON CHANCE

SOME

TABLE I AXIAL LIGANDS POSTULATED

IN

CATALASE

II

Ls

LS

Carbox yl Tyrosine Nonnitrogenous Histidine (imidazole) Carbox yl Carbox yl

-

-

-

Carboxyl H,O OH (hydrogen bond to YH)

Ref. (6) (7.2)

(80) (74, 77, 7 8 , 7 9 ) (76, 81) (76, 81) (17)

distal (YH) components of the active site (76, 81). There are, however, three viewpoints which are consistent with the imidazole nature of one of the two ligands. Thus, Brill and Sandberg pointed out that whenever imidazole is coordinated to the ferriprotoporphyrin (74, 79, 82) the difference spectra of low-spin vs. high-spin complexes are characterized by an absorption band ~ ~ - ~ ~ ~ 6-12 x lo3 M-’ cm-l). Such below 250 nm ( A c ~ approximately a “diagnostic band,” attributable to charge transfer transitions from L, to porphyrin orbitals, was noted in the difference spectrum of ferricatalase cyanide (low spin) vs. catalase (high spin) ( 7 4 ) ,thereby favoring L, 5His. The same thesis was advanced by Blumberg and Peisach (77, 7 8 ) but for different reasons. From the EPR spectra they computed the rhombicity and tetragonality (8%) characterizing crystal fields of various lowspin hemoproteins and found that for low-spin catalase derivatives such parameters parallel those typical of complexes with a proximal (L,) histidy1 group. Finally, the primary structure of bovine liver catalase ( 4 1 ) , specifically between Gly-215 and Lys-220 ( ~ J u )is, reminiscent of “proximal histidine” sequences, particularly in some cytochromes and globins (Table 11) (41, 81. A. S. Brill and R. J. P. Williams, BJ 78, 246 (1961). 82. H. E. Sandberg and M. S. Balegh, BBA 295,37 (1973). 82a. Rhombicity is a function of the geometry of the complex, whereas tetragonality is governed by the charge density at the ferric ion. Hence, necessarily, it is influenced by the charge characteristics of the apical ligands.

TABLE I1 AMINOACID SEQUENCES OF K N O W N A N D PROPOSED "PROXIMAL PEPTIDES" Helical position in hemoglobino Hemoprotein

F1

Cytochrome cz (Rhodospirillum rubrum) R H P (chromatium) Cytochrome c (man) Catalase (bovine liver) Peroxidase (horseradish) Hemolobin (a)(man) Myoglobin (sperm whale)

-

a

(-)

F2

F3

F4

F5

F6

F7

F8

-

-

-

-

Leu Leu Leu

-

Ala Ala -

Leu Leu Leu

-

-

denotes nonidentical residues; (---) denotes deletions in the polypeptide chain.

F9

FG1 Deleted FG2

Ref.

372

GREGORY R. SCHONBAUM AND BRITTON CHANCE

6Sa, 83-87). This parallelism, although intriguing, could be entirely fortu-

itous. Indeed, other evidence, particularly the EPR signature of the ferrocatalase-nitric oxide complex (80, 88) and the optical spectrum of the free enzyme (76) brings into question the histidyl identity of L,. The EPR spectrum of the f e r r o ~ a t a l a s e - ~ ~ N complex O shows only a single triplet hyperfine structure in the x-absorption due to 14N of NO (80, 88). In contrast to the corresponding spectra of myoglobin, hemoglobin, and some peroxidases, no superhyperfine structure is discernible, suggesting a lack of coupling between the 14Nnucleus and the trans proximal ligand. Whether this rules out histidine a t L, is uncertain. Essentially, the data indicate either that the unpaired electron of NO is not significantly delocalized into L, or that its residence time is negligible in the vicinity of the trans nucleus. Equally debatable are the interpretations of the catalase optical spectrum (76, 81) (Table 111),which is distinguished by a band near 880 nm (74,89) and an exaltation of the transition a t 625 nm. These features, although not typical of aquo hemoproteins when L, = His, are apparent in their “anionic” complexes (Table 111) (74, 79, 80, 89-98) and should also characterize high-spin hydroxo derivatives (99, 100). For these reasons, it seems premature to accept the thesis of Williams et al. (76, 81) that L, = carboxyl. 83. M. 0. Dayhoff, ‘[Atlas of Protein Sequence and Structure,” Vol. 4. Nat. Biochem. Res. Found., Silver Spring, Maryland, 1969. 84. K. Dus, R. G. Bartsch, and M. D. Kamen, JBC 237, 3083 (1962). 85. R. E. Dickerson, T. Takano, D. Eisenberg, 0. B. Kallai, L. Samson, A. Cooper, and E. Margoliash, JBC 246, 1511 (1971). 86. K. G. Welinder and L. B. Smillie, Can. J . Biochem. 50, 63 (1972). 87. K. G. Welinder, Abstr., Int. Congr. Biochem., Sth, 1973 p. 79 (1973). 88. T. Yonetani, H. Yamamoto, J. E. Erman, J. S. Leigh, Jr., and G. H. Reed. JBC 247, 2447 (1972). 89. K. Torii and Y. Ogura, J. Biochem. (Tokyo) 64, 171 (1968). 90. F. L. Jajczay, Ph.D. Thesis, University of Alberta, 1970. 91. M. F. Peruts, P. D. Pulsinelli, and H. M. Ranney, Nature (London), New Biol. 237, 259 (1972). 92. A. Hayashi, T. Suzuki, K. Imai, H. Morimoto, and H. Watari, BBA 194, 6 (1969). 93. J. C. Kendrew, Brookhaven Symp. Biol. 15, 216 (1962). 94. H. C‘. Watson and B. Chance, in “Hemes and Hemoproteins” (B. Chance, R. W. Estabrook, and T. Yonetani, eds.), p. 149. Academic Press, New York, 1966. 95. P. Urnes, Ph.D. Thesis, Harvard University, Cambridge, Massachusetts, 1963. 96. G. I. H. Hanania, A. Yeghiayan, and B. F. Cameron, BJ 98, 189 (1966). 97. M. R. Mauk and A. W. Girotti, Biochemistry 13, 1757 (1974). 98. G. R. Schonbaum, JBC 248, 502 (1973). 99. P. George, J. Beetlestone, and J. S. Griffith, in “Haematin Enzymes” (J. E. Falk, R. Lemberg, and R. K. Morton, eds.), p. 105. Pergamon, Oxford, 1961. 100. K. Yoshida, T. Iizuka, and Y. Ogura, J. Biochem. (Tokyo) 68, 849 (1970).

TABLE I11 SPECTROSCOPIC CHARACTERISTICS OF SOMEFERRIC HEMOPROTEINS Proximal ligands Absorption bands

LS

Hemoprotein

LS

Ref.

Ref.

Xrnax/€rnMa

Catalase (horse erythrocyte)

Not Not identified identified

-

-880 1.1,

625 8.1

505 11.4

HemoglobinMil a@z(67 Glu)

His

COz-

(911

-900

623 6.0

500 10.5

Metmyoglobin (sperm whale)

His

F-

(94)

609 7.8

490 8.3

406 133

(95,96)

860 1.15

Peroxidase (horseradish)

0

His

Hz0

His

Not identified

406 115

(89, 90)

(91992)

(93)

1040 0.89

634 3.6

502 9.9

409 168

(95,96)

(74, 79,80,87,97)

1070 0.61

641 3.2

499 11.3

403 102

(98)

Per hemin: at -pH 7, 25".

W

-a

W

374

GREGORY R. SCHONBAUM AND BRITTON CHANCE

1

I

350

400

450

500

550

600

650

700

X(nm) FIQ. 1. Spectrum of horse erythrocyte catalase (-1

(---) and its formate derivative

; pH 4.6, 25’ (101a).

Less controversial is L,. It is assumed to be a labile rraquo”or hydroxo ligand (17, 37, 72), mainly because such a moiety should be both readily replaceable by exogenous ligands and nonoxidizable. Further, water protons exchange at 2 5 O into the environment of the paramagnetic center at a rate exceeding 2.4 X lo5 sec-’ ( 1 0 0 ~ )Even . this minimum rate is a t least tenfold greater than the rate of proton exchange into the inner coordination sphere of metmyoglobin (101), where “water” is known to occupy the L, position. Hence, a carboxyl group (76, 81) is not necessarily a viable alternative to an aquo ligand. Moreover, catalase shows a remarkable affinity toward exogenous carboxylate ligands such as forM ) (37), and the spectra of the resulting derivatives mate ( K a are distinct (cf. Fig. 1) ( 1 0 1 ~ from ) the spectrum of the free enzyme. Neither should obtain if L, were a carboxyl since in a competition between intra- and intermolecular reactions the former would prevail. The above argument assumes that formate invariably ligates to the ferric ion. Hershberg (102) has recently pointed out that this is not necessarily so, on the basis of NMR studies suggesting that formate interacts

<

100a. A. S.Mildvan and G . R. Schonbaum, unpublished observations. 101. A. S. Mildvan, N. M. Rumen, and B. Chance, in “Probes of Structure and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 205. Academic Press, New York, 1971. 101a. G.R. Schonbaum, unpublished observations. 102. R. L. Hershberg, Ph.D. Thesis, University of Pennsylvania, Philadelphia, 1974.

7.

375

CATALASE

principally with the apoenzyme, albeit in the vicinity (approximately 7 A) of the prosthetic group. Accordingly, he assigned the observed spectroscopic changes to some unspecified perturbation of the hemin environment. The preceding discussion illustrates the continuing uncertainty as to the nature of the L, and L, ligands. The properties of low-spin ferricatalase derivatives are consistent with the presence of histidine a t L, but, seemingly, the characteristics of the free enzyme and of the low-spin ferrocatalase-nitric oxide complex are not. It could be argued, of course, that the orientation of L,, or even its identity, is not invariant in different derivatives, but is mobile or interchangeable. Indeed, Samej ima and Kita (103) studied the optical rotatory dispersion and circular dichroism spectra of the free enzyme and its cyanide and azide derivatives and concluded that these compounds had different amounts of helicity. Still another and equally provocative alternative presents itself if (a) the Ls ligand is hydroxo in character, and (b) the iron atom is displaced toward the sixth coordination position, away from the proximal histidine group. Such a configuration is outlined in formula ( I ) ,where YH remains unspecified but, as shown later, might be another histidine residue.

(1)

Note that in such a configuration: 1. Hydroxyl being a weak field ligand (104), the optical absorption spectrum of ferric catalase is similar to that expected for a high-spin form of hydroxy- or fluorometmyoglobin (99) (Table 111). 2. The nonreducibility of catalase, although a pn’ori not predictable, is a t least understandable ( 1 0 4 ~ )First, . the hydroxy group, L,, should stabilize the ferric state. Second, reduction to a high-spin ferrous ion, with an attendant increase in ion radius and its repulsion from the porphyrin lattice, would not be compensated by proximal interactions, as it 103. T. Samejima and M. Kita, BBA 175, 24 (1969). 104. F. Basolo and R . G. Pearson, in “Mechanisms of Inorganic Reactions,” 2nd ed., p. 67. Wiley, New York, 1967. 104a. It is generally assumed that nonreducibility of catalase by dithionite is of thermodynamic origin. The pathways of dithionite-dependent reductions are, however, by no means simple (106, 106), and it would be unjustified to reject kinetic factors in the catalase-dithionite system. 105. C. Creutz and N. Sutin, Proc. N n t . Acarl. Sci. U . S. 70, 1701 (1973). 106. D. 0. Lambeth and G. Palmer, JBC 248, 6095 (1973).

376

GREGORY R. SCHONBAUM AND BRITTON CHANCB

is in myoglobin or hemoglobin. However, the transition to a spin-paired form, as in ferrocatalase-CO or -NO, should-and does-stabilize the low redox state of the enzyme. 3. In low-spin ferric compounds, e.g., cyanide or azide (106~) complexes, the reduction in the ion radius and incursion of r-bonding should facilitate the entry of iron into the porphyrin plane (108,109) and thereby lead to interactions which typify complexes having a proximal histidine (77,79). 4. In contrast, the lack of superhyperfine splitting in the ferrocatalase-NO complex would not have been foreseen if L, = His. As pointed out by Yonetani et al. (88),a fast relaxation of electron spin may explain this behavior. Alternatively, if catalase-NO were to be a square pyramidal complex, the spin density may not extend to the nitrogen atom of the proximal histidine, and hence no superhyperfine coupling need occur. The above rationalizations rest on the assumption that iron is displaced toward L6, putting, perhaps, undue weight on the premise that enzyme properties are solely governed by the relative positions of L6, L,, and iron. At best, it is but one facet of the possible constraints imposed by the apoprotein upon the environment of the prosthetic group.

B. IDENTITY OF A DISTAL LICAND:SELECTIVE MODIFICATIONS OF THE

APOPROTEIN

In spite of the apparent inaccessibility of the distal site to the typical and traditional protein reagents, some progress has been made in delineating its properties. Pivotal to these developments was the observation by Heim et al. (110)that 3-amino-lH-1,2,4- triazole (AT) irreversibly in,Fa

(AT)

H (n)

hibits the catalatic function in liver. The reaction occurs only in vivo and not with an isolated enzyme. This behavior was explained by Margoliash and Novogrodsky (lll), who noted that the inhibition depends 106a. The spin state of ferricatalase-aside is temperaturedependent, the contribution of a low-spin form increasing with the decreasing temperature (89,107). 107. K. Torii, T. Iizuka, and Y. Ogura, J . Biochem. (Tokyo) 68, 837 (1970). 108. J. L. Hoard, in “Hemes and Hemoproteins” (B. Chance, R. W. Estabrook, and T. Yonetani eds.), p. 9. Academic Press, New York, 1966. 109. R. Countryman, D. M. Collins, and J. L. Hoard, JACS 91, 5166 (1969). 110. W. G. Heim, D. Appleman, and H. T. Pyfram, Science 122, 693 (1955). 111. E. Margoliash and A. Novogrodsky, BJ 68,468 (1958).

7. CATALASE

377

upon the presence of peroxides or compounds susceptible to autoxidation, and proposed a reaction between catalase Compound I, an enzyme-peroxide derivative, and AT, the inhibitor: Catalase

H202

AT

Compound I -+ inhibited enzyme

(3)

Only the subunits with intact prosthetic groups are modified (112) and the inhibited product contains approximately one equivalent of AT per hematin bound at His-74 (112, 113) most probably as in formula (111) (11Sa) :

R (ID)

The derivative retains both the ferric, largely high-spin characteristics and the oligomeric structure of the native enzyme (111, 114). It shows, however, only marginal activity in mediating the decomposition of hydrogen peroxide (115)and is correspondingly inert toward the typical hemoprotein ligands such as cyanide and fluoride. This set of properties is best reconciled by assigning the locus of modification to the distal site of the prosthetic group (116). The mechanism of the inhibitory reaction is uncertain, although two proposals have gained some currency (115-118).One of these invokes “activation” of histidine, which is then subject to an unexpectedly selective nucleophilic attack by AT (or some related compounds) (115,116). The other demands compulsory two-electron equivalent oxidation of A T (117,118), giving an electrophilic intermediate which, prior to its solvation, reacts with the imidazole group of histidine. Although AT is an 112. J. Y. Chang and W. A. Schroeder, ABB 148,505 (1972). 113. B. B. L. Agrawal, E. Margoliash, M. I. Levenberg, R. S. Egan, and M. H. Studier, Fed. Proc., Fed. Amer. SOC. E x p . Biol. 29, 732 (1970). 113a. B. B. Agrawal, E. Margolinsh, M. I. Levenberg, R. S. Egan, and M. H.

Studier, personal communication. 114. P. Nicholls, BBA 59, 414 (1962). 115. E. Margoliash, A. Novogrodsky, and A. Schejter, BJ 74, 339 (1960). 116. E. Margoliash, in “Probes of Structure and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 571. Academic Press, New York, 1971. 117. G. R. Schonbaum, in “Probes of Structure and Function of Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 571. Academic Press, New York, 1971. 118. L. P. Hager, in “Molecular Basis of Electron Transport” (J. Schultz and B. F. Cameron, eds.), p. 367. Academic Press, New York, 1972.

378

GREGORY R. SCHONBAUM AND BRITTON CHANCE

exceedingly poor reductant of Compound I, as are long-chain aliphatic alcohols ( k , 0.6 M-l sec-’ a t pH 7, 2 5 O ) (118a),the latter interpretation seems preferable. This hypothesis does not attribute unique reactivities to AT or histidine but suggests that the efficiency ( 1 1 8 ~ of ) the inhibitory reaction is governed by the rate of diffusion of the oxidized AT from the active site, by the rate of its solvation, and by the proximity of histidine to the site of AT oxidation (118d). Such a scheme lends itself to several alternative descriptions of the oxidative reaction (11’7, 118). However, since the AT-Compound I reaction is pH-invariant (116),the pK, of distal histidine could be “atypical” or, more likely, its modification is not rate determining in the reaction sequence of Eq. (4).It is uncertain, however, whether k , or k, represents the slow step of the reaction. Kinetic or analytical demonstration of a Compound I-AT complex is also lacking. Thus, under nonturnover conditions using preformed Compound I, the redox reactions are first order in Compound I and AT when [AT] 570 mM. N

Compound I

-

Catalase- AT (inhibited enzyme)

118a. k, = knbs - L,,/(AT) where k a b s is the observed first-order constant, k., is the rate constant characterizing spontaneous decomposition of Compound I (-3.5 & 0.5 x lo-* sec-’) and AT, the total concentration of 3-aminotriaeole such was determined under nonturnover conditions using prethat t,p 5 40 sec; k, formed Compound I, thus minimizing possible artifacts resulting from competing or side reactions (118b). 118b. R. P. White and G. R. Schonbaum, unpublished observations. 118c. For mammalian catalase, the efficiency of the modification-defined in terms of molarities, as 100 x modified catalase/Compound I reduced-is of the order of 25 2 5% (118b). 118d. Put in those terms, the AT-dependent modifications are not uniquely “catalase-specific” but should also occur with other proteins in oxidatively coupled reactions. Indeed, this has been recently demonstrated with several peroxidases (119, 110).

119. J. Y. Chang and W. A. Schroeder, ABB 156,475 (1973). 120. H. Snyder and J. Schultz, Abstr. Znt. Congr. Biochem., 9th, 1973 p. 79 (1973).

7.

379

CATALASE

Similar pathways are equally plausible in other reactions leading to the inhibition of catalase ; for example, interaction between Compound I and semicarbazide would give nitrogen and an iV-carbamyl derivative, as in Eq. (5) : Catalase Compound I

I

H

Q

o+c,NHa I

semicarbazide

LQ

=

(5)

+N,

The proposal outlined in Eq. ( 5 ) has not been examined, but independent arguments support it. First acylhydrazides (RCONHNH,) are readily oxidized ( 1 2 2 ) and-presumably because of the intermediary formation of acyl diimides or acyl diazonium ions, as in Eq. (6)-the reaction products are powerful acylating agents. Thus, amines are effectively converted to the corresponding acyl derivatives (122) [cf. Eq. (6) ] :

-[

R‘CON=NH

R’CONHNH,

oxidation

]

RNH,

~

R’CONHR

+

N,

(6)

RTON:

Similarly, Compound I-mediated oxidation of semicarbazide should give oxidation product (s) equally reactive toward a proximal nucleophile. Second, under nonturnover conditions, with preformed compound I, the oxidation of semicarbazide is accompanied by uptake of oxygen (118b), suggesting the formation of an intermediate such as diimide (123, 124), followed by its autoxidation [ cf. Eq. (6) 1 . I n a parallel reaction with AT, no uptake of oxygen is to be expected, and none is obtained (118b). The above interpretations are hypothetical, but they have led to other inquiries showing that selective modifications of the active site of catalase are not limited to reactions involving Compound I. Inhibition of catalases with cyanogen bromide is an example of such a reaction (125).This modification occurs only with the free enzyme in a pH-invariant reaction (pH 4.5-7.5) ; the inhibition resulting from the incorporation of one C moiety of BrCN into the apoenzyme of each subunit (Table IV) (63, 90, 121. C. G. Overberger, J.-P. Anselme, and J . G. Lombardino, in “Organic Compounds with Nitrogen-Nitrogen Bonds,” Chapter 6. Ronald Press, New York, 1966. 122. Y. Wolman, P. M. Gallop, A. Patchornik, and A. Berger, JACS 84, 1889 (1962). 123. P. C. Huang and E. M. Kosower, JACS 89, 3910 (1967). 124. P. C. Huang and E. M. Kosower, JACS 89,3911 (1967). 125. G. R. Schonbaum and F. L. Jajcray, in “Hemes and Hemoproteins” (B. Chance, R. W. Estabrook, and T. Yonetani, eds.), p. 327. Academic Press, New York, 1966.

380

GREGORY R. SCHONBAUM AND BRITTON CHANCE

TABLE I V TITRATION O F CATALASE” WITH BrCN A N D THE CATALATIC ACTIVITYOF THE RESULTING DERIVATIVES

BrCN*/ catalase

% Ce incorporation into the apoprotein

0.3 0.6 0.8 1.0 2 12

% Inhibited enzymed 0 28 52 77 33 >99 >99

95

10-8 (M-1

x kl’ sec-l)*

9.0 6.4 4.1 2.2 0.6

% Activity

0.05

100 71 46 25 7 0.6

0.01

<0.1

aUsing enzyme in which approximately 14 sulfhydryl groups were modified by controlled oxidation. * Per hematin; in 5 mM phosphate, pH 7, 25”. c Assayed, following extraction of hemin via Teale’s procedure (126). d Estimated spectrophotometrically (independently of activity assays). Note: (a) Catalase remains fully active when devoid of SH group [see also Morikofer-Zwez et al. (SS)]and (b) there is a close correspondence between loss of activity and degree of inhibition, suggesting the absence of any subunit cooperativities [Jajczay (go)].

166).The stoichiometry of the reaction is given in Eq. (7) : YH

+ BrCN -+

“YCN”

+ Br- + H+

(7)

where YH represents an apoprotein functional group and YCN is the simThe identity plest, but not necessarily the final, reaction product (166~). of Y remains unknown. Almost certainly it is not an -SH, a- or c-amino, thioether, or an -OH group ( 9 0 ) . There are striking similarities in the physical [Fig. 2 (166)and Fig. 3 (126b)l and chemical properties of the cyanylated enzyme (Table V) and the aminotriazole-catalase derivative. Both retain the structural integrity and ferric oxidation state of the native enzyme. In addition, as gauged by their optical spectra [Fig. 2, but see also Figs. 6A and 6B of Margoliash and Novogrodsky (111)I, the modification of the apopro126. F. J. W. Teale, BBA 35, 543 (1959).

0

b

126a. Hydrolysis of YCN to Y- NH1 or an intramolecular cross-linking with a residue X H to Y-C-X would also accord with the analytical data.

II

NH 126b. G. R. Schonbaum, J. Peisach, and W. E. Blumberg, unpublished observations.

7.

381

CATALASE

1

120.0

I

80.01

A4

8 0-

\\

H 60-

4 0-

Y

20.0

20-

0

FIQ.2. Spectrum of (A) horse erythrocyte catalase and (B)its cyanogen bromide derivative in 0.01 M phosphate pH 7.15, 25". Extinction coefficients (M-' cm-') are expressed in terms of heme-Fe (126).

tein elicits in both cases nearly identical heme-linked perturbations which correlate with the loss of activity and ligand binding capacity. It is not unlikely, therefore, that YH = His. With cyanogen bromide, histidine

Mognetic field

FIG.3. Electron paramagnetic spectra in the region of g = 6, 1.4"K for (A) horse erythrocyte catalase and (B) its cyanogen bromide derivative. Note that the rhombicity of the catalase is greater than that of the inhibited enzyme (126b).

W M h3

TABLE V COMP.4RISON

CHEMICAL PROPERTIES O F HORSEERYTHROCYTE CATALASE ITSCYANOGEN BROMIDE DERIVATIVES

O F PHYSICAL A N D

Property Molecular weight 1013 szOnw Absorptivities X,, (nm) 1 O - a ~ (M-1 cm-1)

Native

Ref.

246,000 (*8,000) 11.2 (50.05)

(39)

(39)

277 406 505 538 625 -880 (89, 90) 90 115 11.4 9 . 8 8 . 1 1.1

AND

Modified

Ref.

-250 ,000 11.2 ( & 0.05)

(90) (90)

277 402 506 533 633 -880 (125) 90 102 12.1 11.2 7.0 0.9 0

Circular dichroism in UV A (nm) 10-3 (deg cm*/decimole) Magnetic susceptibility (Bohr magnetons) Oxidation state Reducibility by NazSz04 Affinity for HCN a t pH 7, K ( M )

Catalatic activity’ 10-6kl’ (M-1 sec-1) (Kat.f.) a

190 200 81 4

210 -56

220 -47

5.96

(90)

(30)

x

(90)

11

220 -47

(90)

Nonreducible

2 (12.5)

F Kn

0

(1265)

No evidence of complex when (126) [enzyme] 5 10 p M and [HCN] 5 100 p M 0.06 rf: 0.002b

g 0

Ferric; high spin

Nonreducible 10-6

210 -56 -

(107)

Ferric ; high spin

-5

190 200 72 2 . 5

2

%

‘1 9

1:

U

(90) cj

(95,000)

Per heme group. Possibly resulting from some residual free enzyme.

(-500

5 200)

-

383

7. CATALASE TABLE VI REGENERATION OF ACTIVECATALASE FROM Ligand AFormate Formate Formate Acetate Acetate Fluoride

THE

+

PH

CYANYL.4TED ENZYME, 25'

108 [AAH] (MI

los [AH] (MI

100 100 200 -5 200 95

N O . 06

7.0 4.5 4.5 4.5

4.5 4.5

15 30 -2.8 110 4

lo6 k sec-1)

(M-1

G

15 f 3 15 f 3 -0.6 38 f 2

No significant regeneration within 100 hr [Jajczay (go)].

would give either an N-cyanoimidazole, as in formula (IV) (127) or its solvolysis product, N-carbamyl imidazole as in formula (V) (128) :

H

NCN/ 'NH,

N/-N/CN

LJ

LJ

(N)

(V)

Both (IV) and (V) are stable in nonpolar solvents but not in acidic or basic solutions (127, 128). Similarly, the integrity of YCN is retained, but only in the structurttlly intact protein, in the absence of potential enzyme ligands (formic, acetic, or hydrofluoric acids). Such ligands promote a slow hydrolysis of YCN (Table VI) with a concomitant recovery of full enzymic activity (90). Furthermore, acid denaturation of the cyanylated enzyme at pH 2 is followed by complete hydrolysis of YCN (half-time - 4 hr a t pH 2, 2 5 O ) ( g o ) , and the formation of carbon dioxide, possibly via Eqs. (8) and (9) : YCN HCNO

n

HCNO (+YH2+)

(8)

CO, (+NH,+)

(9)

The characteristics of the BrCN-catalase reaction indicate, therefore, that activation of distal histidine-its conversion into a n electrophilic reagent ( 1 16)-is not necessarily a prerequisite to inhibition. Nevertheless, the remarkable selrctivity of this reaction points to some syfiergic facets of the BrCN-enzyme interaction which are unique to the distal 127. H. Giesenian, J. P m t k . Chem. r41 1, 345 (1955). 128. G. R. Stark, Biochemistry 4, 588 (1965).

384

GREGORY R. SCHONBAUM AND BRITTON CHANCM

region (168a). Such interactions may arise through mutual polarization of the YH-BrCN system and its modulation by iron of the prosthetic group (a Lewis acid) or by hydrogen bonding. These assumptions are central to the hypotheses outlined in Eq. (9a), where the constellation near the ferric ion (17)is represented as in (VI) .

Pr -Fe a:

1

H

Pr-Fe--O(

I

I

k.

H

Y

C

N

Scheme A brings into focus the well-known Lewis acid catalysis, of which the immediately pertinent example is the reaction of cyanogen chloride with benzene under Friedel-Crafts conditions (130-156):

+ AlCls-

q5-CN + HC1 + AlCh (10) Alternatively, as in Scheme B, BrCN forms only an outer sphere complex, with Fe(H,O) assuming the role of an acid catalyst. I n both schemes the essential feature is a polarization of the YH-BrCN system in which YH acts as a donor and BrCN as an acceptor, not unlike the situation in molecular complexes of halogens with u and T donors (133, 134). ClCN

bensene

[NC----ClAlClsl

128a. A similar reaction occurs with sperm whale metmyoglobin (1291, most likely at E 7 (distal) histidine. 129. F. L. Jajcsay and G. R. Schonbaum, Proc. Can. Biochem. SOC. 12, 73 (1969). 130. G. Olah, “Friedel-Crafts and Related Reactions,]’ Vol. I. Wiley (Intersciepce), New York, 1963. 131. S. Nilsson, Actu Cheni. Scund. 27, 329 (1973). 132. A. A. Woolf, J . Chem. SOC.London p. 252 (1954). 133. R. Foster, “Organic Charge-Transfer Complexes.” Academic Press, New York, 1969. 134. L. J. Andrews and R. M. Keefer, Advan. Inorg. Chem. Radiochem. 3, 91 (1961).

7.

385

CATALASE

The convergence of interactions, particularly as in Scheme B, may well explain the specificity of the cyanogen bromide-dependent inhibition. Furthermore, as both schemes imply, the reaction is pH-invariant; it occurs only with the native enzyme; it is competitively inhibited, e.g., by cyanide or formate (126) ; and the resulting product (YCN) like catalase itself, should-and does-exchange water or protons a t the sixth coordination site (1OOa).This, as well as weak acid-dependent reactivations (Table V ) , suggests that Y H is converted to YCN a t a functionally important residue. We propose, therefore, that YH acts as a general acidbase catalyst in both redox and ligand exchange reactions.

C. LIGAND EXCHANGE REACTIONS Catalase reacts reversibly with some weak acids forming spectroscopically and magnetically distinct noncovalent derivatives. Of these, catalase-cyanide, -azide, -fluoride, -formate, and -acetate complexes have been extensively studied (37, 135, 136) and reviewed in some detail (16-18). Briefly, there is a consensus that such reactions do not involve heme-heme interaction ; and, with the possible exception of carboxylate ligands ( l o g ) , all presumably result in replacement of the proximal “aquo” ligand a t L, in a stoichiometric reaction shown in Eq. (11) : Pr-Fe(H20)

+ AH + Pr-Fe(AH) + H2O

(11)

where P r indicates protein and AH, a weak acid ligand. All reactions are of the second order and are independent of the ionization state of the enzyme in the p H range from 4.0 to 9.0. All entail, in the rate-limiting step, an interaction of catalase with the undissociated form of the ligand (Fig. 4) but occur without a concomitant net uptake or loss of protons. The above parameters define the stoichiometric mechanism of the reaction but have little bearing on its intimate character (137); thus, the participation of the weak acid, AH, rather than its conjugate base, A-, in the rate-limiting step of the reaction does not necessarily imply that AH is ligated in the final product. A couple of examples illustrate the issue. Consider first the catalasec,yanide system characterized by the affinity, K , 2 X lo5 M-*. There is no doubt that H C N is the reacting entity ($7, 69) and that net proton release or uptake does not occur (118b). Nevertheless, FeCN appears

-

135. K. Agner and H. Theorell, ABB 10, 321 (1946). 136. M. L. Kremer, Zsr. J. Chem. 8,799 (1970). 137. C. H. Langford and H. B. Gray, in “Ligand Substitution Processes,” p. 2. Benjamin, New York, 1965.

386

GREGORY R. SCHONBAUM AND BRITTON CHANCE 6-

I:

%3

;\

Qp”i& fluoride

I

Acetate

1

1

1

.

1

I

I

I

to be the only possible product. To emphasize this point, note that CH,C = N, CH,N = C, or H C E CH do not form viable complexes-a behavior not attributable to peculiar steric effects but reflecting their intrinsic reactivity, or lack of it. Similarly, the high affinity of “formic acid” ( K , 2 X lo5 M-l) contrasts with the virtual inertness of formamide ( K , 2 0.5 M-I), suggesting again that dissociation of the ligand (AH 4A- H+) is a prerequisite of its effective ligation. Such reactions may be expressed as in Eq. (12) : H

+

It is therefore understandable that the aminotriazole- or BrCN-modified enzyme no longer participates in ligand interchange reactions, particularly if, as implied, YH acts as a general acid-base catalyst. That HA rather than A- is the reactive species may therefore indicate that the distal site, embedded in a hydrophobic environment, is largely inaccessible to anions and cations. The reaction outlined in Eq. (12) is undoubtedly more complex than indicated and may proceed via dissociative, associative, or interchange mechanisms (137).In a t least one case, however, it was shown that direct ligand replacements, via an associative pathway as in Eq. (13), do not occur: Fe(HA)

+

A’H

+

“A’H

+Fe(A’H) + AH

(13)

7. CATALASE

387

where A’H is an entering ligand other than water. For example, the replacement of thiocyanate by cyanide does not proceed via the SN2 (lim) associative mechanism (118b) since the rate of catalase-thiocyanate dissociation (/cap* 2 x sec-’, pH 6.7, 2 5 O ) is essentially the same in the presence and absence of cyanide. Accordingly, the reaction may be governed by dissociation of thiocyanate, or alternatively, the ligand interchange occurs in two steps: the first, involving replacement of thiocyanate by water, being rate-limiting [Eq. (14a) ] : H

Pr-Fe-

(frCS)

+

+

H,O w Pr-Fe-OH

1 YA,

L

Y

H

(HSCN)

(14%)

,

I n the second, fast step, the water molecule is replaced by the entering cyanide as in Eq. (14b) :

+

Pr-Fe-OH L

Y

H

HCN

+Pr-Fe-7N I

,

Y

H

+

H,O

( 14b)

,

Whatever the mechanism, “YH” would facilitate the reaction by participating in proton transfers between the entering and departing ligands, perhaps as shown in Eq. (14c) : Pr-Fe-0.

+

/H

HCN

+Pr-Fe-OH, (144

+ Pr-Fe-CN

(H,O) + P r - F e - C p JH,

+

H,O

&.Iz

Particularly intriguing observations by Ehrenberg and Estabrook (158) point to the differences in catalase-ligand interactions in solution and in the frozen state. In liquid phase, catalase does not effectively ligate ammonia (158) although spectroscopic changes are discernible (SO). However, in the frozen state, low-spin catalase-amine complexes are greatly stabilized (1S9), possibly as a result of a heme-linked conformational change. Analogous conclusions were drawn by Yoshida at al. (100) using frozen glycine-catalase solutions, initially a t pH 2 8, in which the enzyme under138. A. Ehrenberg and R. W. Estabrook, Acta Chem. Scand. 20, 1667 (1966). 139. G. Heimberger and A. Ehrenberg, in “Probes of Structure and Function of

Macromolecules and Membranes” (B. Chance, T. Yonetani, and A. S. Mildvan, eds.), Vol. 2, p. 561. Academic Press, New York, 1971.

388

GREGORY R. SCHONBAUM AND BRITTON CHANCE

goes a transition from a high-spin to a low-spin form with decreasing temperatures. They attributed this to a thermal equilibrium of high- and low-spin aquo and hydroxo states of the enzyme, rather than to glycinedependent effects. A reappraisal of these analyses seems warranted, however, since Ehrenberg failed to observe a low-spin form of catalase at 77OK (initially a t pH 10) and because the optical and EPR spectra of the catalase amine complexes and of the low-spin aquo or hydroxo components are nearly identical. N

IV. Catalase-Mediated Redox Reactions

Cellular control of peroxides is one of the key biochemical reactions, and a process in which catalase plays a significant role. Catalase is not only an exceptional mediator of hydrogen peroxide decomposition (140) as in Eq. (15)

but is also an effective catalyst (141-149) of peroxide-dependent oxidations of hydrazoic, formic, and nitrous acids; of lower aliphatic alcohols; and of hydroxylamine. Except for the oxidation of nitrous and hydrazoic acids such reactions may be represented by a stoichiometric relationship given in Eq. (16). ROOH

-

+ HXOH catalase XO + ROH + HoO

[R = H, alkyl, acyl; X = 0, NH, C = O , (CH,).-1,2,s]. As shown by Chance et a2. ( I @ ) , these reactions occur in a t least two stages, outlined in Eqs. (17) and (18). k + ROOH 4 Compound I Compound I + HXOH 2 catalase + XO

Catalase

(17) (18) Both steps in the catalytic cycle are pH-invariant (36) when the substrates are un-ionized. Except a t very high concentrations of hydrogen peroxide (144, 146) both follow second-order kinetics (146). 140. 141. 142. 143. 144. 145. 146.

B. Chance, D. S.Greenstein, and F. J. W. Roughton, ABB 37, 301 (1952). D. Keilin and E. F. Hartree, BJ 39, 293 (1945). L. A. Heppel and U. T. Porterfield, JBC 178,549 (1949). H. Theorell and A. Ehrenberg, ABB 41,462 (1952). Y . Ogura, ABB 57, 288 (1955). P. Jones and A. Suggett, BJ 110, 617 (1968) B. Chance, JBC 182,649 (1950). I

7.

CATALASE

389

These aspects of catalase-mediated reactions, comprehensively reviewed elsewhere (16-18, 147, 148), highlight two topics of immediate interest: the nature of Compound I and its peroxidatic reactions.

A. THENATUREOF COMPOUND I The nature of the oxygen and peroxide compounds of catalases, peroxidases, and, more recently, hemoglobin was questioned by Philip George, who proposed (9) that electrons were transferred between the ligand and the metal atom in the formation of such “compounds,” in contrast to the more general term, “complex.” Evidence for the generality of this designation has developed slowly over the years to the point where not only can complete electron transfer, as considered by George, be demonstrated but also more recently electron delocalization in a variety of lesser degrees has been shown [ (149) cf. also Caughey et al., Chapter 51. This phenomenon has become especially prominent in the discussion of the oxygen compounds of hemoglobin, where the designations range from “covalently bound oxygen with an extensively delocalized electron” to “loosely bound oxygen with only slight electron delocalization.” These most interesting and significant approaches reiterate George’s query on the point of nomenclature of “compounds” and “complexes”: What is the proper term for an intermediate compound in which modest electron delocalization has occurred as in oxyhemoglobin? This problem has been raised most recently in the discovery of a series of cytochrome oxidaseoxygen compounds (150) in which a plethora of electron transfer possibilities are available, leading to what Greenwood et al. (151) have termed “mixed valency states” of the respiratory chain, with varying degrees of electron delocalization or electron transfer to oxygen (150). It seems appropriate, therefore, to reserve the term “complex” for the first dissociable products of the combination of enzyme and substrate-the “enzyme-substrate complex”-which, according to Michaelis and Menten can be reversibly dissociated without alteration. Examples of this would be afforded by the above-mentioned cytochrome oxidase-oxygen compound and, indeed, oxyhemoglobin compounds which generally fulfill this 147. S. B. Brown, P. Jones, and A. Suggett, in “Inorganic Reaction Mechanisms” 13, p. 159. Wiley (Interscience), New York, 1970. 148. B. Chance, Advan. Enzymol. 12, 153 (1951). 149. W. S. Caughey, C. H. Barlow, J. C. Maxwell, J. A. Volpe, and W. J. Wallace, Ann. N . Y . Acad. Sci. 244, 1 (1975). 150. B. Chance, C. Saronio, and J. S. Leigh, Jr., Proc. N a t . Acad. Sci. U. S . 72, 1635 (1975). 151. C. Greenwood, M. T. Wilson, and M. Brunori, BJ 137,205 (1974).

(J. 0. Edwards, ed.), Vol.

390

GREGORY R. SCHONBAUM AND BRITTON CHANCE

criterion. The chemical equation for an intermediate complex therefore involves the reversible sequence: S

+ E+ES+S + E

(19)

This sequence is apparently applicable to the cytochrome oxidase-oxygen enayme-substrate compound, where the apparent dissociation constant is relatively large (0.5 mM a t -1200) (150). At higher temperatures, however, compounds are formed in which the reversibility can no longer be identified in spite of a number of efforts to do so (152, 169). I n studies of catalase, much effort has been directed toward a determination of whether or not hydrogen peroxide could be dissociated from the enayme-substrate intermediates of catalases and peroxidases. It should be pointed out that catalase, as contrasted with cytochrome oxidase, has been studied only a t room temperature, and if any lesson is to be learned from the study of cytochrome oxidase (150), it is that the “complexes” are most likely to be identified a t low temperatures, as precursors of the “compounds.” I n this sense, they are of first importance and not to be ignored in our understanding of the mechanism of enzymic reactions. B. THE CATALASE REACTION MECHANISM By analogy to other catalase-ligand interactions, Compound I was initially taken to be an enayme-peroxide complex, even if a rather unusual one. However, the optical spectrum of Compound I (154, 155), its magnetic susceptibility (156), and the lack of a discernible E P R signature (126b) are unlike those of other high- or low-spin catalase-ligand derivatives. In spite of these anomalies, the idea of a “complex” remained a working hypothesis, particularly since kinetic studies failed to reveal any intermediate (9) preceding Compound I formation (144, 157) and since the interaction of bacterial catalase with methyl hydrogen peroxide could be expressed in terms of an equilibrium reaction (158).The latter proposition 152. F. A. Schindler, Ph.D. Thesis, University of Pennsylvania, Philadelphia, 1964. 153. S. Muraoka and E. C. Slater, BBA 180,227 (1969). 154. B. Chance, ABB 41, 404 (1952). 156. A. 5.Brill and R. J. P. Williams, BJ 78,253 (1961). 156. A. S. Brill, Free Radicals Biol. Syst., Proc. Sump., 1960 p. 53 (1961). 157. B. Chance, in “Currents in Biochemical Research” (D. E. Green, ed.), p. 308. Wiley (Interscience), New York, 1956. 158. B. Chance and G. R. Schonbaum, JBC 237,1962 (1962).

7.

CATALASE

jibh

391

.14

152 .SO

100

200 Sacondr

3

(A1

Fra. 5. (A) Formation of Compound I using EtOOH. (B) Decomposition of EtOOH ; and in the absence of enzyme ( W). I n (A) and (B) : in the presence of catalase (0) M . lysodeikticus catalase (hematin) N 6.2 p M ; (EtOOH) 19.5 p M , pH 8, 25" ( 1 0 1 ~ ) . AAW (maximum at 300 sec) Zl.8 p M Compound I. Concurrently, less than 0.15 r M Compound I1 was formed, as determined independently.

was based on the assumption that Compound I is adequately stable and that it is the sole product of the enzyme-peroxide reaction-a point considered in detail in that publication. More recent data suggest that acetaldehyde is formed from ethyl hydrogen peroxide in the presence of M . lysodeikticzls catalase (Fig. 5 ) . Similar results are obtained with bovine liver and horse erythrocyte catalases (159, 159u), the major fraction of ethyl hydrogen peroxide being converted into acetaldehyde ( 1 7 ) . It appears that the equilibrium of catalase and peroxide involves problems similar to those of cytochrome oxidase (150) in that electron transfer between enzyme and substrate occurs and that low temperature and rapid reaction techniques may be required to study this equilibrium. The formation of aldehyde appears to be an intrinsic property of the catalaseethyl hydrogen peroxide system, as noted in the early work of Stern (160); it is not an artifact attributable to the presence of some adventitious ethanol. Even in the presence of methanol or nitrite (both excellent reductants of Compound I ) , approximately 60% 4 10% of EtOOH is converted to acetaldehyde (Table V I I ) . Moreover, the extent of conversion is nearly invariant of the donor concentrations. Both facets of the reaction suggest that the enzyme mediates either dehydration of EtOOH (Scheme 1) or a redox rearrangement (Scheme 11). 159. G. R. Schonbaum, Abstr., Wenner-Gren Symp. Struct. Funct. Oxidatiori-Reduclion Enzymes, 1970, p. 48 (1970). 159a. G . R. Schonbaum, Abstr., Int. Congr. Biochem., Sth, 1979 p. 49 (2b10) (1973). 160. K. G. Stern. JBC 114,473 (1936).

392

GREGORY R. SCHONBAUM AND BRITTON CHANCE

TABLE VII FORMATION OF ACETALDEHYDE FROM ETHYLHYDROQEN PICROXIDE MEDIATED BY M. lg8OdeikliCUS (MLC) AND HORSEERYTHROCYTE (HEC) CATALASESO Acetaldehyde from Donor concn. range (mM) Ethanolb 1.64.1 Methanol 2 .O-8.2 Nitrite 0.3-1 .3

MLC

HEC

pM

%of (EtO0H)i

pM

%of (EtO0H)i

51 f 1

100

51 f 1

100

36 It 1

70

32 f 1

64

37 & 2

72

27 f 2

52

(MLC)re 3.2 p M , (HEC) 0.76 p M , (EtOOH) 51 p M in 0.1 M phosphate, pH 7, 25". Control experiments showing conservation of redox equivalents and not the origin of aldehyde [Schonbaum (10Ia)l. E

f

+

RCHO

+

H,O

(Catalase)

EO

+

RCH,OH

(Compound I)

SCHEME I E

+

RCH,OOH

+(E .RCH,OOH)

(EO .RCH,OH)

Y k 2 E

+

RCHO

SCHEME

+

H,O

\ EO +-RCKOH

Ir

Scheme I1 is preferred because with methyl or butyl hydroperoxides

k, > k, (IOIa).Essentially, then, Compound I is not the primary enzyme-substrate complex (161). The formation of Compound I entails the reduction of substrate (peroxide) a t the active site (compare Schemes I and 11). The recent discovery that nearly one mole of Compound I is formed in the 1 :1 reaction between catalase ferriheme and peracetic 161. P. George, ABB 45,21 (1953).

7.

393

CATALASE 25pM CH,CO,H

1 $ 4.25pMCH,COSH

I,

8.5pM CH,CO,H

,i \

4 -

I

I

0

10 10' [CH,CO,

20

30

H] (M)

FIG.6. Titration of -15 fiM horse erythrocyte catalase-ferriheme with peracetic acid, 0.08 M phosphate, pH 7.23, 25" ( 1 0 1 ~ ) .

acid (Fig. 6) further supports this conclusion ( 1 6 1 ~also ; see 169, 166, 163). As in enzyme-hydrogen peroxide reactions, the unionized peracid is the immediate substrate (162) and, according to Jones and Middlemiss (163),one mole of acetic acid is released per mole of catalase ferriheme converted to Compound I. No information is available as to the individual steps of the reaction, and none is likely to issue from kinetic studies with higher alkyl peroxides (C 2 2) or peracids. The reasons are implicit in the data of Table VIII (1,69, 101a, 162, 164), which show that the observed rate with peracetic acid, hyconstants for Compound I formation ( k ,),, droxymethyl hydroperoxide, and ethyl hydrogen peroxide are of the same order of magnitude (2.5 0.5 X lo4 M-* sec-') in spite of differences in the following:

*

0-0

dissociation energies for EtOOH (43 kcal/mole) and peracids kcal/mole) , the basicities of the leaving groups (pK,:acetic acid, 4.76; ethanol, 18), and the ionization properties of peroxides (pK, :peracetic acid, 8.2; EtOOH, 11.8) (-35

161a. The stoichiometry of the reaction is unaccountably variable, the average of several titrations showing that approximately 0.87 & 0.08 mole of Compound I is formed per mole of peracetic acid. 162. P. Jones and D. N. Middlemiss, BJ 130,411 (1972). 163. P. Jones and D. N. Middlemiss, BJ 143,473 (1974). 164. S. Marklund, BBA 289,269 (1972).

394 GREGORY R. SCHONBAUM AND BRITTON CHANCE

e:

7.

395

CATALASE

We must assume, then, that the rate-limiting step is virtually independent of the above parameters. Nor can it be involved in the redox-dependent rearrangement of iron-protoporphyrin since the reactions with H,O, and CH,OOH are faster than those with EtOOH or CH,CO,H. A relationship between k, ap,, and van der Waals volumes of peroxides is the only discernible pattern. This strongly suggests that enzyme-peroxide association is the rate-determining step and, as such, is rather irrelevant to the elucidation of the redox transformations. The cocatalytic role of the apoprotein in facilitating Compound I generation is increasingly regarded as that of a general acid-general base (17, 75,159, 159a), although different functional groups are seen as fulfilling this role. Jones and Suggett ('?5),pointed to the possible involvement of >C(NHr),+ group from an arginine residue (BH') and a carboxylate (A-) as in Eq. (20). H ,

H

+

Hgoz

B H ,F

H,O-Fe

l

I

,

?I

H,

(20)

O/O--Fe

I

H

9_ _ _ _ 0_ _ _ _ _ _ Fe I

H

,

As already mentioned, a less detailed scheme may also be developed, using only Y H 3 His [formulas (VII) and (VIII) ]

' 7 p-".

Enzyme

3,

,'

Enz-yy

2

H, -

R

or

I

H.,,

Y'

"0

3

H (VE)

(VIII)

The central theme that the apoprotein facilitates the scission of the 0-0 bond is based on the established mechanisms of peroxide heterolysis (165).By invoking "concerted" proton transfer (s) in the transition state, such schemes illustrate that oxygen-oxygen heterolysis need not be attended by an electrostatically unfavorable charge separation. I n addition, they offer some rationale for the observed high entropy of activation in 25 cal mole-' deg-I) (166). the primary H,O,-catalase reaction (-AS* This should be the case in a rigid lattice of interactions implied in Eq. (20) and formulas (VII) and (VIII). All such suggestions are clearly most tentative since the nature of the oxidation product, Compound I, is still unresolved. The stoichiometry of the enzyme-peroxide reaction merely demands that its formal oxidaN

165. L. Bateman and K. R. Hargrave, Proc. Roy. Soc., Ser. A 224,339 (1954). 166. G. K. Strother and E. Ackerman, BBA 47,317 (1960).

396

GREGORY R. SCHONBAUM AND BRITTON CHANCE

tion state be Fe(V), i.e., two oxidation equivalents above the native enzyme, Fe (111).Such an assignment tallies well with the magnetic susceptibility for a compound with three unpaired electrons (6000-6500 X lo-’’ emu) (166) but does not uniquely define their distribution. For example, assuming that the oxidation is confined to the prosthetic group, other structures compatible with the magnetic susceptibility data can be expressed as a radical combined with Fe(1V) or as a diradical in conjunction with low-spin Fe (111).None of these structures singularly reflects the nature of Compound I but, as emphasized by Hamilton (167),all in varying degrees could contribute to its resonance form, for the term “oxidation state” has little chemical significance in compounds with a substantial covalent character, as evident in coordination compounds with delocalized ground states such as metal dithienes (168, 169) and metal-nitric oxide complexes (170). In Compound I, such polarization interactions should involve an extensive delocalization of electrons from the porphyrin toward the metal ion; an extreme case of which is a porphyrin-r-cation radical combined with Fe(1V) as shown in Eq. (21),

where X.6+denotes a radical moiety. Pertinent to this discussion are recent studies on porphyrin-r-cation radicals derived from magnesium and cobaltic oct.aethy1 porphyrins (171, 172),and zinc and magnesium tetraphenyl porphyrins (171).In all cases, the optical spectra of such radicals share features also found in Compound I (Fig. 7). These are (a) a decrease of r-r” transitions associated with the Soret band and (b) the appearance of bands between 600 and 700 nm. The chief objection to the proposal implicating “a free radical” moiety stems from the absence of a distinct E P R signature for Compound I. This is not an overriding restriction since, conceivably, an electron localized on the porphyrin will couple through exchange interactions with spin localized on the metal (173),resulting in broadening of the EPR signal beyond the limit of detection. 167. G. A. Hamilton, Advnn. Enzymol. 32,55 (1969). 168. G. N. Schrauzer, Accounts Chem. Res. 2,72 (1969). 169. J. A. McCleverty, Progr. Znorg. Chem. 10, 49 (1968). 170. J. A. Lewis, Sci. Progr. (London) 46,506 (1959). 171. J. H. Fuhrhop and D. Mauzerall, JACS 90,3875 (1968). 172. R. H. Felton, D. Dolphin, D. C. Borg, and J. Fajer, JACS 91, 196 (1969). 173. D. Dolphin, A. Forman, D. C. Borg, J. Fajer, and R. H. Felton, Proc. Nut. Acud. Sci. U.S. 68, 614 (1971).

397 120’ 100-

7 80E

-

-

‘I60-

5 40: 20OJ

400

350

450

500

550

600

I

700

650

‘ 0

A (nm)

FIG.7. Spectrum of horse erythrocyte catalase (---) and its peracetic acid deriva(Compound I) ; 0.08 M phosphate, pH 7.23, 25” ( 1 0 1 ~ ) . tive (-)

Among other factors contributing to electron delocalizations, the influence of environment, particularly that of the trans ligands, must also be taken into account. Ideally, such ligands should be polarizable but not readily oxidizable. An imidazole group a t L, could fulfill such a function and, as L,, the oxygen anion would be appropriate because of its ability to enter into u-type coordination and x-donor bonding. The nature of Compound I being still unresolved, it is not surprising that the mechanisms of its reduction have been variously expressed as: transfer of oxygen from Compound I (EO) to the reductant (HXOH) followed by the rearrangement of the “hydroxylated” intermediate (118) [Eq.

hydride transfer (17,18,75,174,175) [Eq. (2311 EO

+

XHOH

+

c .I EO

HiCQ-,,

E(H,O)

+

XO

(23)

outer sphere electron exchange (17.9) inner sphere electron transfer (17,159~4167) [Eq. (24)] EO

+

HXOH

[ -

+

E+-X

(/]

p---H

E(H,O)

+

XO

(24)

174. L. L. Ingraham, “Biochemical Mechanisms,” p. 71. Wiley, New York, 1962. 175. P. Nicholls, BJ 00, 331 (1964).

398

GREGORY R. SCHONBAUM AND BRITTON CHANCE 17mM CH,OOH

I.3pM CH,OOH

421-480nm

FIG.8. Reduction of horse erythrocyte catalase Compound I by methyl hydrogen peroxide; at pH 7, 4". Note also the ready oxidation of Compound I1 to Compound I11 by H201where knpp 3 x 1oJ M-'sec-' (164).

-

The reactions outlined in Eqs.. (22)-(24) are widely recognized in "model" redox systems. Their relevance to the catalase-mediated oxidations is the central issue-the subject of a lively current debate. I n this context, the following criteria and guidelines must be considered : 1. The HXOH donors (hydrogen peroxide, hydroxylamine, formic acid, and alcohols) and nitrous acid reduce Compound I to ferricatalase without detectable participation of Compound I1 (176). Accordingly, the reaction occurs either via two-electron equivalent reductions, or the rate of Compound I1 formation is smaller than the rate of its reduction (17'7, 178) [Eq. (25)]. Compound I + HXOH +[Compound I1 XOH] +catalase + XO (25) slow fast

If so, then Compound I1 should be reducible by radicals derived from HXOH (e.g., HOz' and NO,'). Contrary to these expectations, Nicholls (1'79) failed t o adduce evidence indicating reduction of Compound I1 by NO,'; and Compound I1 is the dominant derivative in the presence of reagents which generate superoxide. Apparently, as in the corresponding peroxidase reactions (180) catalase Compound I1 is not readily reducible by an externally generated superoxide. These observations, although not entirely conclusive, do bring into question the scheme of Eq. (25) but pertain only to compound I-HXOH interactions. With other substrates, for example, with hydroperoxides (16.4) (Fig. S), phenols (178), or oximes (101a) one-electron equivalent reductions of Compound I do occur. 2. No pH dependence is apparent in EO--(HXOH) reactions, when ionization state of the reductant is taken into account (36)(Fig. 9). All 176. B. Chance and D. Herbert, BJ 4,402 (1950). 177. D. Keilin and P. Nicholls, BBA 29, 302 (1958). 178. P. George, BJ 52, XIX (1952). 179. P. Nicholls, BBA 81, 479 (1963). 180. B. H. Bielski, D. A . Comstock, A. Haber, and P. C. Chan, BBA 350, 113 (1974).

7. CATALASE

399

4

15' 0

tb

6 PH

Fro. 9. Effect of pH upon activity of Compound I toward hydrogen donors (38).

reactions conform t o the rate law

+

(26)

d(EO)/dt = ~ ~ ' ( E O ) ( X T ) / [(Ka/H+)I ~

where XT is the total concentration of the reductant, Ka the ionization constant [Ka = (H+)(HXO-)/HXOH], and k:/[l Ka/H+] = k4 app. Thus, as in the reactions of the resting enzyme with ligands, only the un-ionized donor interacts with Compound I. Yet, following the formation of a n outer sphere complex, EO(HXOH), ionization of the substrate is a likely prerequisite t o HXOH oxidation. The point a t issue is illustrated by the relative reactivities of formic acid and formamide: the former is an excellent donor (k:Tpp 9 X 105 IT-' s-l ) (36) and the latter is inert ( 1 0 1 ~ )Similarly, . N-methyl hydroxylamine is a reductant comparable t o ethanol but 0-methyl hydroxylamine is relatively inactive ( 1 0 1 ~ ) . It is also a much weaker acid.

+

-

Donor k:',",,(M-' sec-1)

CHsCHZOH 1000

CHSNHOH 350

CH30NHz < 10

3. The reactivity pattern ( k 4 a p p : CHaCH20H> CHaNHOH > CH30NH2) would not be expected if either exchangeability of X--H hydrogen or nitrogen nucleophilicity were of importance in governing k d a p p , and the results are not peculiar to methoxy amine. I n general, amines are not effective Compound I reductants (k!c;p <0.2 Ri1-l sec-' a t 25") and unlike in nonenzymic reactions (181-183), attempts to demonstrate hydroxylations of R N H 2 (R = H, CHa) have so far proved unsuccessful (118b). Nor is the failure to effect hydroxylations limited to amines. Thus, reduction of Compound I in the presence of cyanide does not result in the formation of cyanate (118b) and thiols, which are superior to alcohols as reductants of peroxides, are not the preferred donors 181. W. R. Dunstan and E. Goulding, J. Chem. Soc., London 75, 1005 (1899). 182. I(.M. Ibne-Rasa and J. 0. Edwards, JACS 84, 763 (1962). 183. S. N . Lewis, in "Oxidations" (R. L. Augustine, ed.), 1701. I, p. 213. Dekker, New York, 1969.

400

GREGORY R. SCHONBAUM AND BRITTON CHANCE

in catalase-mediated oxidations (146): kis&,

CHaCHnOH

Donor (M-I 8ec-I)

CHaCHnSH

<70

1000

Hence, barring the formation of EO (RSH) complex in a rate-determining step, the nucleophilic attack by sulfur at EO oxygen, or transfer of hydrogen atom from S-H, does not appear to be the preferred reaction pathway, Further, the reduction of Compound I by aaide does not conform to the stoichiometric mechanism [Eq. (27) J (143, 176). EO + HNa +I+E(N0-) + N, + H+ (27) Nitrous oxide, and not nitrogen, is one of the major products (143). The reaction sequence HNa

EO

N=N=N--OH 2 NO-

.--)

Nn

+ NO- + H+

+ HzO

2 €I+

NzO

is thus precluded. Otherwise, Nz should be the dominant product. Only in the coupled oxidation of nitrite using glucose oxidase-glucosel8OZ (as H1sOISOHgenerating system) and beef liver catalase, which results in the formation of isotopically enriched nitrate (183u),are the results compatible with an oxygen atom transfer mechanism [Eqs. (28) and (29) 1. Even in this case, alternative interpretations are possible, such as oxidation of nitrite via outer or inner sphere electron transfers [Eqs. (30) and (31) ] followed by, or concurrent with, fast solvation:

~ 4 8 0

+

HNO,

-L

HBl

6+ 1

E"0 (NO,)

H,O -E(H,O)

+ '%NO,- + 'H

(30)

where B denotes a base. 183a. The degree of '*O incorporation is variable, but contrary to the preliminary report (184) it is not marginal. 184. R. J. Olcott and G . R. Schonbaum, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 33, 1245 (No. 119) (1974).

7.

401

CATALASE

TABLE IX REDUCTION O F COMPOUND 10B Y SATURATED PRIMARY ALCOHOLS Y-CHzOH Y ~~

k,

(M-l sec-1)

v Y ~

(La)

PK.

~~

OH

CH3

H

FCHz

ClCHa

CF3

CHZOH

1200 13.4 -

1020 22.7 15.9

830 5.7 15.1

470 26.9 -

95 36.8 14.3

32 35.3 12.4

26 30.4 14.8

Horse erythrocyte catalase; 5 m M phosphate, pH 7, 25". bvan der Waals volumes of Y (186, 186) [Chance (56) and White and Schonbaum (118b)l.

Little evidence is available, therefore, that reductants induce redistribution of electronic charge within the prosthetic group in such a manner that oxygen of EO becomes the locus of the nucleophilic attack. The best reductants of Compound I are "hard" donors rather than polarizable substrates (cyanide, thiols, carbon monoxide, and isonitriles)-observations which cannot be readily reconciled with the "hydroxylation" mechanisms (118) outlined in Eqs. (22), (32), and (33):

"'" [ :Hy>m] 0- 1EO-N

EO

+

E(H,O)

EO

+

H,O,

E05O@

E(H,O)

+ +

HN Y N * N NJ

(32)

0,

(33)

[H

4. The data on alcohol oxidation further underscore the difficulties of mechanistic interpretation [Table IX (35, 118b, 185, 186) and Table X (118b)1. Thus, although k,, values for the reduction of horse erythrocyte catalase Compound I by ethanol, n-propanol, and n-butanol (Table X) support the classic view (35) that steric (volume) constraints influence the reaction rates, other effects became apparent on comparing the rates of oxidation of methyl, n-propyl, allyl, and propargyl alcohols (118b) (Tables IX and X ) . The latter is clearly a superior reductant, not only relative to other three-carbon alcohols but also compared to methanol. An inductive (electron withdrawing) effect of the acetylenic group may be of importance here but hardly dominant since 2-fluoro-, 2-chloro-, and 2,2,2-trifluoromethyl ethanols are by no means the prefer185. J. T. Edwards, J. Chem. Educ. 47, 261 (1970). 186. A . Bondi, J. Phys. Chem. 68, 441 (1964).

402

GREGORY R. SCHONBAUM AND BRITTON CHANCE

REDUCTION OF COMPOUND Ia BY

TABLE X THREE,A N D FOURC.4RBON ALCOHOLS

TWO,

Alcohol Ethyl

Propyl

Butyl

Ally1

Propargyl

0.76 1020 15.9

1.0 6.5 16.1

1.24 0.4 16.1

0.91 330 15.5

0.90 2500 13.6

vRb

kc llpp (M-l sec-l) PK.

0 Horse erythrocyte catalase; 5 mM phosphate, pH 7, 25' [White and Schonbaum (118b)l. * VR denotes van der Waals volumes relative to V R = 1 for propyl.

TABLE XI STEREOSPECIFICITY OF ETHANOL OXIDATION I N BEEF LIVER (PRODUCT ANALYSES) CATALASE-MEDIATED REACTIONS Products" Substrate

CHaCHO

(%I

CHsCDO (%)

S-( -)-Ethanol-1-d Ethanol-1-d (racemic)

0 43k3

100 57f3

aFollowing approximately 65 f 10% of the reaction [Schonhaum (169)].

red donors (Table IX) . Accordingly, neither the steric nor electronic factors are, per se, adequate in accounting for the variation of k, app values, but their interplay as well as the residence time of alcohol in an EO (ROH) complex should be taken into account. That orientation effects are important is particularly well illustrated in the case of ethanol. Its oxidation occurs stereospecifically (Table XI) (169,187,188) and involves loss of the pro-R hydrogen (188). H,C

*

-

H,C-C9 iI'

0

*

(+2H+)

(34)

S-(-)-1-H*-ethanol

To account for other results in Tables IX, X, and XII, it couid be argued that significant interactions in the EO (HXOH) complex would 187. H. Gang, A. I. Cederbaum, and E. Rubin, BBRC 54,264 (1973). 188. R. J. M. Corrall, H. M. Rodman, J. Margolis, and B. R. Landau, JBC 249, 3181 (1974).

7.

403

CATALASE

I

'6

-i

-i

-i -5

-j

-+

-6

TAS.(kcdmle-' 1

FIG.10. Enthalpy-entropy compensations in reductions of Compound I by hydrogen donors: (1) 2-fluoroethanol, (2) ethanol, (3) ally1 alcohol, (4) methanol, (5) propargyl alcohol, (6) hydrogen peroxide, and (7) formic acid. Horse erythrocyte catalase in 5 mM phosphate, pH 7 ( I I H J ] .

result in a more negative entropy of activation and that given a proper orientation of the reactants the enthalpy barrier would be lower. Such an entropy-enthalpy correlation is discernible in the oxidation of several alcohols (Table XIII) but cannot be extended to oxidations of hydrogen peroxide or formic acid (Fig. 10). This is not entirely unexpected since the observed activation parameters reflect the sum of the standard quantities pertaining to the equilibrium between Compound I and the reductant EO

+ HXOH+

EO(HX0H)

AFo

=

-RT In K.,

(35)

and the activation quantities for the transformation of EO(HX0H) t o the transition state. Moreover, different reaction steps appear t o be rate limiting in the Compound I-donor interactions. Thus, significant isoTABLE XI1 REDUCTION

OF

COMPOUND 1 B Y

SOME " T W O

ELF:CTRON

I)ONORS''''

Y-OH

Y OH k4 npp

PKa

1 . 8 x 107 11.6

N=O 1.2

x

3.3

HC=O 107

9

x

106

3.75

N Hz

x 104 > 12

24

Data from Chance (36), Chance and Herbert (176), and Schonbaum (10Ia).

404

GREGORY R. SCHONBAUM AND BRITTON CHANCE

TABLE XI11 THERMODYNAMIC ACTIVATION PARAMETERS FOR SOME APPARENT CATALASE COMPOUND I-MEDIATEDOXIDATIONS' Donor Methanol Trideuteromethanol Ethanol 1,1-Dideuteroethanol Ally1 alcohol 2-Propy n-l-ol 0

AHS (kcal mole-')

ASS (cal mole-' deg-1)

8.4 9.3 10.8

-17.4 -17.6 -8.9 -9.1 -10.2 -21.9

11.1

9.3 0.4

Data from White and Schonbaum (118b).

tope effects, which are principally attributable to AH* (Table XIII) obtain in the oxidation of 1-deutero alcohols (CDaOH and CHaCD20H) but not in the oxidation of deuteroformate (DC02H) (Tables XIV and XV) or deuteroethanol (CHaCH20D)(Table XVI). In particular, the large differences in k4 values and kH/kD ratios for formate and methanol oxidations cannot be simply attributed to steric constraints. Rather for formate, the formation of the Compound I-donor complex must be assumed to be rate determining. This being the case no isotope effect is expected; indeed, kFapp and k?,,, are nearly equal ( k H / k ~ 1.1 f 0.05) (Table XIV). Note also that rate constants characterizing the interaction of formic acid with the resting enzyme (kl loe M-* sec-I) (37) and with Compound I (k4 0.9 X 106 M-1 sec-1) are nearly the same. Apparently, the active site is equally accessible to substrates in different

-

.,,-

ISOTOPE

TABLE XIV EFFECTSI N BEEF LNER CATALASE-MEDIATED OXIDATIONS' Formic acid

Substrate k4 sppo

10-6

(M-1 sec-1)

k4'b

(M-1 sec-1) kdkn

-

Ethanol

Methanol

HCOzH DCOpH CHaCHzOH CH,CDzOH CHsOH CDIOH 486 8.0s

400 7.64 1.16

890

1.96

455

720

-

130

5.54

In 10 mM potassium phosphate, pH 7, 24.7'. For un-ionized form using pK.(HC02H) 3.75; pK.(DC02H) 3.78 [White and Schonbaum (118b)l. (I

7.

405

CATALASE

TABLE XV ISOTOPE

EFFECTS I N CATALASE-MEDIATED OXIDATIONS O F A N D DEUTEROETHANOLS~ ETHANOL

Rate constant (M-l sec-l) Horse erythrocyte

Substrate

1020 f 20 1020 f 20 460 f 10 2.22

Ethanol S-(-)-1JDeuteroethanol 1,l-Dideuteroethanol Pentadeuteroethanol kcE,c~,on/kcHIcDtoH a

Beef liver

&I. lysodeikticus

890 f 20

21.5 f 0 . 5 20.0 f 0 . 2 14.2 0 . 3 1.51

455 f 10 445 f 10 1.98

*

I n 10 m M potassium phosphate, p H 7, 24.7" [Schonbaum (lola)].

oxidation states of the enzyme. The simplest permissible reaction scheme is therefore EO

+ H X O H kk-il ' E O ( X H 0 H ) 2 E(H20) + XO

(36)

and

Hence, if k ) > k-l, then k 4 k l . Seemingly, such conditions are met in the Compound I-formic acid reaction. I n contrast, since in the oxidation of methanol kH/kD > 5 , scission of the C-H bond must be rate limiting, suggesting that k t < k-1, and k l a p p = ( k l k z ) / k _ , where k l a p p 830 M-l sec-'l (Table IX). N

TABLE XVI SOLVENT ISOTOPE EFFECTSI N HoRSle ERYTHROCYTE-MI.:DIATED OF ETHANOL OXIDATION Solvent

kc npp (M-I sec-l)

H20n 90 % D20-10 % H i 0 (v/v)* knlo/kDno

1020 20 960 k 30 1.06

*

I n 10 m M potassium phosphate, p H 7.0. In 10 m M potassium phosphate, pD 7.1; 24.7' [White and Schonbaum ( l l 8 b ) l .

406

GREGORY R. SCHONBAUM AND BRITTON CHANCE

Therefore, k2/k-1 = 8 X 10-4, provided that as in the reactions of catalase with methyl hydrogen peroxide (Table VIII) or with formic acid (37), kl- 106 M-' sec-1. Further, since even a t 2 mM MeOH, the kinetics of Compound I reduction by methanol obey the second-order rate law ( I & ) , it follows that k-l/lcl 2 20 m M . Hence, k-1 2 2 X lo'sec-' and kt 2 16 sec-'. The above analysis, which merely indicates the possible lower limits of k-1 Llpp and kz app, presupposes the formation of one EO(HX0H) intermediate in the course of the redox reaction. On this score definitive evidence is lacking. Even this minimum postulate may be inadequate to account for the observation that in reactions of catalase with methyl or butyl hydroperoxides less than 15% of the initial peroxide is converted into the corresponding aldehydes unlike the catalase-ethyl hydrogen peroxide system where formation of acetaldehyde from EtOOH exceeds 50% (Table V I I ) . I n the reactmionwith MeOOH, the low yield of formaldehyde could be attributed to a rapid diffusion of methanol from the active site:

%EO + CH,OH

EO(HOCH*)

(38)

Conversely, high conversion of EtOOH to CH,CHO would result from slower diffusion of ethanol from EO (HOC,H,) ; accordingly, the yield of butyraldehyde from BuOOH should be a t least equally high. Since this is not so, we can only conclude that, compared to ethanol, either the residence time of butanol a t the active site is unaccountahly shorter or that prior to oxidation of ROH, the outer sphere complex, TO (HOR b , rearranges to another intermediate ; an intermediate having a configuration in which the slowly reacting alcohols (Tables IX and X) cannot be readily accommodated. An outer sphere complex differing from EO(HOR), the possible initial product of E ( R O 0 H ) reaction, or an inner sphere complex [Eq. (24) ] could meet such criteria. However, no distinctions can be even attempted a t IUP,S(i i r We shall merely note the following: (a) Lack of isotope effect in the oxidation of C H,OD (Table XV) is inconsistent with the reaction pathway outlined in Ey. (23) ; (b) Formation of an inner sphere complex would be subject to a stringent steric control ; (c) Significant displacement of metal ion frorn the porphyrin plane,. or doming of the porphyrin macrocycle should be prerequisites to an inner sphere ligation ; (d) Addition a t iron, if any, cannot be random; otlierwise oxidations would not proceed stereospecifically (Table XI) ;

7.

407

CATALASE

(e) Oxidations proceeding by an inner sphere mechanism are formally analogous to various metal-catalyzed reactions, among others by Cr (VI) (189, 190), Pb(OAc), (191-193), or V ( V ) (194-196), and could be expressed as suggested by Taqui Khan and Martell (197) [Eqs. (38a) and (38b)l: /--

,.o -P0, H

OxE-F e y ’“OH

T__

E-Fe-

+

0,

+

OH-

(384

where OxE is a two-electron oxidized form of the active site, or as outlined in Eqs. (39)-(41) and Fig. 11.

A reminder that the above suggestions are no more than working hypotheses should hardly be necessary. 189. K. B. Wiberg, in “Oxidations in Organic Chemistry” (K. B. Wiberg, ed.), Part A, p. 69. Academic Press, New York, 1965. 190. J. K. Beattie and G. P. Haight, Jr., i n “Inorganic Reaction Mechanisms” J. 0. Edwards, ed., Vol. 17, Part 11, p. 93. Wiley (Interscience), New York, 1972. 191. K. Heusler and J. Kolvoda, Angew. Chem., Int. Ed. Engl. 3, 525 (1964). 192. Y. Pocker and B. C. Davis, JACS 95,6216 (1973). 193. R. Criegee, in “Oxidations in Organic Chemistry” (X. B. Wiberg, ed.), Part A, p. 277. Academic Press, New York, 1965. 194. K. Kustin and D. L. Toppen, Inorg. Chem. 12, 1404 (1973). 195. J. S. Littler and W. W. Waters, J . Chem. Soc., London p. 2767 (1960). 196. J. S. Littler, A. I. Mallet, and W. W. Waters, J. Chem. SOC.,London p . 2761 (1960). 197. M. M. Taqui Khan and A. E. Martell, “Homogeneous Catalysis by Metal Complexes,” Vol. I, pp 139-142. Academic Press, New York, 1974.

408

GREGORY R. SCHONBAUM AND BRITTON CHANCE

ACKNOWLEDGMENTS Research in the authors' laboratories and preparation of this review were supported by USPHS GM-12202, AA-00292 and HL-16061 (to B.C.),by the Medical Research Council of Canada (MT-12701, and by NSF-GB 41635 (to G.R.S.)

Author Index Numbers in parentheses are reference numbers and indicate that an author’s work is referred to, although his name is not cited in the text. Alm, R., 70, 88(130) Alonso, C., 65 Alpert, Y., 317, 318(109) Abacherli, E., 26 Altschul, A. M., 345, 346(1, 2), 347(1, 21, Abatwrov, L. B., 14, 31(56) 348(1, 21, 353(1) Abe, K., 367, 368, 393(69), 394(69) Amelunxen, R. E., 2(17), 3, 19(17) Abeles, R. H., 143 Anan, F., 367,368,393(69), 394(69) Abney, R., 133 Anderson, B. M., 30 Abraham, R. G., 274,275(344) Abrams, R., 345, 346(1, 21, 347(1, 21, 348 Anderson, L., 256 Andrkasson, L. E., 317, 336(108), 336(108) (1, 2), 353(1, 2) Andreev, V. P., 300 Abrams, W. R., 279, 280 Andreeva, N. S., 9 Ackerman, E., 395 Ackrell, B. A. C., 177, 245, 247, 248(26), Andreoli, T. E., 70, 76(127), 77(127) Andrews, E. C., 313 24906, 1991, 253(184) Andrews, L. J., 384 Adams, M. J., 10, 24 Anfinsen, C. B., 86, 132 Adelson, G. I., 105 Anselme, J. P., 379 Adija, D. L., 29 Antonini, E., 335, 336(192), 349 Aebi, H., 365, 367,379(63), 280(63) Appell, G., 366, 367(41), 370(41), 371(41) Agar, N. S., 142 Appleby, C. A, 264 Agatova, A. I., 25 Appleman, D., 376 Agner, K., 366, 385 Archakov, A. I., 153 Agrawal, B. B. L., 377 Arigoni, D., 252 Aikawa, T., 109 Arnold, H., 48 Akagi, J. M., 286, 295(388) Arnold, L. J., 29 Akeson, A., 200 Alben, J. O., 317, 321(113), 322(113), 323 Arnon, D. I., 54, 55(12), 66(12), 66(12), 366 (1131, 337(113) Arosio, P., 246, 247(191) Alberty, R. A., 188, 189(57), 199(57) Arrigoni, O., 237 Albracht, S. P. J., 187 Arscott, L. D., 92, 93, 94(36), 100(36, 371, Alexander, A. G., 300 103(36, 63), lOQ(61,6 3 , 105(85), 106 Alifano, A., 73 (36), 107(36), 118(63), 119(63, 85), Alimov, G. A,, 153 120(61, 63), 121(63), 123(61), 129, 135 Allen, G. A., 23,24(85), 34 (601, 136(60), 137(60), 143(36), 144 Allgyer, T. T., 110 (36) Allison, W. S., 2(15), 3, 20,21(66), 28(15), Artavanis, S., 5 30, 39 409

A

410 Arvy, L., 300 Asada, K., 274,286(359), 295(359) Asahi, T., 91, 133(7), 143(7), 279 Asakura, T., 346,347,348(27, 28,29, 30, 31,

AUTHOR INDEX

Barlow, C. H., 314, 315(93), 319, 321, 322

(1421, 323(140), 329(93), 334(94), 337 (117, 118), 338(94), 339(117,118), 365, 368, 389 Baron, J., 150 32, 37), 349(28, 30, 321, 360(27, 31) Barrett, J., 301 Asano, A., 66, 72(89), 73(89), 76(89) Barldn, E. S. G., 261 Asnis, R. E., 91 Barry, R. E., 164 Asriyants, R. A., 48 Bartlett, G. R., 261 Asyis, R. Aat., 48 Atchison, R. W., 54, 56(18), 57(18), 58 Bartsch, R. G., 371(84), 372 Basford, R. E., 222 (18) Basolo, F., 375 Atkin, C. L., 143 Atkinson, D. E., 274, 277(339), 286(339) Basu, D. K., 106, 107(98) Bateman, L., 395 Atkinson, M. R., 109 Batke, J., 25, 40 Auda, B. V., 260 Auts, S. D., 165, 166(369), 167(374), 168 Bathe, G. R., 84 Baudhuin, P., 368 (374), 169(374) Baudras, A., 265, 268(287, 289), 269(287, Averback, B. C., 274,275(344) 299, 302) Awasthi, Y. C., 312; 313(84) Bauer, V. A., 143 Assi, A., 72 Baugh, R. F., 182, 187, 188, 189, 190(43) Azzone, G. F., 67, 72(102), 214, 282 Baum, H., 70 B Bayne, R.. A., 315,317,337(115) Bearden, A. J., 210 Baccarini-Melandri, A., 2(28), 3 Beattie, D. S., 81(176), 217 Bach, S. J., 263.264 Beattie, J. K., 407 Bachmanova, G. I., 153 Beck, W. S., 143 Bader, P., 235 Baggott, J. P., 33, 34(156), 166, 167(380) Bednars, A. J., 258 Baginsky, M. L., 106, 107(112), 108(112), Beetlestone, J., 346, 372, 375(99) 112(112), 205, 224, 236(156), 243, 244 Behme, M. T. A., 43 Beinert, H., 100, 101(69), 179, 184(34), 185 (156), 246(156) (34, 46), 186(46), 187(34,46), 193(46), Bailey, K., 3 205, 214, 215(46, 541, 216(116), 219, Bakker, E. P., 315, 316(102), 320(102), 220(136), 221(46, 1361, 226(218), 235, 321(102) 244, 245, 253, 297, 307, 309, 316(66), Balastero, F., 237 320, 321, 323, 330, 331(139), 332(126, Baldesten, A., 93, 143(46), 144(46, 275) 1391, 333(139), 335(66, 126), 336(66, Balegh, M. S., 370 194) Balthasar, W., 31 Bell, J. J., 65, 83(84), 84(84) Baltscheffsky, H., 74, 254, 256(220) Benjamin, B. M, 39, 45(181) Baltscheffsky, M., 74, 254,256(220) Benohr, H. C., 130,131 Banaszak, J. J., 10, 11(51), 12(51) Berger, A., 379 Banaszak, L. J., 9, 10, 35, 317, 318(112) Berger, T. J., 68,71(115), 79(115) Bandi, L., 83 Bandurski, R. S., 91, 133(7), 143(7), 274, Berghauser, J., 24 Berglund, O., 143 279, 286(359), 295(359) Bernath, P., 222, 223(144), 225(144), 226, Banerjee, R., 317, 318(109) 271, 273 Baranowski, T., 9 Bernhard, S., 33, 34, 35(160), 36(160), 37 Barea, J. L., 274 (174, 175), 41(160), 42(165), 48(165), Barela, T. D., 15 49(165) Barker, H. A., 143

411

AUTHOR INDEX

Bernheim,. F., 263 Betheil, J. J., 31,34(137) Betz, G., 150 Beutler, E.,131,132(212),132 Beychok, S.,123, 125(168) Bianchi, G.,262 Bielski, B. H., 398 Biggs, D. R , 217 Bilimoria, M. H.,167, 168(384, 386), 170 (386) Bittmann, R.,31 Bjorkhem, I., 83 Black, S.,91,133, 141, 143(6) Blair, P.V.,237 Blakley, R. L.,143 Bloch, W., 34,42(165),48(165),49(165) Blumberg, W. E.,368, 369, 370(77, 78), 376(77), 380,381(126b), 390(126b) Bock, R. M.,310 Boger, P.,54, 62 Boeri, E.,264, 270, 272 Boers, W., 28,33 Bolotina, I. A., 25, 31(105) Bond, J. C, 24 Bondi, A., 401 Bonner, W.D., Jr., 215 Bonnichsen, R. K.,366 Booth, F.W.,300 Borg, D. C., 356,396,397(173) Boross, L., 28, 43 Borst, P.,80 Bossi, E., 365 Bourne, G.H., 300 Boxer, G.E.,47 Boyd, G.S.,83 Boyer, P.D., 39,40,74, 75, 101 Braams, R.,335 Brady, A. H.,123,125(168) Brady, W. T.,93 Bragg, P. D., 64,68(62), 72(62), 79, 80 Bramlett, R.,284,285(382),297 Branden, C. I., 10,11(51),12(51) Brandt, K.G.,94,112(52,53),113(52,53), 114(53), 135(52,53),137(53), 139(53), 141(53) Branzoli, U.,101 Bresters, T.W., 106, 107(114),114(114) Bridgen, J., 5,22 Brierley, G.,180,312,313(85) Brill, A. S.,351, 365,369, 370(74, 791,372.

(74,79,81),374(81), 376(79), 385(16), 389(16), 390,396(156) Brocklehurst, E. S., 321,322(138), 326(138 327(138),331(138), 333(138), 335(138) Brodie, A. F., 64, 65, 66(60), 68(60), 72 (60) Brodie, J. D.,237 Bronk, J. R.,62,65(34), 67(34) Brosemer, R. W., 2(23), 3 Brosnan, J. T., 82,86(186) Brown, J. P.,104,118(86),119(86), 120,121 (86) Brown, N. C., 142. 143(262) Brown, S.B., 389 Brownie, A. C., 83,84(192) Brumby, P. E.,114, 122(155), 123(155) Bruni, A., 239,244,246 Brunori, M.,315, 316(99), 323(99), 324 (99),335, 336(99, 192), 389 Bryla, J., 110 Bucher, T., 81 Buege, J. A., 165, 166(369, 3741, 167(374), 168(374),169(374) Buehner, M., 9,10(46), ll(46, 48),24(47), 29(47), 39(47, 48), 44(46, 47) Bulger, J. E.,94, 112(52, 53), 113(52, 53), 114(53),135(52,531, 137(53), 139(53), 141 (53) Burgoyne, L. A.,267 Burleigh, B. D., Jr., 92, 100(33), 103(33), 104(33,61), 105(33), 119(33), 120(61), 123(61) Burma, D. P., 106,107(98) Butler, W. L.,239,240(178) Butlin, J . D., 68,78,79 Butow, R.A.,186,203(48) Buzard, J. A.,129

C Caifa, P., 247 Caldwell, B. V.,84,88(225) Camerino, P. W.,320, 333(131) Cameron, B. F.,372,373(96) Cammer, W., 65,84(83) Canellakis, Z.N..142,143(262) Cantz, M.,365, 367,379(63), 380(63) Capaldi, R. A., 180, 311, 312(75) Capeillere, C., 268, 269(299) Caputo, A., 349

412 Caputto, R., 2,3(7) Cardenas, J., 274 Carlson, C. W.,2(23), 3 Carpenter, E.,312 Cam, M.L.,150 Carr, N. G.,2(27), 3 Carraway, K.L., 48 Carroll, W.R.,10s Casida, J. E.,177, 204, 205(22, 881, 206 (22) Casola, L., 112(150), 113, 114(150), 122 (150, 155), 123(155) Caswell, A. H.,215 Caughey, W. S., 302, 304, 305(40), 306 (52), 307, 308(52), 313(52), 314(60, 62), 315(52, 93), 316, 317(52, 651, 318 (52, 65), 319, 321(113, ll6), 322(62, 65, 113, 114, 115, 1421, 323(65, 113, 114, l40), 325(65), 328(65), 329(65, 93), 330(104), 334(60), 337(15, 113, 115, 117, 118),338(27, 40, 80,62, 651, 339(117, 1181, 340(116), 343(60, 621, 364, 389 Cavallini, D., 304 Cederbaum, A. I., 81(176), 402 Cepure, A., 101 Cerletti, P.,222, 225(150), 230(157), 231, 234, 237, 246, 247(191), 248 Challoner, D. R., 300 Chamalaun, R.A. F. M.,82 Chan, P. C., 398 Chan, S. H.P.,310,311(69) Chance, B.,31, 72, 204, 207, 215,219,258, 259(229), 301, 321, 324, 335(21), 336 (1551, 346, 348(22), 351(22), 352(6, 22),353(4,6, 221, 356(5), 363(5), 364, 365, 366, 369(1), 372, 374(37), 385 (37), 388(37, 52), 388(36), 389, 390 .(150), 391(150), 393(1), 394(1), 398 (36,1541,399(36), 400(146), 401, 403, 404(37), 406(37, 146) Chang, J. Y.,377,378 Chang, 5. H.,133 Changeaux, J. P.,31 Chantrenne, H.,347, 348(43) Chappell, J. B.,207 Charache, S., 322 Charalampous, F. C., 312 Chen, W.L.,312 Christensen, J. R.,261

AUTHOR INDEX

Christian, W., 2 Chuang, T.F.,312,313034) Chung, A. E., 53, 55, 56(10, 18), 57(10, 181, 58(10, 181, 60 Chung, C. W., 274 Cilento, G.,28 Ciotti, M.M.,30, 52, 53(1), 54(1), 56(l), 58(1, 31, 62(1), 65(31), 86(75) Claisse, M., 269 Clark, W.M.,130 Clegg, R.A.,217,219(131), 220(135) Cleland, W.W.,40, 76, 88, 139 Click, E. M.,288 Clodfelder, P.,39,45(185) Cobley, J. G.,219,220(136), 221(136) Cochran, D.G.,258,259034) Cocriamont, C., 347 Cohen, B. S.,151, 152(318, 320), 153 Cohen, I. A., 302, 304, 338(27) Cohen, P. T.,53, 54, 56(17), 57(6), 58, 59 (17), 60, 61 Cohn, M. L., 100, 103(65), lOQ(65) Coleman, R.,180, 181(41) Coles, C. J., 225, 226, 230(158), 234(15@, 248(158) Colli, W.,88 Collins, D. M., 376 Collipp, P.J., 216 Colman, R. F.,141 Colowick, S. P.,3, 28, 29(13), 38(13), 39 (131,40(13), 44, 52, 53(1), 54, 56(1), 58(1, 2, 3, 5), 62(1 ), 65(30), 67(30), 69(30), 70(30), 73(30), 76(30), 167 Comstock, D. A.,398 Conn, E. E.,92 Conney, A. H.,149 Connors, M.J., 30, 39 Conover, E.,72 Constantinides, S. M.,25 Conway, A., 31, 33, 34(143), 42(143) Cook, D.E.,153 Cook, K.A.,274 Coon, M.J., 91, 149, 150(295), 151(295), 153, 165(286), 166(370, 3711, 167(370, 371), 169 Cooper, A., 371(85), 372 Cooper, D.Y.,83, 91, 152, 166 Coratelli, P.,88 Corcoran, D., 148, 151(284a), 164 Cordes, E. H.,43

413

AUTHOR INDEX

Cori, C. F., 2, 3(8), 48 Cori, G. T., 2, 3(8), 48 Cori, O., 261 Cornforth, J. W., 252 Corrall, R. J. M., 402 Correa, W., 131 Corte, E. D., 132 Coulson, A. F. W., 347, 350, 353(34), 355 (34, 36)

Countryman, R., 376 Cox, C. D., Jr., 274 Cox, D. J., 93 Cox, G. B., 64,68, 78(73), 79, 81(73) Crane, F. L., 68, 79, 312, 313(84) Crawford, I. P., 254, 256(220) Creaghan, I. T., 110 Cremona, T., 69, 78, 177, 184, 188, 189, 190(58, 841, 202, 203, 270, 271, 272, 273 (320) Cresswell, C. F., 274 Creutz, C., 375 Criddle, R. S., 310 Criegee, R., 407 Crifo, C., 304 Cronin, J. R., 259, 260(243), 262(243) Cross, D. G., 29 Cseke, E., 28, 43 Cunningham, L. W., 39, 45(185) Cunningham, W., 313 Curdel, A., 272, 273(329) Curnyn, C., 66, 72(.87) Curti, B., 101 Cusanovich, M. A., 325 Cutolo, E., 264 Czerlinski, G., 117 Czygan, F. C., 274

D Dade, E., 46 Dahlen, J. V., 88 Daigo, K., 93, 108(43) D’Allessio, G., 2(16), 3, 4(16) Dalling, M. J., 274 Dallner, G., 166 D’Aloya, R., 63,87(53) Dalziel, K., 139 Dandliker, W. B., 24 Daniel, L. J., 261

Danielson, L., 64, 65(56, 57, 58), 67(56, 57, 58), 68(56, 57, 581, 72(56, 57, 58), 73(57, 581, 74(56, 57, 581, 77(56, 57, 58) Danielsson, H., 83 Darnall, D. W., 15 Davidson, B. E., 2(25), 3, 5,9(33), 20(33)

Davidson, D. W., 83 Davidson, J. T., 282, 284(377) Davies, J. L., 304, 305(40), 338(40) Davies, P. L., 80 Davis, B. C., 407 Davis, K. A., 179, 191, 192(71, 73), 193 (32, 691, 200(69),. 203, 222, 224(141, 142, 143), 225(141, 142, 143), 226(32), 227, 228, 229, 230(73, 141, 142, 143), 231, 232(71, 166), 233, 234, 235(143), 236, 237(166), 239,240(166), 241(166), 242(166), 243(166), 244(143), 245 (143), 253,254,246(143), 248(71,166), 254, 255, 256(220) Davis, P. S., 170, 288, 290(413, 414, 4151, 291(414), 292(413, 415), 294(414) Davison, A. J., 324 Dayhoff, M. O., 105, 371(83), 372 Deacon, T. E., 282, 284(377) Deal, W. C., Jr., 25, 30(108), 48(108) DeBernard, B., 155, 189 de Duve, C., 365, 367(23), 368 de Haan, E. J., 63, 82, 86(191), 87(53) Deisseroth, A., 365, 385(18), 389(18), 397 (18) De Kok, A., 101, 106, 107(114), 114(114), 124, 125(172), 238 De La Chica, G., 133 De Lorenzo, F., 132 De Luca, C., 65 De Luca, H. F., 83 De Marco, C., 304 Dennis, D. T., 2(26), 3, 39, 40(!26), 45 (188), 48(26) De Rosier, D., 126 DerVartanian, D. V., 124, 125(172), 187, 221, 222, 224(149), 226, 236(149), 237 (149), 238(149), 251, 252, 253(149), 281, 284, 285(282), 288, 295(373a) Deutsch, H. F., 366,382(39) Devichensky, V. M., 153 De Vijlder, J. J. M., 31, 32, 33(144, 1511, 34(144), 41(159)

414

AUTHOR INDEX

Devlin, T. M., 62, 65(33) Eichner, R. D., 29 De Wael, J., 131 Eichorn, J., 83 Dewan, J. D., 2 Eigen, M., 31 Diamond, L. S., 66 Eik-Nes, K. B., 83 Dick, A., 259 Eisele, B., 28, 33 Eisenberg, D., 371(851, 372 Dickerson, R. E., 371(85), 372 Ejima, A., 108 Diehl, H., 152 Ekstrand, V., 93, 94(40), 138(40), 139 di Franco, A,, 268 (40), 140(40) Dixon, M., 2,3(7), 109, 263, 264, 267 Djavadi-Ohaniance, L., 212, 213, 296, 297 Eldjarn, L., 130 Eley, M. H., 108 Doeg, K. A., 224,236(153), 239 Elias, H. G., 24 Dolphin, D., 356, 396, 397(173) Ellfolk, N., 347, 348(40, 41, 42) Dontsov, A. E., 74,75(148) Elliott, J., 46 Dorfman, R. I., 83 Elliott, P., 143 Dorsey, J. A., 88 Dounce, A. L., 365, 366, 367, 385(18), 389 Elodi, P., 2(14), 3, 23, 25, 26, 31 Engel, P. C., 91,100 (18), 397(18) Englard, S., 31, 34(137) Drabikowska, A. K., 258 Eriksson, A., 133 Drabkin, D. L., 300 Eriksson, L. E .G., 235 Dragoni, N., 286, 295(389) Eriksson, S. A,, 132 Dreyfuas, J., 286, 287(387), 288 Dro t t, H R., 347, 348(32), 349(32), 360 Erman, J. E., 347, 350, 353(34), 355(34), 372, 376(88) DuBus, R., 91 Ernest, M. J., 130 Due& E., 9, lO(45) Duggleby, R. G., 2(26), 3, 39, 40(26), 45 Ernster, L., 64, 65(56, 57, 58), 67(56, 57, 58), 68(56, 57, 581, 68(66), 69(68, 69, (188), 4806) 106), 70(59, 67), 72(56, 57, 58, 67), Duncan, H. M., 216 73(57, 58, 59, 102, 103, 106, 135, 136, Duncan, I. W., 93, 94(40), 138(40), 139 137), 74(56, 57, 58, 106, 136, 1371, 75 (40), 140(40) (67, 68, 69), 76(59, 67, 68,69), 77(56, Dunstan, W. R., 399 57, 58, 59, 67, 68, 69, 106, 135, 136, Duppel, W., 149, 150(295), 151(295) 137), 78, 82, 86(191), 87(74, 1841, 88 Dupre, S., 340 (129, 130), 166, 168, 204, 207, 212, 214, Durchschlag, H., 32 246, 249(188), 261 Dus, K., 371(84), 372 Escamilla, E., 133 Dutton, P. L., 215, 325 Estabrook, R. W., 63, 65, 66(37), 81(37), E 83, 84(83), 86(37), 91, 151, 152(318, 320, 321), 153, 168, 186, 203(48), 258, Eberhard, C. A., 106, 107(97) 259(233), 261, 262(262), 387 Eberspaecher, H. I., 305 Evang, A., 368 Ebisuzaki, K., 260 Eylar, E. H., 317, 318(112) Eby, D., 29, 30(126) Edelhoch, H., 101, 189 Edelstein, S. J., 101 F Edmondson, D., 222 Faeder, E. J., 170, 287, 288, 290/415), 292 Edwards, J. O., 399 Edwards, J. T., 401 (415) Fahien, L. A., 26, 31(110) Egan, R. S., 377 Ehrenberg, A., 200, 235, 330, 331(174), Fairs, K., 302 346, 348(22), 351(22), 352(22), 353 Fajer, J., 356, 396, 397(173) Feeney, R. E., 2(22), 3, 26(22) (20,22), 356,387,388,400(143)

.

415

AUTHOR INDEX

Fehrniann, H., 106, 107(106), 112(106) Feigin, L. A,, 366 Feinberg, R. H.,262 Feinstein, R. X., 365 Feldberg, N. T.,249 Feldberg, R.,90,282 Felsenfeld, G., 330 Felton, R. H.,356, 396,397(173) Felton, S. P.,179, 188(37), 189, 195(63), 203(63) Fenselau, A.,23,42,43(202) Ferri, G., 22,44(76),101 Fife, T.H.,39,45(181) Filmer, D.,32,33(149) Fisher, H.F., 29 Fisher, R. J., 64, 65, 67, 68(61), 72(61), 79 Fisher, R. R., 68, 69(112), 71(109, 110, 111, 112), 76, 77(118), 78(109, 110, 111, 112), 207, 213 Fitzgerold, B., 48 Flashner, M.I. S., 101 Fleischer, S.,180,312,313 Flohe, L.,130 Fluharty, A. L.,105 Forster, T.,359 Fogo, J. K.,202 Fonzo, D., 84 Forchielli, E.,83 Forcina, B. G., 22,44(76) Ford, G. C.,9, 10(46), 11(46, 48), 24(47), 29(47), 39(47, 48), 44(46, 47) Forestier, J. P., 268,269(302) Forman, A,, 396,397(173) Forti, G., 100 Foster, R., 384 Foust, G. P.,90,94,98(50), 146, 147(284). 282 Fowler, L. R., 179, 258(33), 306(50), 307, 308(50) Fox, J. B., Jr.. 24 Frampton, V. L.,366,367 Francavilla, A,. 63,86(50), 87(54) Francis, S. H., 24,42, 44, 45(90), 48(200) Franklin, M.R.,168 Fraser, D.R.,83 Frech. M.E.,62,65(31) Fredericks, W. W.,54, 59(13). 66(13) Fricke, H.,302 Friedrirh, P.,23

Frisell, W. R., 259, 260(243), 262(243) Furhs, S.,132 Fuchsman, W. H., 304, 305(40), 314, 334 (941,338(40, 94) Fuhrhop, J. H., 396 Fujihara, Y.,320, 325(125), 326(125), 331 (125),332(125) Fukuyoski, Y.,108, 109, llO(132) Furfine, C.,3, 19,31(12), 34(137),40(60), 41(60), 42(60), 44(60), 48(60) Furuta, H., 366,367(50)

G Caber, B. P., 105 Gal, E. M.,106, 107(94) Galante, Y.,178, 214, 246, 247(191), 296, 297 Gallego, E., 31 Gallop, P.M., 379 Galston, A. W.,366 Gang, H., 402 Ganther, H.E.,132 Garhe, A.,24 Garland, P. B., 63, 86, 87(52), 88, 204, 217,219(131), 220(135) Garrett, R. H., 274,275(351),275(350) Garwood, D.C.,119 Garwood, D.S.,119 Gaylor, J. L.,151 Gehl, J. M.,54, 59(13), 66(13) George, P.,304, 346, 361% 9), 364, 369, 372, 375(99), 389(9), 392 Gerari, G., 333 Gerth, E., 48 Ghalambor, M. A., 222, 224(141), 225 (141), 230(141), 232(166), 236, 237 (1661, 240(166), 241(166), 242(166), 243(166), 248(166), 253( 166) Ghazarian, J. G., 83 Ghiretti-Magaldi, A.,271, 273 Ghisla, S.,97, 235 Gibson, F., 64, 68, 78(73), 79, 81(73) Gibson, Q . H., 92,94(24), 97(24),98(24), 107(24), 109, 111(24), 112(24), 113 (24, 1371, 115(24, 137), 116(24), 136, 167(245), 168(245, 386), 169(245), 170(245,3861, 172(245,398), 286, 287 (3941, 290(394), 291(394), 308, 322, 324, 327, 333, 335(153), 336(153)

416 Gierisch, W., 367 Gieseman, H., 383 Gilboa-Garber, N., 286, 287(391) Gillette, J. R.,153 Gilmour, M. V., 311, 312(78), 324, 335 Gioeli, R. P.,65 Giordano, M.,237, 247 Giovenco, M.A.,237,247 Giovenco, S.,247 Girotti, A. W.,372 Giuditta, A.,246,247(189) Givol, D.,37 Glatzle, D.,131 Gleason, F.K.,143 Glenn, J. L.,199,260 Goldman, D.S.,106, 107(111) Gonze, J., 63, 186, 203(48), 262 Goodall, D.,31 Gordon, M.S.,300 Gorjunov, A. I., 9 Goto, M.,150 Gottesman, D.P.,110 Goulding, E.,399 GouLian, M.,143 Grab, D.J., 365 Gralen, N.,366 Grandchamp, S.,217 Grande, H. J., 126 Grant, J. K.,83,84(192) Grataer, W.B.,32 Gray, H.B.,385,386(137) Gray, R. W.,83 Grebner, D.,310,319 Green, A. P.,65 Green, D.E.,2, 75,222,237,256,257(223) Greenbaum, A. L.,46,47(211), 81,82 Greene, J. C.,2(22), 3,26(22) Greenfield, R.E.,366 Greengard, P.,83 Greenstein, D.S.,388 Greenwood, C., 315, 316(99), 322, 323 (99), 324, 327, 335(153), 336(99, 153, 192), 389 Gregolin, C., 270, 271, 272(312) Greville, G. D.,87, 273 Griffith, J. S.,372,375(99) Griffiths, D. E.,64, 68(63), 69, 72(63), 73(63), 178, 179(27, 281, lSl(281, 183 (28), 184(28), 190(27, 281, 245(27),

AUTHOR INDEX

258(33), 306(45), 307, 308(45, 53), 327(53), 330(53) Grinius, L. L., 69,74(120), 75(120, 148) Groot, G. S.P., 73,312 Grossman, S., 219, 220, 221(136) Groudinsky, O.,264, 265, 267 Gudat, J. C.,275 Guerra, F.,84 Guerrero, M. G., 274, 276 (342, 345) Guest, J. R.,110 Guiard, B.,267, 296 Guillory, R. J., 68, 69(112), 71(109, 110, 111, 112, 113, 117), 78(109, 110, 111, 112), 213 Guindon, A. H., 106, 107(97) Gumaa, K.A.,46, 47(211), 81 Gunsalus, I. C.,91,92, 106, 107(110, 115), lOS(lS), 172 Gupta, R. K.,347, 348(38), 357(38) Gurd, F. R.N., 317, 318(112) Guseva, M.K.,2(24), 3,25(24), 30(24) Gustafsson, J., 83 Guthenberg, C.,133 Gutierrez, J., 133 Gutman, M.,69, 177, 186(23), 188(19, 23), 200(19), 201(19, 23), 203(19, 231, 204 (19,23), 205(19, 23,881,206(19), 214, 216(116), 223(23), 224(23), 235(23), 236(23), 238(23), 247(20, 231, 248(23, 24), 249(23, 197), 250(23, 195), 251 252 Gutnick, D. L., 66,79

H Haaker, H., 106, 107(114), 114(114) Haas, E.,167 Haavik, A. G., 178, 179(27, 28), 181(28), 183(28), 184(28), 190(27,28), 224(27), 245(27), 258(33) Haber, A., 398 Hachimori, A.,366, 367(50) Hackert, M.L.,24 Hageman, R. H., 274 Hager, L. P.,377, 378(118), 397(118), 401 (118) Hagihara, B.,323 Hagman, L. O.,348, 364 Haight, G. P.,Jr., 407 Hainfeld, J., 126

AUTHOR INDEX

Halkerston, I. D. K., 83 Hall, D.E.,93,143(46), 144(46) Hall, P.F.,83, 85(209) Halsey, Y.D.,130 Hamada, M.,108, 109, llO(132) Hamberg, M.,83 Hamilton, G. A., 396, 397(167) Hamilton, L.,108 Hammes, G. G.,126 Hanania, G.I. H., 372, 373(96) Haniu, M.,100 Hansen, R. E.,179, 180, 181(41), 184(34), 185(34, 461, 186(46), 187(34, 46), 193 (461, 215(46, 54), 221(46), 309, 316 (66),335(66), 336(66, 194) Hanstein, W. G., 78, 178, 179, 181(80), 191, 192, 193(32), 199(68), 200, 203 (68),206(42), 207(42), 208, 209, 210, 211(80), 212(80), 214(80), 222, 224 (141, 1421, 225(72, 141, 1421, 226 (321,230(141, 1421, 232(71), 236, 237 (1661,240(166), 241, 242,243, 248(70, 71, 721, 253(166) Harano, Y., 84 Harding, B. W.,65, 83(84), 84(84, 211) Hardman, K. D.,317, 318(112) Hargrave, K. R.,395 Harlow, D.R.,66 Harmsen, B. J. M., 32, 33(151) Harrer, C.J., 167 Harrigan, P. J., 21, 23, 35(69), 37(83), 39(69, 831, 40(83), 41(69, 1901, 43 (831, 44 Harrington, W. F, 5, 24 Harris, J. I., 2(18, 19, 211, 3, 4(18, 19), 5, 9(18,33), 14(35, 371, 18, 19, 20(19, 29, 33, 371, 21(29, 30, 34, 37), 22(35, 37), 23, 24(35, 85), 25, 26, 27(103), 34, 36(35), 39, 45(29) Harrison, P. M., 100, 103(68) Harte, E.M.,91, 143(6) Harting, J., 30,39,45(184) Hartree, E.F.,301, 346, 366,367(30), 382 (30),387(30), 388 Hartzell, C. R.,309, 316(66), 317, 320, 323, 325(107, 125), 326(107, 125), 327 107), 330(165), 331(125), 332(125, 126), 335(66, 107, 1261, 336(66, 194) Hasson, E. P.,66 Hatano, S., 274

417 Hatch, M. D., 143 Hatefi, Y., 78, 178(30), 179(27, 291, 180, 181(29), 183, 184(28, 291, 190(27, 28), 191, 192(71, 73), 193(69), 194(40, 671, 195, 196, 197, 198(40), 199(88), 200 (29,45,69),203,204, 205,206(40,42), 207(40, 421, 208, 209, 210, 211(80, 107), 212(80), 213, 214, 216(40, 108), 222, 224(141, 142, 1431, 225(72, 141, 142, 143), 226, 227, 228, 229, 230(73, 141, 142, 143), 231, 232(71, l66), 233, 234, 235(143), 236(156), 237(166), 239,240(166, 178),241(166), 242(166), 243(166), 244(143, 1561, 245(143), 246 (143, 156), 248(70, 71, 72, 1661, 253, 254, 255(220a), 256, 258(29, 331, 296, 297, 306(50), 307 308(50) Hattori, A.,274 Hauber, J., 237,240,247,255 Havsteen, B. H., 31 Hayaishi, O., 189 Hayakawa, T.,106, 107(96), 108, 109, 110 (132) Hayashi, A.,372, 373(92) Hayashi, H., 311 Hayaski, T., 108 Hayes, J. E.,30 Hechter, O.,83 Heidema, J., 149 Heim, W.G.,376 Heimberger, G.,387 Heineman, W.R.,325, 330 Helleman, P. W.,131 Hemmerich, P.,170, 235, 308 Hemmes, R. B., 54, 55, 56(16), 58(16), 59(16), 80 Hems, D. A., 46 Henderson, R.W.,325,326(162) Henley, K. S., 82 Henning, U.,106, 107(107), 108(107) Heppel, L.A.,388 Herbert, D.,366, 367(45, 46), 398, 403 Hershberg, R. L.,364, 374, 385(102) Heusler, K.,407 Hewitt, E.J., 274 Hildebrandt, A., 151,152(321) Hill, E.,10 Hill, F. L.,48 Hilvers, A. G.,33,39,41(159) Hinkle, P.C.,88

418

AUTHOR INDEX

Hiraga, M., 367, 368, 393(69), 394(69) Howard, R . L., 191 Hirayama, K.,165 Howell, L.G.,90,91,146, 147(284), 282 Hiromi, K.,268 Howland, J. L.,63 Hoagland, V. D., 25 Hou, C., 64, 68(62), 72(62), 79, 80 Hoard, J. L.,376 Hrycay, E.G., 150,152 Hoberman, H. D.,69 Huang, J. J., 172 Hochberg, R . B.,85 Huang, P. C.,189, 190(59), 379 Hocking, J. D.,2(19), 3, 4(19),5, 14(37), Hucho, F., 106, 107(92), 108(92) 20(19, 37), 21(37), 22(37) Hudson, B.,91, 133, 143(6) Hockstein, P., 168 Huennekens, F. M.,106, 107(112), 108 Hodgins, D.S., 143 (112), 112(112), 179, 188(37), 189, Hodson, R. C.,282, 283 195(63), 203(63), 216 Hojeberg, B., 53, 58(8), 61(8, 231, 69(8), Huheey, J. E., 334 Hultquist, D.E.,165 70(8), 71(8), 7603) Hoek, J. B., 53, 58(8), 61(8, 231, 68, Humphrey, G. F.,62, 65(32), 67, 69(32), 69(8), 70(8), 70(107), 71(107),76(8), 70(32), 76(32) 78(107), 82,86(191), 87(184), SS(130) Hundal, T.,68,70(107), 71(107), 78(107) Hoeksema, W. D., 286 Husain, M.,297 Hoffman, B. M,360 I Hofmann, T.,92, 100(32), 103(32, 68), 124(32) Ibers, J. A.,302 Hogeboom, G. H., 67 Ibne-Rasa, K.M.,399 Hogenkamp, H. P.C., 143 Icen, A.,93, 94(39), 100(39), 129(39), 132 Hogness, T.R., 167,345,346(1, 21, 347(1, (39), 138(39), 140(39), 141 2),348(1,2),353(1,2) Ichihara, K., 149 Hogue, P.,248 Ide, S.,106, 107(96) Hoguem, P.K.,109, llO(130) Ii, I, 129 Holleman, W.H., 25 Iizuka, T.,347,351, 356(55), 360,372,376, Hollocker, T. C.,249 382(107), 387(100) Holloway, M.R., 25,26; Imahori, K.,2(20), 3 Holloway, P.W., 151 Imai, K.,66, 72(89), 73(89), 76(89), 372, Hollunger, G.,207 373(92) Holmes, R. S.,365 Imai, Y., 151 Holmgren, A., 93, 119(45, 471, 143(46, Ingraham, L. L.,397 47), 144(46,47) Inomata, H., 165 Holtaman, J. L.,150 Irukulla, R., 365 Hommes, A.,70,77(126) Isaacs, G. H., 259 Hommes, F.A.,207 Isaacson, E. L.,150 Hommes, R. W., 63 Isherwood, F.A., 129, 130(191), 138(191) Hong, J., 243 Ishimoto, M.,286,295(395) Hood, W.,2(27), 3 Ishizawa, S.,165 Hooper, M.,65 Ito, A.,154 Hopkins, F.G.,245 Iwatsubo, M.,268, 269(299), 272, 273 Horecker, B. L.,166,167(375) (329) Horgan, D.J., 177, 205(22), 206(22) Iyanagi, T.,165, 166(373), 167(373), 170 Horie, S.,306(47), 307, 322 (373), 171(373), 172(373, 402) Horio, T.,323 J Horney, D.L.,108 Hoskins, D.D.,164 Jacobs, E. E., 306(48), 307,308(48), 313 Hosoya, T.,346 Jacobs, N.J., 260

AUTHOR INDEX

419

(390, 397), 289(390, 3971, 290(394, 413), 291(394), 292(390, 397, 4131, 293(390), 295(412) Kaneda, T., 108 Kanner, B. I., 79 Kanzaki, T., 108 Kaplan, N. O., 2(15), 3, 20, 28(15), 29, 30, 52, 53(1), 54(1), 56(1, 17), 57(6, 71, 58(1, 2, 3, 5, 7, 21, 221, 59(17), 60(7), 61, 62(21), 65(30, 311, 67, 69 (30, 78, 96, 971, 70(30, 961, 72(78), 73(30), 76(30), 77(118), 78(97), 86 (75), 87(24), 110, 167, 168, 169(388), 207 Karpukhina, S. Ya., 366 Karr, G. M., 5, 24 Karuzina, I., 153 Karyakin, A. V, 153 Kaschnitz, R. M., 153, 206 Kaufman, B. R., 67, 69(96), 70(96), 110 Kaufmann, H., 367, 379(63), 380(63) Kawahara, Y., 106, 107(102, 104), 126 (104) Kawakita, M., 187 Kawasaki, T., 65, 69(78), 72(78) Kearney, E. B., 69, 176, 177, 188, 189, 190 (58, 83), 202, 203, 217, 222, 223(25, 140), 224(25), 234(25), 235(25), 236 (25), 237(25), 238(25), 245, 24700, 25), 248(24, 25, 261, 249(25, 26, 197, 199), 250(25, 195, 196, 1971, 251(197), 252(197), 253(184), 254(16, 251, 270 Keefer, R. M., 384 Keenan, T. W., 312, 313(84) Keilin, D., 222, 223, 240(147), 264, 300, 301, 315, 338(14), 346, 363(4), 364, 366, 367(30), 382(30), 387(30), 388, K 398 Keirns, J. J., 311, 312(78) Kadlubar, F, F., 153, 154 Keister, D. L., 54, 55, 56(11, 16), 58(16), Kadziauskas, Y.P., 69, 74(120), 75(120) 59(11, 16), 64, 66(11, 64, 65), 68(64, Kagawa, T., 263 65), 69(65), 72(64, 65), 73(65), 76 Kajiwara, S., 346 Kallai, 0. B., 371(85), 372 ( 6 9 , 80 Keleti, T., 24, 25, 40 Kalra, V. K., 65 Keller, R., 267 Kalse, J. F., 124, 125(172, 173). 238 Kelso, J . R., 300 Kamen, M. D., 371(84), 372 Kamin, H., 91, 136, 166(10), 167(10, 245), Kendrew, J. C., 349, 364, 372 168(10, 245, 384, 3861, 169(245), Kenney, W. C., 109, 110(130), 177, 222, 223(25), 224(25), 225, 226(158), 230 170(245, 3861, 172(245, 398), 176, 278 (158), 234(25, 158), 235(25), 236(25), (3), 286, 287(390, 394, 397, 412), 288

Jacobson, M., 149 Jacq, C., 264, 265, 266, 268(290, 291), 296 Jacquot-Armand, Y., 267 Jaenicke, R., 25, 32 Jajczay, F. L., 372, 373(90), 379(90), 380 (1251, 381(125), 382(90, 125), 383, 384, 385(125) Jalling, O., 261 James, B. R., 302 Jasaitis, A. A., 69, 74(120), 75(120, 148) Jecsai, G., 25 Jedeikin, L. A., 46, 49(214) Jefcoate, C. R., 83, 151 Ji, S., 75 Jick, H., 150 Jocelyn, P. C., 129 Jornvall, H., 5 Johansson, B. C., 254, 255(220), 256(220) Johnston, J. M., 151 Jollow, D., 153 Jones, E. T., 92, 94(35), 100, 103(35), 104 (35), 119(35), 120(61), 123(61), 141 (35) Jones, G. L., 365 Jones, G. M. T., 5, 14(35), 22(35), 24(35), 36(35) Jones, 0. T. G., 274, 275(346) Jones, P., 151, 368, 369, 388, 389, 393, 394 (1621, 395(75), 397(75) Jose, J., 2(16), 3, 4(16) Joy, K. W., 274 Joyner, T. B, 301 Junge, J. M., 2(10), 3 Junk, K. W., 149, 165(286) Juntti, K., 65, 72, 212

420 237(25), 238(25), 247(25), 24805, 158), 249(25), 250(25), 254(25) Kensler, C. J., 260 Kepler, C., 106, 107(97) Kessler, E., 274 Kielly, W. W., 62, g5(34), 67(34) Kierkegaard, P., 348,364 Kim, I. C., 217 Kim, K. H., 130 Kimura, T., 172, 240, 247, 260, 261(256), 262(262), 263(266) King, T. E., 78, 176, 182, 187, 188, 189, 190(43, 43a), 191, 222, 223, 226(151), 235, 236, 237, 240(147, 148, 1671, 242, 244(164), 245(167, 1731, 246, 248, 253 (164), 296, 306(51), 307, 308(51), 310 (51), 312, 319, 320, 322(124), 323 (124), 324, 325(124), 326(124), 331 (124), 332(124), 333(131), 334 Kinon, B. J., 131 Kirkman, S. R., 86 Kirkpatrick, F. H., Jr., 313 Kirsohner, K., 25, 26, 31(101), 32, 33 (101), 41(146) Kirtley, M. E., 29, 30(126) Kita, M., 375 Kitamura, T., 108 Klaaae, A. D. M., 237, 238(172)

AUTHOR INDEX

Konings, A. W. T., 68, 71(113, 117) Kopko, F., 129 Korgenovsky, M., 260 Koritz, S. B., 83 Koshland, D. E., Jr., 26, 31, 32, 33(149),

34(143), 35(164), 36(113), 37(113, 172), 42(143) Kosow, D. P., 47 Kosower, E. M., 28, 97, 131, 379 Kosower, N. S., 131 Koster, J. F., 124, 125(172), 126(180), 238 Kotani, M., 347, 351, 356(55), 366 Kowal, J., 84 Koyama, J., 286 Kramer, R., 67, 69(100), 70(98), 78(100) Kratky, O., 32 Krebs, E. G., 2(10), 3 Krebs, H. A., 46, 47, 49, 63, 79, 81(175), 82(175), 86 Kremer, M. L., 305,366,385 Krimsky, I., 13,28,39(54), 40,41, 42(196) Kronman, M. J., 25 Krul, J., 106, 107(114), 114(114) Krupas, M., 269 Kubose, A., 106, 107(94) Kuboyama, M., 306(51), 307, 308(51), 310, 334 Kuma, F., 165 Kumar, S. A., 189, 195(63), 203 Klein, K. O., 84 Kupriyanov, V. V., 70 Klein, S. M., 106, 107(113), 108(113) Kustin, K., 407 Kleineke, J., 82 Kusunose, E., 149 Kleinsek, D. A., 88 Kusunose, M., 149 Kleppe, K., 101 Kuwana, T., 317, 320, 325(107, 125), 326 Klima, J., 167 (107, 125), 327(107), 330(165), 331 Klingenberg, M., 56, 57(20), 63, 66(20), (125), 332(125), 335(107) 81(20, 38), 86(38, 45, 461, 258, 259 Kuenetsova, G. P., 153 (228) Klouwen, H., 180, 312 1 Klybas, V., 44 Knof, S., 25 Labeyrie, F., 265, 267, 268,269, 272(306) Knutson, J. C., 83 Ladany, S., 85 Kobayashi, K., 286, .295(395) Lafferty, M. A., 274, 275(351) Kodicek, E., 83 Lakatos, S., 26 Koeppe, 0. J., 31 Lakshmanan, M. R., 88 Koike, K., 108 Koike, M., 93, 106, 107(96, 109), 108, 109, Lam, K. W., 79 Lambert, J. M., 45 110(132), 114(109) Lambeth, D. O., 375 Kolb, E., 2(21), 3,5 Lambowitz, A. M., 215 Kolvoda, J., 407 Lamprecht, W., 24 Komai, H., 311, 312(75)

AUTHOR INDEX

Land, G., 143 Landau, B. R.,402 Landriscina, C.,88 Lands, W.,168 Lang, G.,331,356 Langdon, R. G., 91,129,130, 142, 166(11), 167(11, 380), 168(11) Lange, L. G., 111,213 Lange, R.,20, 368 Langemann, H.,260 Langford, C. H., 385,386(137) Lardy, H.A., 110, 256, 259, 263 Lardy, M. A., 62, 65(35), 67(35) Larsson, A., 143, 144 Larsson, L. L., 364 Larsson, L. O., 348 Laskawska-Klita, T.,310 Laughrey, E.G.,82 Launay, A. N.,84 Laurent, T. C., 92, 142(23), 143(23) Lazdunski, M.,39 Lazzarini, R.A,, 274,277(339), 286(339) Leadbetter, E.,274, 276(342) Lean, J. D., 57,58(21),62(21) Lebeault, J. M., 149,150(295), 151(295) Lebherz, H.G., 9,26 Lederer, F., 264, 265, 266, 267, 268(290, 2911, 296 Lee, C.P., 64,67,68(66), 69(106), 70(59), 72(59, 102, 103, lW),73(106, 135, 136, 1371, 74(106, 136, 137, 166), 76(59), 77(59, 106, 135, 136, 1371, 78, 87(74), 204,207,214,246 Lee, C.Y., 29 Lee, J. P.,286,288,295(396) Lee, K.L., 259 Lee, R., 168 Lee, Y.P.,259 Lees, B.D,329 Le Gall, J., 281, 286, 288, 295(373a, 389, 396) Lehninger, A. L., 65, 92 Leigh, J. S., 187, 296, 320, 322(124), 323 (124), 324, 325(124), 326(124), 327, 331, 332(124, 168), 333(168), 336 (155),350,360,365, 372,376(88), 389, 390(150), 391(150) Leinweber, F. J., 286,287, 288(404) Lemberg, R.,301, 315, 316(18, 1001, 320 (18), 322, 323(100), 324, 335(18)

421 Lenaz, G., 70 Lenhoff, H.M.,168,169(388) Lents, P.J., 24 Leonard, J., 347, 348(39), 349(39), 359 (39) Lerner, D. A.,126 Leterrier, A. F.,317,318(109) Levenberg, M.I., 377 Levin, W.,149, 152 Levitzki, A.,34,35,36 Levy, H.R.,39 Lewis, J. A.,396 Lewis, S. N.,399 Liberman, E. A., 69, 74(120), 75(120) Libor, S., 23, 25 Lichtenberger, F.,153 Lieber, C. S.,81(176) Lieberman, S.,83,85 Light, P. A., 204, 217, 219 Lilienthal, H.R.,314,334 Liljas, A., 10,11(51), 12(51) Lim, J., 235, 244(164), 253(164), 296 Lindberg, O.,261 Lindsay, J. G.,302, 321, 322(138), 323, 326(138), 327(128), 331(138), 333 ( 1381, 335(138) Linn, T. C., 106, 107(92), 108(92) Linnane, A. W.,216 Lipmann, F.,38,74, 279 Lippard, S. L., 331 Lipscomb, J. D.,172 Lipscomb, M.D., 83, 84(210) Listowsky, I., 31, 34037) Littler, J. S.,407 Lo, S., 361 Low, H.,261 Loewus, F. A., 39 Lombardino, J. G.,379 Long, R.W.,88 Lopez-Colome, M., 133 Losada, M.,274,276(342, 345) LoSpalluto, J., 150 Louie, D.D.,53,57(7), 58(21, 22), 60(7), 62(21) Love, B., 310, 311(69) Lovenberg, W.,176,243 Loverde, A., 156,164(349) Lovrien, R.,367 Lowe, G., 43 Lowry, C., 24

AUTHOR INDEX

Lowry, 0. H., 47 Lu, A. Y.,149,152,165(286) Lund, P.,46,82,86(185) Lundsgaard, E.,2 Lusty, C. J., 106, 107(91, 93), 109(91), 202, 262, 263(266) Luthy, J., 69, 203 Lutwak-Mann, C.,245 Luzikov, V. N.,70 Luzzati, M., 264 Lyanagi, T., 169 Lyric, R. M., 281, 282(375), 284 Lyster, R.J. L., 369

M McAllister, J. K., 92, 94(36), 100(36), 103(36), l06(36), 107(36), 143(36), 144(36) McCann, L. M., 68,79 McCarthy, J. L., 84 McCarthy, K., 243 McCay, P. B.,168 McCleverty, J. A., 396 McCormick, D. B.,101, 125 McCoy, S.,304, 305(40), 317, 321, 322 (114),323(114), 337(115), 338(40) McDaniel, M. C.,334 McDonald-Gibson, R. G.,247 McGraw, J. C., 164 Machinist, J. M., 78, 190(84), 202 Machleidt, W.,311 McIntosh, E.N.,65 McKee, E. M., 154 Mackerer, C. R.,87 Mackey, L. N.,317, 325(107), 326(107), 327(107), 335(107) Mackler, B., 179,188(37),189,195(63), 203 (63),216 McLean, P.,46, 47(211), 81 McManus, I. R.,100, 103(65), 106, 107 (1051,1O9(65) MacMunn, C. A.,300, 315 McMurray, C. H.,41 McMurray, W. C., 62, 65(35), 67(35) McPherson, A., 10,24 MacQuarrie, R. A., 34, 36, 37(175), 42 (165),48(165), 49(165) Magar, M. E.,14,25(57) Mager, J., 286,287(386, 391)

Magni, G., 247 Mahler, H. R., 188, 189, 199, 217, 307, 308(59) Mahoney, A. J., Jr., 261 Major, J. P.,21 Makhlis, T.A., 70 Makino, N.,171, 172(402) Maley, G. F.,62,65(35),67(35) Malhotra, 0. P.,33, 35(160), 36(160), 37(174), 41(160) Mallet, A. I., 407 Malmstrom, B. G., 300, 301, 316(11), 317 (ll), 326, 328, 330,334(11), 335(108), 336(108, 171) Maltempo, M., 365 Malviya, A. N.,335 Mandula, B., 131 Mann, P.J. G., 260,261(248) Mann, T., 346 Mannervik, B., 132, 133, 139, 140, 141 (253, 254) Manocha, S.,300 Mansley, G. E.,322 Manzocchi, A., 248 Mapson, L. W.,129, 130(191), 138(191) Margoliash, E.,368,371(85), 372,376,377 (lll), 378(115), 380(111), 383(116) Margolis, J., 402 Margolis, S. A,, 70 Marklund, S.,393, 394(164) Markovich, D.S.,25,31(105) Martell, A. E.,407 Martin, R. B.,304 Martinez, J., 132 Marver, H.S., 150 Maskasky, J., 307, 314(60), 334(60), 338 (60), 343(60) Mason, H. S., 165, 166(373), 167(373), 170 (373), 171(373), 172(373, 4021, 176 Mason, T.L.,307, 308(58), 311, 312 Massey, V., 90, 91, 92, 93, 94(15, 24, 27, 29),95(27), 96(27), 97(1, 24, 29, 55), 98(1, 24,50, 55), 100(29, 32), 101, 103 (29,32),106(4,27),107(4,24,97,101), io8(4i), 109,111, ii2(24,55,ioi, 118, 150), 113(4, 24, 27, 29, 59, 137), 114 (27, 55, 150), 115(24, 137), 116(4, 24 27), 117(27, 1171, 120(83), 122(150, 154, 155), 123(154, 1551, 124(32), 125, 126, 133, 134(27, 1181, 135(29), 136

423

AUTHOR INDEX

(118), 137(1, 29, 59, 244), 138(29), 139(244), 140(244), 141(29), 146, 147 (284), 148(58), 170(1), 177, 153(23), 188(23), 200, 201(23), 203(23), 204 (23), 205(23), 223(23), 224(23), 235 (23), 236(23), 238(23), 246, 247(23), 248(23), 249(23), 250(23), 254, 282 Masters, B. S. S., 136, 150, 151, 154, 165 (337), 166, 167(245, 3371, 168(245, 386), 169(245), 170(245, 386), 172 (245,337,398) Masters, C. J., 365 Mathew, E.,11, 21, 22(52, 701, 42(70), 45(70) Mathews, C.T., 302,303(25) Matsubara, H.,310 Matsubara, S., 365 Matsurnura, Y.,311 Matthews, J., 106, 107(99), 108, 112(99), 114(99) Matthews, R. G., 90,91, 100, 103, 104(63), 114, 118(63, 156), 119(63, 156), 120 (63),121(63), 122(156), 146, 147(284), 282 Mauk, M. R., 372 Mauleon, P.,300 Mauzerall, D.,396 Mavis, R.D., 92, lOO(34) Maxwell, J. C.,319, 321, 322(142), 323 140), 337(117, 118), 339(117, 118), 340, 389 May, H.E.,168 Mayhew, S. G.,90, 91, 100, 123, 282 Mayr, M, 247. 249(199) Medina, A.,274,275(346),276(367) Mehler, A. H.,9 Meighen, E. A., 25, 26(99) Meigs, R. A,, 83 Meijer, A. J., 80,88 Melandri. B. A..2(28), 3 Meldrum, N. U.,92 Menon. K.M. J., 83 Mercer, W. D., 9, lO(45) Meriaether. B. P.,5. 11. 20(29), 21(29), 22(52, 70), 24, 39(29), 42(70), 44(90), 45(29, 70, 90, 185), 48(200) Merola, A. J., 180, 181(41), 312, 313(85) Mersmann, H., 69, 203 Mevel-Ninio, M.,269 Meyer, A.J., 88

Meyerhof, O., 2, 28(la) Michaels, G,B.,282 Michejda, J. M.,84 Middleditch, L. E.,55, 56(18), 57(18), 58(18) Middlemiss, D. N., 393,394(162) Midelfort, C.F.,9 Mihara, K., 151 Mildvan, A. S.,374, 385(100a) Millard, S. A., 106, 107(94) Miller, Z. B.,261 Mills, G. C.,48 Minakami, S.,176, 187, 188(56), 189(56), 190(56) Minchiotti, L.,100, 103(66), 143(66) Minnaert, K.,307, 325 Misaka, E.,106, 107(102, 1041, 122 Mitchell, C. H.,153 Mitchell, J. R.,153 Mitchell, P.,74, SO(155, 157), 81(157), 87 Mize, C.E.,129, 142 Mochan, E.,334,357 Moe, 0.A,, Jr., 126 Moir, N. J., 151 Molbert, E.,366, 367(48) Moleski, C.,334 Monod, J., 31 Monroy, G.L.,72 Montal, M., 72 Monteilhet, C., 264 Monty, K.J., 286,287(387), 288(404) Moore, E. C.,91, 92(8), 93(8), 99(8), 142 (23), 143(8, 231, 144(8), 145(8). 156 (9)

Moore, J., Jr., 23 Moran, T.,312 Moras, D.,9, 10(46), ll(46, 481, 18, 24 (47),29(47), 39(47, 48), 44(46, 47), 129, 141(188) Moreno, C. G., 274 Morgan. E.,245 Mori, M., 304 Morikofer-Zwex, S.,367, 379(63), 380 Morimoto, H., 372,373(92) Moroff, G.,141 Morrison, G.,83 Morrison, M., 176, 306(47), 307, 322 Morton, R.K.,264,269(300) Moss, T.H., 314, 334(94), 338(94), 356 Moyle, J.,74, 75(157), 81(157), 87

424

AUTHOR INDEX

Mozolovsky, A., 25 Muller, F., 282 Muller, M., 67, 69(100), 78(100) Muesing, R. A., 88 Muijsers, A. O., 308, 315, 316(101, 1021, 317, 318(111), 320(101, 102), 321(102, 132), 324,325,326(162), 335(157) Muiswinkel-Voetberg, H., 117, 118(160), 119(160), 124, 125, 128(181, 182) Mukherjee, B. B., 108 Muller, F., 90, 123, 126 Mulrow, P. J., 84 Munk, P., 108 Muraoka, S., 390 Murdock, A. L., 31 Murphy, M. J., 286, 287(397), 288(397), 289(397), 292(397), 295(410, 412) Murphy, T. A., 259 Murthy, P. S., 64, 65, 66(60), 68(60), 72

(60) Myer, Y. P., 319

N Nagahisa, M., 367 Nagai, Y., 286 Nagradova, N. K., 2(24), 3, 25(24), 30 (241, 48

Naiki, N., 286, 287(399), 288,292(399,424) Najjar, V. A., 274 Nakajima, H., 165 Nakamura, H., 217 Nakamura, M., 122 Nakamura, T., 286,295(402, 403) Nakanishi, K., 106, 107(102, 104), 122, 126 (104), 128(104)

Nakatani, M., 367 Namba, Y., 108 Narashimhulu, S., 166 Naslin, L., 267, 272 Nason, A., 274, 275(344), 277(337), 278 (356)

Navazio, F., 86 Nawa, H., 93 Needham, D. M., 2 Negelein, E., 301 Nelson, E. B., 150 Nelson, D. H., 65, 83, 84(211) NkmBthy, G., 32, 33(149) Nepokroeff, C. M., 88 Nesbakken, R., 130

Neas, G. C., 88 Neufeld, E. F., 52, 53(1), 54(1), 56(1), 58(1, 2, 3), 62(1), 65(30), 67(30), 69 (30), 70(30), 73(30), 76(30), 167

Newsholme, E. A., 47 Nicholas, D. J. D., 274, 275(346), 276 (367), 286, 287(401), 288, 292(401, 4221, 293(401)

Nicholls, D. G., 63,86,87(52) Nicholls, P., 237, 301, 320, 321(133), 334, 335(21), 346, 357, 386, 369(17), 370 (17, 721, 374(17, 721,377, 384(17), 385 (171, 389(17), 391(17), 395(17), 397 (17), 398, 400(175) Nieuwenhuis, F. J. R. M., 69, 74(119), 80 (1191, 214 Niluson, S., 384 Nise, G., 132 Nisley, S. P., 63, 66(37), 81(37), 86(37) Noeler, H. F., 5, 9(33), 20(33) Norling, B., 246, 249(188) Northrop, D. B., 141 Notani, G. W., 106, 107(110) Novogrodsky, A., 376, 377(111), 378(115), 380(111) Nurnberger, H., 368 Nygaard, A. P., 38, 44(179), 162, 264, 267 (280), 270, 271

0 O’Brien, P. J., 150, 152 Oda, T., 237 Oesper, P., 45 Oestereicher, G., 248 Oguchi, M., 48 Ogura, Y., 187, 372, 373(89), 376(89), 382 (89, 1071, 387(100), 388, 390(144)

Ohlsson, P. I., 365 Ohnishi, T., 187, 204, 215, 219, 235, 244, 253, 296, 346, 347(19), 353(19), 361 (19)

Ohno, H., 205 Ohta, Y., 366, 367(50) Okabe, K., 106, 107(96) O’Keeffe, D. H., 307,314(60), 315(93), 329 (931, 334(60, 94), 338(60, 941, 343(60)

Okunuki, K., 306(46), 307, 308(46), 274, 275(364, 365), 276, 310, 311, 315, 316 (1031, 323, 334

425

AUTHOR INDEX

Olah, G., 384 Olcott, R.J., 400 Oldham, S. B.,65, 83(84), 84(84) Olsen, K.W.,9, 10(46), ll(46, 481, 18, 24 (47), 29(47), 39(47, 481, 44(46, 471, 129, 141(188) Olson, E.J., 23, 45(81) Olson, J. A.,86 Olson, M. S., 110 Omdahl, J. L.,83 Omura, H., 274 Omura, T.,91, 150, 154, 165, 166(372), 167(372) Ondarza, R. N.,132, 133 Oosthuizen, C.,33 Oppenheimer, C.,363 Oppenheimer, N.J., 29 Orii, Y., 310, 311, 315,316(103), 320(103), 321(128), 324, 334, 336(134) Orlando, J. A., 66, 68, 71(114, 115), 72 (871, 79(115) Orme-Johnson, N. R., 179, 184(34), 185 (34), 186, 187(34, 461, 193(46), 215, 221 (46) OrmeJohnson, W. H.,83, 179, 184(34), 185(34), 187(34), 226(218), 253, 323 Orr, M.D.,143 Orrenius, S.,150, 152, 166 Orsi, B.A.,40 Osajima, Y.,274 Osborn, M.,366 Osenga, G., 222, 225(150) Oshino, N.,151, 365 Ostroumov, S. A.,69,74(121) Ota, A.,274,275(364) O’Toole, M.C.,308,314,315(93), 317(65), 318(65), 322(65), 323(65), 325(65), 328(65), 329(65, 93), 337(65), 338(65) Otsuka, J.,366 Otsuka, K.,108 Otto, J., 311 Ovadi, J.,24,25 Overberger, C.G.,379 Owen, C. S.,320, 322(124), 323(124), 325 (124),326(124), 331(124), 332(124) Ozawa, T.,234, 335

P Packer, L., 259,261, 262(262) Pain, R.H., 368

Pajot, P., 264,265,269 Palmer, G., 90,92,95,97(l,55), 98(1, 55), 100(32), 101, 103(32), 112(55), 113 (591, 114(55), 120(83), 124(32), 137 (1, 591, 170(1), 205, 322,324, 330, 335 (1531,336(153), 375 Paltauf, F., 151 Panebianco, P., 219 Paneque, A.,274,276(345) Paniker, N.V.,131 Panov, A. V., 70, 88(129), 212 Papa, S.,63, 73, 86(50), 87(53, 54, 55) Paradies, G.,70,88(129), 212 Park, J. H.,3, 5, 11, &(29), 21(29), 22 (52, 701, 23, 24, 28(13), 29(13), 31, 38(13), 39(13, 291, 40(13), 42(70), 44 (13, QO), 45(29, 70, 81, 90, 1851, 48 (200) Park, R., 84 Parker, D.J., 20, 21(66), 30 Parkhouse, R. M. E., 132 Parkhurst, L. F.,333 Passon, P. G.,165 Passoneau, J. V.,47 Patchornik, A., 379 Paul, K.G., 365 Pauling, L.,364 Payne, M.,33, 35(160), 36(160), 41(160) Payne, W.J., 274 Pazur, J. H., 101 Pearson, R.G.,375 Peck, H.D.,Jr., 279, 281, 282, 284,285, 286(372), 288,295(396), 297 Peczon, B. D., 33, 41(161) Pederson, T. C., 165, 166(369, 374), 167 (374), 168(374), 169(374) Peisach, J., 368, 369, 370(77, 781, 376(77), 380, 381(126b), 390(126b) Pelley, J. W.,106, 107(92), 108(92) Perham, R. N.,5, 24, 21(30, 34), 45, 104, 118(86), 119(86),120, 121(86) Peron, F. G., 84,88(225) Persson, B.,246,249(188) Perutz, M.F.,349,372,373(91) Pesch, L. A.,70 Petering, D.H.,360 Peters, J. M., 92,97(28),104(28), 114(28), 127(28) Peterson, J., 70

426 Pette, D., 48 Pettit, F. H.,106, 107(92), 108(92), 110 Pfennhger, O., 33, 35(160), 36(160), 41 ( 160) Pfleiderer, G., 24 Pharo, R. L., 70, 76(127), 77(127), 189, 190(59), 195(64), 198(64), 205 Phillips, A. H.,91, 166(11), 167(11), 168 (11) Pihl, A,, 20, 368 Pillai, P., 2 Pincus, G., 83 Pinsent, J., 366, 367(45, 46) Plaut, G.W.E., 86,87 Plumley, H.,130 Pnafili, E.S.,155 Pocker, Y.,407 Podack, E.R., 88 Poff, K.L.,239,240(178) Pogson, C.I., 41 PolgBr, L., 21, 22(73, 741, 23, 43, 45(73, 74) Poole, B., 365 Popowsky, M.,202 Porque, P. G.,143,144(275) Porter, J. W.,88 Porterfield, U.T., 388 Postma, P. W.,79 Poulsen, L. L., 106, 107(100), 153 Poyton, R. O.,307,308(58), 311 Prabhakararao, K., 286, 287(401), 288, 292(401, 422), 293(401) Prados, R., 304 Prager, G., 255 Prakash, O.,274,277,278,297(341) Presswood, R. P., 170, 286, 287(394), 290 (3941,291(394) Preabindowski, K.S.,313 Price, N.C.,33 Price, V. E., 366 Pronk, J., 120, 148(163) Prough, R. A., 151,166,169,172 Psychoyos, S.,83 Puchwein, G.,32 Pullman, M.E.,72, 88 Pulsinelli, P. D.,372,373(91) Pupillo, P., 2(28), 3 Purvis, J. L.,84 Pyfram, H.T., 376

AUTHOR INDEX

0 Quagliaridllo, E., 63, 73, 87(53, 54), 88 Quastel, J. H.,260, 261(248)

Rabinowitz, J. C., 243 Racker, E., 13, 28, 38, 39(54), 40, 41, 42 (196), 44, 66, 179, 180(36), 181(36), 183, 184(36), 186, 187, 201(35), 214, 239,244,246,302 Radcliffe, B. C.,274 Radda, G.K.,33 Rafter, G. W.,2(10), 3 Ragan, C. I., 78, 179, 180(36), 181(36), 182, 183, 184(36), 188, 187, 190(43a, 133), 201(35), 204, 214, 217, 218(133), 219( 1311, Raijman, L., 46 Rall, T. W., 92 Ramasarma, T., 247 Randall, D.D.,106, 107(92), 108(92) Ranney, H.M.,372,373(91) Rao, N. A,, 179, 188, 189, 195(63), 203 (631,216 Rapkine, L., 2,20(5) Rasmussen, 0.L.,302 Rawitch, A. B.,101 Ray, D.K.,317, 318(112) Ray, G. S.,334, 335(185), 336(185), 346, 348(14), 352(17), 353(14) Recheigl, M., 365 Redfield, A. G.,357 Redline, R., 148, 151 (284a) Redman, C.M., 365 Reed, D.W.,165 Reed, G.,350, 360 Reed, G.H., 372, 376(88) Reed, J. K.,106, 107(95), 110, 116(95), 117(95), 118(95), 119(95), 139(95) Reed, L. J., 92,93, 106, 107(92, 99, 1091, 108(19, 43, 92),109(03), 110,112(99), 114(99,log), 126(126) Reichard, P., 91,92(8), 93031, 99(81, 142 (23),143(8,23,261,2621, 144(8, 275), 145(8), 156(9) Reiske, J S., 310 Rendina, G.,260,261(254, 255) RBtey, J.,252

427

AUTHOR INDEX

Richardson, S. W.,306(50), 307, 308(50) Righetti, P.,225, 230(157), 231,234 Rikihisa, T.,39,45(181) Ringler, R. L.,176, 187, 188(56), 189, 190 (56),238, 257, 258(226) Riordan, J. F., 213 Rippa, M.,270 Rider, J. L.,264 Rivas, J., 274, 276(345) Robberson, B., 366, 367(41), 370(41), 371 (41) Robbins, P. W.,279 Roberts, K.D.,83 Robertson, A. M., 69 Robins, R. G.,302,303(25) Robinson, J., 65, 85(86) Robinson, J. R., 108 Rocca, E., 177, 217,254(16) Rodkey, F. L.,113 Rodman, H.M.,402 Rogers, M. J., 148, 151(284a), 154, 156 (341, 342), 161(341, 342, 344), 161 (341,,342) Rogers, L.A., 285 Rogers, W.I., 261 Rolleston, F. S.,47 Ronchi, S.,2!2, 44(76), 100, 101, 103(62, 661, 104(61,62), 105(62), 120(61), 123 (611, 143(66), 144(62), 145, 146, 147 (284) Roos, D., 73 Roper, M.,150 Rose, I. A., 47 Rosemeyer, M.A,, 131 Roskn, S.,331 Rosenthal, O.,83, 91 Rosenthal, S.,152,166 Rosing, J., 74 Rosman, J., 310 Ross, E.,66 Rossi, C., 78, 190(84), 202, 222, 225(150) Rossi, E.,246, 249 Rossi, F., 222, 225(150) Rossi-Fanelli, A.,349 Rossmann, M. G.,9, 10(46), ll(46, 48, 51), 12(51), 18,24(47), 29(47), 39(47, 48),44(46,471,129, 141(188) Rothschild, H. A.,261 Rotilio, G.,304 Roughton, F.J. W., 388

Rouslin, W.,312 Roussos, G.G.,274,278(356) Rubin, E.,81(176), 402 Rubin, M.S.,307,308(57),311(57) Rueger, D. C., 170, 286, 287(394), 290 (3941,291(394) Rumack, B.H.,150 Rumen, N. M., 374 Rutter, W.J., 9 Ruzicka, F.J., 245,297 Ryan, D.,149, 152 Ryan, K.J., 83 Ryback, G.,252 Rydstrom, J., 53, 58(8), 61(8), 64, 66, 68, 69(8, 68, 69, 70, 71, 721, 70(8, 67, 70, 72, 107), 71(8), 72(67), 75(67, 68, 69, 70,711,76(8, 67,68,70,71,721, 77(67, 68, 69, 70, 71, 72), 78(70, 1071, 87, 88 (70,71,129, 1301,207,212 Rytka, J., 272

S Sabeson, M. N., 11 Sabo, D.,66, 72(87) Sacktor, B.,258, 259(229, 233, 234) Sadana, J. C.,274,277,278, !297(341) Sadasivan, N.,305 Sagers, R. D.,106,107(113), 1W113) Saggerson, E. D.,47 Sajg6, M.,5,9(33),20(33) Sakai, H., 129 Sakamoto, Y.,150 Sakurai, Y.,108,109,llO(132) Salach, J. I., 222, 235, 247, 249(200), 258 Salhanick, H.A.,65 Salmon, D.M.W., 46 Saltzgaber, J., 312 Salvenmoser, F., 67, 69(100), 70(98), 78 (100) Samejima, T., 366,367(50),375 Samson, L.,371 (85),372 Samuelsson, B.,83, 168 Samuilov, V. D.,69, 74(121) Sanadi, D. R., 64, 65, 67, 68(31), 70, 72 (61), 76(127), 77(127), 79, 91, 92, 97 (26, 281, 104(28), 106, 107(25), 108, 113, 114(26, 28), 116(116), 117(151), 127(28), 189, 195(64), 198(64), 205 (64)

428 Sandberg, H. E., 369, 370(74, 791, 372(74, 791, 376(79) Sandborn, B. M.,249 Sanders, E., 91 Sands, R. H., 98, 113(59), 137(59), 235, 244, 330 Sani, B. P., 67 San Pietro, A., 53, 54, 55(11), 56(11), 58 (5), 59(11), W11) Santema, J. S., 55, 56(19), 57(19), 58(19), 59(19), 62, 80(19), 106, 107(114), 114 (114) Sarkar, N. K., 188, 189(57), 199(57) Saronio, C., 324, 336(155), 389, 390(150), 391(150) Sasame, H. A., 153 Ssto, R., 66, 72(89), 730391, 76(89), 151, 154, 288, 287(400), 288, 292(400, 421, 423, 424), 293, 294, 295(402, 403) Satoh, K., 65,69(78), 72(78) Sauer, L. A., 84 Savage, B., 26 Savage, N., 94, 98(54) Schachman, H. K., 25,26(99), 34(112) Schacter, B. A., 150 Schatz, G., 66, 167, 183, 214, 307, 308(58), 311, 312 Schejter, A., 377, 378(115) Scherz, B., 367 Schevitz, R. W., 10, 24 Schiff, J. A., 279, 280, 282, 283 Schindler, F. A., 390 Schlessinger, J., 34 346s 348(ZZ)3 351(n)3 Schleyer, (n), 353(20, '22) Schmid, D., 25 Schneider, W. C., 67 Schoenhard, D. E., 286 Schollmeyer, P., 63,81(38), 86(38) Scholz, R., 88 Schonbaum, G. R., 361, 365, 369(17), 370 (17), 372, 373(98), 374(17), 377, 378 (117), 379(118b), 2"25), 381(125, 126b), 382(125), 384(17)9 385(17, 100at llsb, 125), 387(118b), 389(17), 390 (lola), 391(17, lola), 392, 393(101a, 159), 394(101a), 395(17, 159, 159a), 397(17, lola), 398(101a), 399(101a, 118b), 400, 401, 402(118b, 159), 403 (118b), 404, 405

AUTHOR INDEX

Schramm, M.,44 Schrauzer, G. N., 396 Schroeder, W. A., 366, 367(41), 370(41, 63a), 371(41, 63a), 372(63a), 377, 378 Schultz, J., 378 Schuman, M.,90,91, 101,282 Schuster, I., 25,26,31(101), 32,33(101) Schutte, H. R., 368 Scott, E. M.,93, 94(40), 138(40), 139(40), 140(40), 164 Scouten, W. H., 106, 107(105), 109 Seamonds, B., 346, 351(21) Searls, R. L., 92, 97(26, 281, 104(28), 106, 107(25), 108, 114(26, 281, 116(116), 127(28) Sebald, W., 311 Segal, H. L., 39, 40 Segel, I. H., 129,132(193), 138(193) Sehested, K., 302 Seibl, J., 252 Seifried, H. E., 151 Sekuzu, I., 306(46), 307, 308(46), 323, 334 Sellinger, 0. Z., 259 Sels, A. A., 347 Seng, R., 222 Setlow, B., 148, 151(284a) Seubert, W., 88 Severina, I. I., 74, 75(148) Seydoux, F., 33, 35(160), 36(160), 41(160) Shah, P. C., 106, 107(109), 114(109) Shakespeare, P. G., 307, 308(59) ShaltiB1, S., 20, 22 Shaw, D. C., 11, ZZ(52) Shelton, J. B., 366, 367(41), 370(41), 371 (41) Shelton, J. R., 366, 367(41), 370(41), 371 (41) Shepley, K., 269 Shibata, H., 150 Shibata, Y., 25 Shifrin, s.,28 Shimakata, T.,151 Shin, B. c.,48 Shin, M., 54, 55, 58(12), 66(12) c. E**47 Shusterf L.*59f 150 Siege19 L. M.9 170, 286, %7(390, 394, 3979 412), 288(390, 397, 404), 289(390), 290 (394,413,414,415), 291(414), 292(390, Shonkj

AUTHOR INDEX

397, 415), 293(390), 294(414), 295(410, 412) Sies, H., 130, 365 Sih, C. J., 83 Simard-Duquesne, N., 69 Simon, A. M., 265 Simon, I., 34 Simplicio, J., 303 Simpson, E. R., 84 Simpson, R. J., 2(25), 3 Singer, T. P., 69, 78, 106, 107(91), 109(91), 110(130), 176, 177, 184, 186(23), 187, 188(19, 23, 561, 189(56), 190(56, 83, 841, 200(15, 191, 201(14, 15, 19, 231, 202, 203(19, 2 3 , 204(19,23), 205(19, 23), 205(19, 22, 23, 88), 206(19, 221, 214, 216(116), 217, 219, 220(136), 221 (1%) 222, 22303, 25, 140, 144), 225 (23, 25, 1441, 226(158), 230(158), 234 (25, 158), 235(23), 236, 237(15), 238 ( 2 3 , 240(15), 245, 246, 247(15, 20, 23, 25, 189), 248(23, 24, 25, 26, 249(23, 25, 26, 197, 199, 200), 250(23, 25, 195, 196, 1971, 251(197), 252(197), 253(1&1), 254(15, 16, 251, 255, 257, 258(=), 260, 261(%7, 254, 2557 256)* 262(227, 2621, 263(266), 284, 2709 271, 272(312), 273 Singh, A. P., 79 Singh, J., 275 Sivak, A., 261 Sjoberg, B. M., 93, 119(47), 143(47), 144 (47) Skulachev, P., 214 Skulachev, v. P., 69, 74(120, 121), 75(120, 146, 147, 1481, SO(146, 147) Skvaril, F., 367 ‘later’ E’ “’ 28’ 31’ 33(144)’ 34(144)9 41 (159), 63, 74,86(39, 49), 176, 187, 237, 238(172), 251, 307, 390 Slein, M. W., 2, 3(8), 48 Slencaka, W., 56, 57(20), 63, 66(20), 81 (20), 86(20) Sloan, D. L., 32, 33(155) Slonimski, P. P., 217, 272 Sluse, P. E., 88 Smiley, I. E., 24 Smillie, L. B., 371(861, 372 Smith, C. M., 40, 41, 42, 45(194), 49(194), 110

429 Smith, D. W., 369, 370(76), 372(76), 374 (76) Smith, G. D., 26,34(112) Smith, J. E., 132, 142 Smith, M. H., 300 Smith, M. L., 307, 314(60), 334(60), 338 (60), 343(60) Smith, T. E., 40 Smith, W., 168 Smythe, G. A., 307, 314(60), 315, 334(60, 94),338(60,94), 343(60) Snyder, H., 378 Soling, H. D.,82 Somlo, M., 269,272(306) Sone, N.,260 Sordahl, L. A., 189, 195(64), 198(64), 205 (64) Sorger, G. J., 274 So&, S., 20 Sosfenov, N. I., 366 Spallhols, J. E., 321, 322(142) Spats, L., 100, 148, 151(284a), 154(74), 155, 156(74), 161(74), 166(74), 167(74) Spencer, D., 274,277(337) Spencer, R. L., 100, 103(67) Speranaa, M. L., 100, 103,143(66) 41(161) Spivey, H. O., 3, Sportorno, G. M., 25,26 Spyridakis, A., 265,268(289) Sreenivasan, A., 260 Srere, P. A., 88 Srivastava, s. K., 131, 132 Staal, G. E. J., 94, 125, 131, 137(51), 139 (51), 140(51) Stachiewicz, E., 272 Stadtherr, L. G., 304 Stadtman, T. C., 143 Stallcup, W. B., 26, 35, 36(113), 37(113, 172) Stanbrough, E. C., 143 Stanbury, J’, 315, 316(100), 322, 323(100) Stansell, M. J., 366,382(39) Stark, G. R., 383 Staudt, H., 153 Steele, W., 244 Stein, A. M., 65, 86(75), 109, 110, 114, 117, 118(153) Stein, J-H., 86,109, 1 1 4 3 118(153) Stellwagen, E., 92, 100(34)

430 Stempel, K. E.,180, 181(40), 191, 194(40, 67), 195, 196, 197, 198(40), 199(68), 203(68), 206(40, 42), 207(40, 42), 216 (40) Stern, K. G., 363,366,391,397(160) Stevenson, P. M.,65, 85(86) Stiggall, D.L.,178,297 Stockell, A.,31 Stolzenbach, F. E.,30, 54, 55(11), 56(11), 59(11), 66(11) Stotz, E.,310,311(69) Straub, F.B.,106, 107(90), 189 Strength, D.R.,261 Strickland, S., 91 Strittmatter, P.,91, 100, 124, 148, 151(9, 17, 284a), 15407, 74, 307, 308), 155 (347), 156(74, 307, 341, 3421, 157 (3501, 158(354), 159(308, 351), 160 (308,354, 355), 161(74, 341, 342, 343, 344, 3501, 162(171, 341, 342), 163 (171, 352), 164(347, 349, 352, 359, 360), 165(347), 166(74), 167(74), 168 (360) Strobel, H. W.,149, 153, 169 Stromberg, C., 317,335(108), 336(108) Strom, R.,237,247 Strother, G.K.,395 Stryer, L.,93,349,364 Studier, M.H., 377 Sturani, E.,100 Sturtevant, J. M., 33, 34(156), 268, 269 (300) Stynes, D.V.,302 Stynes, H.C., 302 Su, G., 117, llS(l61) Suematsu, T., 108 Suggett, A., 368, 369, 388, 389, 395(75), 397(75) Sugita, Y.,317,318(110), 349 Sullivan, P.A.,90,91,282 Sulmovici, S.,83 Sumner, J. B.,38,440791,366 Sun, F.F.,306(48), 307, 308(48), 313 Sund, H.,366, 367(48) Susheela, L.,247 Suter, H., 365 Sutin, N.,375 Suzuki, I.,281, 282(375), 284 Suzuki, K.,2(18, 20), 3, 4(18), 9(18), 19, 25, 26, 27(103)

AUTHOR INDEX

Suzuki, T., 372, 373(92) Swartz, M.N.,62,65(31) Sweat, M.L.,83, 84(210) Sweetman, A. J., 64, 65, 68(63), 72(63), 73(63) Swoboda, B. E. P., 98, 113(59), 137(59) Symons, R.H., 267 Szabolcsi, G.,23, 31 Szarkowska, L.,258 Szorhyi, E.,2(14), 3 Szumilo, T., 273 Szymona, M., 273

T Tagawa, K., 54, 55(12), 56(12), 66(12) Tager, J. M.,63, 73, 81(40), 82, 86(39, 40,49,191),87(53,54),88 Takahashi, E.,286, 295(395) Takahashi, H.,274, 277(337) Takahashi, Y.,335 Takano, T., 371 (85),372 Takemori, A.,259 Takemori, S.,237, 245(173), 306(46), 307, 308(46),334 Takesue, S., 154, 165, 166(372), 167(372) Tallan, N.H.,83 Tamaoki, B.,83 Tamura, G., 274, 286(359), 295(359, 405) Tanaka, M.,100 Tanaka, N.,108 Tanford, C., 367 Taqui Khan, M.M., 407 Tarr, H.L.A., 92 Tasaki, A., 366 Taube, H., 303 Tauber-Finkelstein, M.,22 Taylor, E.L.,39 Taylor, J. F.,24 Taylor, P. L.,85 Taylor, W.E.,150 Teale, F.W.J., 348, 380 Tedeschi, P.,28 Teeter, M.E.,205 Teipel, J., 34, 35(164) Teixeira da Cruz, A., 64, 69(68, 691, 70 (67), 72(67), 75(67, 69), 76167, 68, 691,77(67,68,69),207 Telegdi, M., 25 Teller, D.C.,25

431

AUTHOR INDEX

ter Welle, M. F., 63, 72(47), 73(47), 81 (47) Tepley, L. J., 189 Testolin, G., 222, 225(150) Thelander, L., 91, 92(8), 93(8), 98(31), 99(8, 311, 100(31), 101(31), 103(31), 104, 142, 143(8, 262, 2631, 144(8, 38), 145(8, 31), 156(9) Theorell, H., 162, 200, 356, 364, 366, 369, 370(6), 385, 388, 400(143) Thomas, J. O., 23 Thompson, T. E., 142 Thor, H., 150 Thorgeirsson, S. S., 153 Thorn, M. B., 247 Thornber, J. M., 109 Thorpe, C., 123 Thurman, R. G., 88 Tiesjema, R. H., 317, 318(111), 319, 324, 325, 326(162), 335(157) Tietre, F., 130, 143(202) Tisdale, H., 205, 222, 225, 226(158), 230 (158), 234(158), 248(158), 255 Tolbert, N. E., 274 Tomizawa, H. H., 130 Topali, V. P., 69,74(120), 75(120) Topper, D. L., 407 Toren, D., 83 Torii, K., 279, 372, 373(89), 376(89), 382 (89, 107) Torndal, U. B., 65, 73(166), 78, 207, 212 Tosi, L., 264 Tottmar, S. 0. C., 190(133), 217, 218 (133) Tove, S., 287(412), 288,295(412) Trentham, D. R., 21, 23, 28, 35(69), 37 (83, 1171, 39, 40(83, 117), 41(69, 189, 190), 42(116, 117), 43(83, 1171, 44, 45 Trudgil, P W., 91 Trudinger, P. A., 286, 295(392) Triiper, H. G., 285 Tsai, C. S., 117 Tsai, P., 150 Tsao, M. S., 302 Tsernoglou, D., 10 Tsofina, L. M., 69,74(120), 75(120) Tsou, C. L., 222, 223(146), 226(146), 246 (146) Tsuchihashi, M., 367 Tsudzuki, T., 325, 326(163), 327

Tu, S. C., 101 Tubbs, P. K., 88, 273 Tucker, A., 300 Tung, T., 256 'Turini, P., 255 Turkki, P. R., 262 Turner, J. F., 143 Tyler, D. D., 186,203, 262 Tysarowski, W., 272 Tyson, C. A., 172 Tzagoloff, A., 307, 308(57), 310, 311(57), 317

U Ueda, T., 91 Uesugi, I., 274 Ullrich, V., 152, 153 Urnes, P., 372, 373(95) Urry, D. W., 319 Uigiris, V. I., 65

v Vanngard, T., 317, 330, 335(108), 336(108, 171) Vainshtain, B. K., 366 Vallee, B. L., 213 Van Ark, G., 332 Van Buuren, K. J. H., 308, 315, 316(101, 102), 320(101, 102), 321(102, 132,133), 335 van Dam, K., 63, 69, 72(47), 73(47, 481, 74(119), 80(119), 81(47), 86(49), 214 Van Demark, P. J., 260 van den Brock, H. W. J., 53, 55, 56(19), 57(9), 58(9), 59(9, 19), 60(9), 62, 80, 143 Van der Hoeven, T. A., 153, 165, 166 (3701, 170(370) van de Stadt, R. J., 69,74, 80(119), 214 Van Drooge, J. H., 332 Van Eys, J., 3, 28(13), 29(13),.38(13), 39 (13), 40(13), 44(13) Van Gelder, B. F., 307, 308, 315, 316(101, 1021, 317, 318(1111, 319,320(101, 102), 321(102, 132, 133), 324, 325, 326(162), 331(139), 332(139), 333(139), 335 (157) van Haefen, H., 63, 86(45)

432 Van Heerikhuizen, H., 187 Vanko, M., 260 Van Lis, M. J., 33,41(159) Vanneste, W. H., 315, 316(98), 317(98), 320(98), 322(98) Varandani, P. T., 130 Varshavsky, Y. M., 14,3166) Veech, R. L., 46, 79, 81(175), 82(175), 86(175) Veeger, C., 63, 56, 58(19), 57(9, lg), 58(9, 191, 59(9, 191, 60(9), 62, 80(19), 92, 94(24, 27), 95(27), 96(27), 97(24), 98(24), 106(27), 107(24, 97, 106, 114), 111(24), 112(24, 108, 1181, 113(24, 27), 114(27, 114), 115(24), 116(24, 27), 117(27, 157), 118(157, 160), 119 (1601, 120, 122(154), 123(154), 124, m ( i 7 2 , 173, 174, 1751, i26(174, i79, 180, 181, 1821, 131, 133, 134(27, 1181, 136(118), 137(51), 139(51, 157), 140 (51), 143, 148(163), 222, 224, 226 (218), 236(149), 237, 238(149), 251, 252, 253 Vega, J. M., 274, 276(342) Velick, S. F., 3, 9, 19, 29, 30(125), 31 (12), 32(127), 33(155), 34(156), 39, 40(60), 41(60), 42(60), 44(60), 45 184, 194), 48(60), 49(194), 91, 92, 151(9) Vennesland, B., 39 Vennesland, J. W., 92 Vermilion, J. L., 165, 166(371), 167(371) Vernon, L. P., 188, 189(57), 199(57) Vignais, P. V., 84 Vinogradov, A., 248 Visser, J., 116, 117(157), 118(157), 120, 124, 125,(175), 126(179, 180, 1811, 139 (1571, 148(163) Vitols, E., 143, 216 Vladimirova, M. A., 69, 74(120), 75(120) Voetberg, H., 116, 117(157), 118(157), 120, 125, 126(180), 139(157), 148(163) Vogel, O., 106, 107(107), lOS(l0t) Voigt, B., 31 , Volkstein, M. V., 25,31(105) Volpe, J. A., 306(52), 307, 308(52), 313 (52), 315(52), 316, 317(52, 651, 318 (52, 65), 319, 321, 322(65), 323(65, 1401, 325(65), 328(65), 329(65), 333, 337(65, 118), 338(65), 339(118), 389

AUTHOR INDEX

von Ellenrieder, G., 32 von Wartburg, J. P., 367,379(63), 380(63) Vore, M., 152 Vorona, M. K., 48 Vyas, S. R., 189, 195(64), 198(64), 205 (64)

W W d a m , F., 150 Waentig, P., 367 Wainio, W. W., 306(49), 307, 308(49), 310, 319, 322(49), 324 Wainwright, T., 286, 287(398), 288, 292 (398, 420) Wakil, S. J., 88, 151 Walajtys, E. I., 110 Walker, G. C.,274, 275 Walker, W. E., 222, 235 Wallace, W. J., 319, 321, 323(140), 337 (1171, 339(117), 340, 389 Wallenfels, K., 28, 33 Waller, H. D., 130, 131 Wang, L., 109 Wang, T. Y., 222, 223, 226, 246(146) Wang, Y. L., 222, 223(146), 226(146), 246(146) Warburg, O., 2, 301, 315, 363, 364(3) Wartofsky, L., 91, 14303 Warnarman, P. M., 19,21, 28(61) Wassink, J. H., 55, 56(19), 57(19), 58(19), 59(19), 80(19), 108, 107(114), 114 (114) Watari, H., 190(83), 202,267, 372, 373(922) Waterman, M. R., 360 Waters, W. W., 407 Watson, H. C., 9, 10(45), 19, 21, 28(61), 35, 364, 372 Webb, L., 10 Weber, F., 131 Weber, K., 366, 367(48) Weber, M. M., 110, 168, 169(388) Wedding, R. T., 106, 107(100) Weenen, J. H. M., 39 Weil, J. A., 304 Weinbach, E. C., 66 Weinhouse, S., 46, 49(214) Weiss, L., 88 Welch, M., 85 Welinder, K. G., 371(86, 871, 3?2

AUTHOR INDEX

Wells, I. C., 261 Welton, A. F.,165, lsS(369) Wendel, A., 130 Wendell, P. L.,133 Wenske, G.,63, 86(45) Werner, A,, 303 West, C.A.,66 West, S., 149, 152 Wever, R.,315, 316(102), 320(102), 321 (1021, 332 Wharton, D. C., 275, 301, 306(45), 307, 308(45), 310, 312, 322, 324, 325, 327 (53), 330(53), 335(153), 336(153) White, H.B., 111, 39 White, R. P., 378, 379(118b), 385(118b), 387(118b), 399(118b), 401, 402(118b), 403(118b), 404, 405 Wiberg, K. B., 407 Widger, W. R.,78, 182, 190(43a) Wieland, O.,88 Wikstrom, M. K. F., 215 Wilken, D.R., 133, 263 Wilkins, R. G.,303 Williams, C. H, Jr., 91,92,93, 94(29, 35, 36), 97(29), 98(30, 501, 99(30), 100 (29, 33, 36, 37), 101, 103(29, 631, 104(33, 35, 61, 62, 63), 105(33, 62, 85), 106(36), 107(36, 103, 108), 113 (29, 59), 114(108), 118(63, 156), 119 (33,35, 63,85, 1561, 120(61, 631,121 (631, 122(156), 123(61), 129, 135(29, 601, 136(60), 137(29, 59, 60, 244), 138 (291, 139(244), 140(244), 141(29, 351, 143(36), 144(36,62), 145(30), 146(30), 147(284), 148(58), 166(10), 167(10, 245), leS(l0, 245, 3861, 169(245), 170 (245), 172(245) Williams, G.R., 263 Williams, J. N.,Jr., 260, 334 Williams, J. R.,81 Williams, R.J. P., 317,318(109), 351,369, 370(76), 372(76, 811,374(76, 811, 390 Williamson, D. H.,46, 82, 86(185, 186) Williamson, J. R., 46, 47(215), 48(219), 110 Willms, B., 82 Willms, C. R., 108 Wilmarth, W. K.,302, 304 Wilson, D.F.,215,302, 320,321, 322024, 138), 323(124, 137), 325(124), 326(124,

137, 138, 1631, 327, 328(1379, 331(124, 138), 332(124, 168),333(137,138,168), 335(138), 346, 351(21) Wilson, J. E., 100, 101(64), 103(64), 109 (64), 117, 118(161) Wilson, L. D.,83, 84(211) Wilson, L.G.,91,133(7), 143(7), 279 Wilson, M.T.,315, 316(99), 323(99), 324 (99),335,336(99,192),389 Wilting, J., 335 Winell, M.,133 Winter, D. B., 235, 244(164), 248, 253 (164), 296 Wiss, O., 131 Witkowski, P.E.,262 Wit-Peeters, E. M.,88 Woenckhaus, C., 24 Wold, F.,100,103(67) Wolf, B., 109 Wolman, Y., 379 Wolny, M., 9 Wonacott, A. J., 10, 19, 20 Wong, D.,204 Wong, S.H., 83,84(211) Wood, P. M.,303 Woodin, T. S., 129, 132(193), 138(193) Woodrow, G.V.,111,360 Woodward, H.E.,260,261(248) Woolf, A. A.,384 Worthington, D. J., 131 Wren, A., 106, 107(101), 112(101), 117 ( 117)

Wiithrich, K.,267, 357 Wyman, J., 31 Wyss, S.R.,367

Y Yagi, K.,176,234,335 Yamafuji, K.,274 Yamamoto, H.,350, 360, 369, 370(80), 372(80), 376(88) Yamamoto, T., 311 Yamanaka, T., 274,275,276 Yamasaki, I., 122, 169 Yang, C.S.,198,311,312(78) Yang, J., 366,367 Yang, P. C.,310 Yang, S. T.,25, 30(108), 48(108) Yasunobu, K.T.,100

AUTHOR INDEX

Yeghiayan, A,,372,373(96) Yike, N. J., 64, 66(64, 651, 68(64, 65), 69(65), 72(64, 65), 73(65), 76(65) Yonetani, T.,306(44,46), 307,308(44,46), 320, 330, 331(174), 334, 335(185), 336 (1851, 346, 347(19), 348(14, 26, 27, 28, 29, 30, 31, 32, 33, 37, 38, 39), 349 (26, 28, 30, 32, 39, 461, 350, 351(21, 22), 352(17, 22), 353(14, 16, 18, 19, 20, 22, 34), 355(34, 361, 356(55), 357 (17,38), 359(39), 360(27, 31), 361(17, 19), 364, 369, 370(80), 372(80), 376 Yoneyama, Y., 317, 318(110), 349 Yong, F. C.,306(51), 307, 308(51), 310 (511, 319 Yoshida, K., 372,387 Yoshikawa, S.,320, 321(128), 336(134) Yoshimoto, A., 286, 287(400), 288, 292 (400, 421, 423, 424), 293, 294, 295 (402, 403) Yoshiaawa, K.,108 You, K. S.,179, 193(32), 214, 226(32) Yu, C.,312 Yu, L.,312

Z Zahler, W. L., 313 Zakim, D., 109, llO(130) Zanetti, G.,92, 94(36), 97, 98(30, 501, 99 (30), 100(36), 103(36), 106(36), 107 (36), 143(36), 144(36), 145(30), 146 (30), 147(284), 148(58), 222,225(150), 216, 247(191) Zapponi, M. C., 22,44(76) Zatman, L.J., 53 Zawodsky, P.,14, 25, 26, 31(56, 105) Zerfas, L. G.,263 Zeszotek, E.,222 Zeylemaker, W. P., 222, 224(149), 236 (149), 237(149), 238(149, 1721, 251, 252, 253(149) Zherebkova, N. S., 153 Zidoni, E.,305 Ziegler, D. M.,153, 154, 165(337), 167 (3371, 172(337), 224, 236(153, 1541, 239 Zumft, W. G., 274 Zuurendonk, P. F.,308, 320, 321(132)

Subject Index A

glyceraldehyde-3-phosphate dehydrogenase and, 30 Absorption spectra small NADH dehydrogenases and, 196 adenylyl sulfate reductase, 282, 284, 285 transhydrogenases and, 59,69 catalase, 372-374, 381, 382, 397 ubiquinone reductase and, 181, 184, 186, cytochrome b?, 265, 267 215-216 cytochrome b, reductase, 155-156, 157Acetylpyridine adenine dinucleotide 158 phosphate, sulfite reductase and, cytochrome c oxidase, 315-319, 322,323, 288, 290, 292 327 Achromobacter fischeri, nitrite reductase, cytochrome c peroxidase, 351,353,354 physical properties, 277-279 glutathione reductase, 95, 135-136, 137Active site, lipoamide dehydrogenase, 105 138 Acyl hydrazides, catalase and, 379 a-glycerophosphate dehydrogenase, 257 Acyltransferase activity, glyceraldehydelipoamide dehydrogenase, 118, 122, 123, 3-phosphate dehydrogenase, 44-45 126 Adenine nucleotides nitrite reductase, 278 glyceraldehyde-3-phosphate dehydrosmall NADH dehydrogenase, 193, 194 genase and, 2, 45, 46, 48 succinate dehydrogenase, 232, 233 transhydrogenase and, 70 sulfite reductase, 288, 289,291, 293 Adenosine, transhydrogenase and, 71 thioredoxin reductase, 98 Adenosine diphosphate ubiquinone reductase, 183-184 choline dehydrogenase and, 262,263 Acatalasemia, form of, 367 lipoamide dehydrogenase and, 125 Acetaldehyde, catalase and, 391-392, 406 NADH dehydrogenase and, 207 Acetate succinate dehydrogenase and, 250 catalase and, 383, 385 transhydrogenase and, 71 small NADH dehydrogenases and, 192 Adenosine diphosphate ribose, cytoAcetoacetate, succinate dehydrogenase chrome b, reductase and, 163 and, 238 Adenosine diphosphate sulfurylase, Acetyl coenzyme A, transhydrogenase reaction catalyzed, 282 and, 71 Adenosine monophosphate Acetyl dephospho coenzyme A, transadenylyl sulfate reductase and, 282, hydrogenase and, 71 283, 284 N-Acetylimidazole, catalase and, 367 cytochrome P-450 reductase and, 167 Acetyl phosphate, glyceraldehyde-3-phosNADH dehydrogenase and, 188, 207 phate dehydrogenase and, 21, 28, 38transhydrogenase and, 71 39,43, 44-45 Adenosine 2’-monophosphate Acetylpyridine adenine dinucleotide sulfite reductase and, 293 cytochrome bs reductase and, 156, 157, transhydrogenase and, 57,58,59,60, 159, 160, 163 61, 71 435

436 Adenosine 3'-monophosphate, transhydrogenase and, 71 Adenosine 3',5'-monophosphate, transhydrogenase and, 71 Adenosine 5'-phosphosulfate, formation of, 279 Adenosine triphosphatase cytochrome c oxidase and, 321 transhydrogenase and, 79 Adenosine triphosphate choline dehydrogenase and, 263 cytocrome c oxidase and, 300, 302, 343 glyceraldehyde-3-phosphate dehydrogenase and, 25,26, 48,49 lipoamide dehydrogenase and, 125 NADH dehydrogenase and, 207 ,redox potentials and, 215, 216 succinate dehydrogenase and, 247, 248, 249 sulfate reduction and, 279 transhydrogenation and, 63-64, 65, 6768, 72, 73-74, 77, 80, 81, 207, 214 Adenylyl sulfate reductase(s) occurrence of, 282 properties, 279-286 Adipose tissue glyceraldehyde-3-phosphate dehydrogenase in, 47, 48 glycolytic enzymes, diabetes and, 47 Adrenal cortex transhydrogenase of, 65 function, 83-85 Aerobacter aerogenes, sulfate reduction by, 281 Aeromonas punctata, sulfate reduction by, 281 Alanine residues, glyceraldehyde-3-phosphate dehydrogenase, 11, 12 Alcohols, catalase and, 388, 398, 401 Aldehydes, glyceraldehyde-3-phosphate dehydrogenase and, 39 Algae, sulfate assimilation by, 279, 280 Alkali cytochrome c oxidase and, 311 respiratory particles and, 240-241, 248 Alkyl bromides, succinate dehydrogenase and, 246 Alloxan diabetes, cytosolic redox state and, 46-47

SUBJECT INDEX

Ally1 alcohol, catalase and, 401-402, 403, 404 Amine oxidase, mixed function, 153-154 Amino acid (s) conservation, glyceraldehyde-3-phosphate dehydrogenase, 14-19 cytochrome bs reductase composition, 155 cytochrome c oxidase composition, 310 cytochrome c peroxidase composition, 348 lactate dehydrogenase, 266 pyridine nucleotide-disulfide oxidoreductases, composition, 100, 102 sequences glyceraldehyde-%phosphate dehydrogenases, 6-8 hemoproteins, 371 thioredoxins, 144 small NADH dehydrogenase composition, 195 synthesis, transhydrogenase and, 80-81 3-Arnino-1H-1,2,4-triazole, catalase and, 376-378 Ammonia, catalase and, 387 Ammonium sulfate glyceraldehyde-3-phosphate dehydrogenase and, 4 lipoamide dehydrogenase and, 124,125, 126 succinate dehydrogenase and, 229-231 Amytal choline dehydrogenase and, 261-262, 263 respiratory particles and mitochondrial, 199 yeast, 217 ubiquinone reductase and, 181,182, 204, 205 Anaerobes, succinate dehydrogenase of, 254 Aniline, hydroxylation of, 150 Anilino-naphthalene sulfonate cytochrome c peroxidase and, 349,359 transhydrogenase and, 69 Antimycin A cholesterol side chain cleavage and, 85 choline dehydrogenase and, 262 a-glycerophosphate dehydrogenase and, 268

437

SUBJECT INDEX

L-lactate dehydrogenase and, 269 small NADH dehydrogenase and, 199 succinate dehydrogenase and, 239, 250 ubiquinone reductase and, 182, 197 yeast NADH dehydrogenase and, 219 Antiparallel sheet, glyceraldehyde-3phosphate dehydrogenase, 13, 14 Arginine residues catalase, 395 glyceraldehyde-3-phosphate dehydrogenase, 12, 22, 24 transhydrogenases and, 213-214, 296 Arsenate, glyceraldehyde-3-phosphate dehydrogenase and, 38, 44-45 Arsenite lipoamide dehydrogenase and, 95-97, 110, 111, 113-114 sulfite reductase and, 289 thioredoxin reductase and, 99 Arsenocholine, choline dehydrogenase and, 261 Arterial tissue, transhydrogenase in, 65 Ascites cells, glyceraldehyde-3-phosphate dehydrogenase in, 47 Ascorbate, cytochrome c peroxidase and, 353 Asparagine residues glyceraldehyde-%phosphate dehydrogenase, 11, 12, 30 lipoamide dehydrogenase, 101 Aspartate residues, glyceraldehyde-3phosphate dehydrogenase, 11, 12, 30 Atebrin, L-lactate dehydrogenase and, 264 Azide catalase and, 376,385,400 cytochrome c oxidase and, 320-321,328, 333 cytochrome c peroxidase and, 350,353 Azotobacter lipoamide dehydrogenase of, 114 transhydrogenase molecular properties, 57-59 reaction mechanism and regulation, 59-62 Azotobacter anile, transhydrogenase of, 54 Azotobacter chroococcum nitrite reductase of, 275,276-277 transhydrogenase of, 54 Azotobacter vinelandii NADH dehydrogenase of, 221

nitrite reductase of, 275, 277 transhydrogenase of, 53,54 function, 80 purification, 55, 56

B Bacillus stearothermophilus glyceraldehyde-3-phosphate dehydrogenase of,2, 4, 9, 23 apoenzyme, 19 pyridine nucleotide binding, 34 Bacteriophage T4, thioredoxin and, 144 Barbiturates, NADH dehydrogenases and, 203, 204, 206 Bathocuproin sulfonate, cytochrome c oxidase and, 308 Bathophenanthroline, succinate dehydrogenase and, 246 Bathophenanthroline sulfonate, NADH dehydrogenase and, 206 3,4-Benzpyrene hydroxylation, pyridine nucleotides and, 152 Benzyl viologen, nitrite reductase and, 275, 276 Betaine, formation of, 260 Betaine aldehyde, choline dehydrogenase and, 261, 262 Betaine aldehyde dehydrogenase, occurrence of, 260 Bicarbonate, succinate dehydrogenase and, 238 Borohydride, glutathione reductase and, 136, 140 Brain glyceraldehyde-3-phosphate dehydrogenase in, 47, 48 a-glycerophosphate dehydrogenase of, 256-258 Bromelain, cytochrome P-450 reductase and, 166, 167 Bromide small NADH dehydrogenases and, 192 succinate dehydrogenase and, 247, 248 3-Bromoacetylpyridine, glyceraldehyde-3phosphate dehydrogenase and, 24 Bromopyruvate glyceraldehyde-3-phosphatedehydrogenase and, 24 succinate dehydrogenase and, 249

SUBJECT INDEX

Bumilleriopsis filiformis, transhydrogenase of, 54 2,3-Butanedione, transhydrogenation and, 296 n-Butanol, catalase and, 401402 Butylhydroperoxide, catalase and, 392 Butyraldehyde, catalase and, 406

C Cadmium, lipoamide dehydrogenase and, 114 Calcium choline dehydrogenase and, 263 transhydrogenases and, 58, 61, 70 Candida utilis, NADH dehydrogenase of, 216-218, 219, 221 Carbohydrate degradation, a-glycerophosphate dehydrogenase and, 259 synthesis, transhydrogenase and, 80 Carbon monoxide catalase and, 401 cytochrome c oxidase and, 301, 317418, 322, 327-328,329, 333, 336-337, 338 cytochrome c peroxidase and, 353 heme iron and, 321-323 nitrite reductase and, 275, 278-279 sulfite reductase and, 289, 293 Carboxylate groups, catalase, 395 Cardiolipin cytochrome c oxidase and, 313 ubiquinone reductase and, 183 Carotenoids, transhydrogenase and, 69 Catalase active site distal ligand identity, 376-385 ligand exchange reactions, 385-388 ligand identity at fifth and sixth coordination positions, 369-376 apoprotein, selective modifications, 376-385 general properties, 366-369 historical background, 363-366 hydroperoxide and, 356 redox reactions, 388-389 nature of Compound I, 389-390 reaction mechanism, 390-408 Catalytic domain, glyceraldehyde-3phosphate dehydrogenase, 16

Cetyldimethylethylammonium bromide, succinate dehydrogenase and, 234 Chemiosmotic hypothesis, transhydrogenase and, 74-75 Chloramphenicol, yeast NADH dehydrogenase and, 217 Chlorella pyrenoidosa, adenylyl sulfate reductase in, 282, 283 Chloride, succinate dehydrogenase and, 247, 248 Chlorobium limicola, adenylyl sulfate reductase of, 286 5-Chloro-3-t-butyl-2-~11loro-4’-nitrosalicylamide, cytochrome c oxidase and, 321 2-Chloroethanol, catalase and, 401 p-Chloromercuriphenyl sulfonate glutathione reductase and, 141 small NADH dehydrogenase and, 203 thioredoxin reductase and, 146 ubiquinone reductase and, 197 Chloroplasts, transhydrogenase of, 66 Chlorosuccinate, succinate dehydrogenase and, 237-238 Cholate NADH dehydrogenase and, 189 ubiquinone reductase and, 178, 182, 183 Cholesterol, side chain cleavage, 83, 84-85 Choline, oxidation to betaine, 260 Choline dehydrogenase electron transport system and, 261-263 properties, 260-261 Chromatium sulfate reduction by, 281 transhydrogenase of, 54 function of, 80 molecular properties, 58, 59 purification, 55, 56 Chromium complexes, oxygen and, 304 Chymotrypsin, cytochrome b5 reductase and, 155 Circular dichroism catalase, 382 cytochrome c oxidase, 319 transhydrogenase and, 62 Clostridium kluyveri, sulfate reduction by, 281 Clostridium nigrijicans, sulfate reduction by, 281

SUBJECT INDEX

Clostridium pasteurianum, sulfate reduction by, 281 Cobalt ammine complexes, oxygen and, 303304 Coenzyme A, transhydrogenase and, 70, 71 Complex I, see under Nicotinamide adenine dinucleotide Complex 1-111 properties of, 197 Complex 11, see also Succinate dehydrogenase properties of, 239 Compound I, nature of, 389-390 Configuration, pyridine nucleotide binding and, 31 Conformation hypothesis, transhydrogenase and, 75 Copper complexes, oxygen and, 304 cytochrome c oxidase and, 302,307309, 314, 315, 317,319, 322, 329-330, 338 lipoamide dehydrogenase and, 114, 122-123 nitrite reductase and, 275 Corpus luteum, transhydrogenase of, 65, 85 Cyanide catalase and, 376, 377, 382,385, 387, 399,401 cholesterol side chain cleavage and, 85 cytochrome c oxidase and, 301,308, 320-321, 335, 336 cytochrome c peroxidase and, 350,353 2-hydroxyacid dehydrogenase and, 272 L-lactate dehydrogenase and, 269 microsomal electron transport and, 148, 151 nitrite reductase and, 275, 276, 277,278 succinate dehydrogenase and, 223,246247, 250 sulfite reductase and, 289, 293 Cyanogen bromide, catalase and, 379-385 Cycloheximide, yeast NADH dehydrogenase and, 217,220 Cystamine, glutathione reductase and, 132 Cysteine, lipoamide dehydrogenase and, 122

439 Cysteine residues glyceraldehyde-3-phosphate dehydrogenase, 5, 12-13, 14, 22, 28, 29, 34, 38, 39, 44, 45, modification of, 20-21 pK. of, 43 Cystine, glutathione reductase and, 132 Cystine residues lipoamide dehydrogenase, 120-122 pyridine nucleotide-disulfide oxidoreductases and, 95, 101, 104 thioredoxin, 92 Cytochrome(s) ubiquinone reductase and, 179, 180 Cytochrome a, see Cytochrome c oxidase Cytochrome b choline dehydrogenase and, 261, 262 succinate dehydrogenase and, 224, 239, 244-245 Cytochrome b2, L-lactate dehydrogenase and, 263-264,266267,296 Cytochrome bs, cytochrome b, and, 267, 296 Cytochrome b5 reductase cytochrome P-450 reductase and, 151153 function of, 150-151 mechanism, microsome bound, 161-162 mechanism of Strittmatter, review, 156-161 methemoglobin reductase and, 164-165 molecular properties, amphipathic and soluble forms, 154-156 structural studies, 162-164 Cytochrome c adenylyl sulfate reductase and, 281, 285-286 choline dehydrogenase and, 261 cytochrome b2 and, 267, 268-269 cytochrome b5 reductase and, 156 cytochrome c oxidase interaction, 334335 356-360 cytochrome c peroxidase an!, cytochrome P-450 reductase and, 167, 168, 169, 170 dehydrogenases and, 90 D-lactate dehydrogenase and, 270 nitrite reductase and, 275 small NADH dehydrogenases and, 194, 195, 196, 198, 199-200, 203,206

440 succinate dehydrogenase and, 250 sulfite reductase and, 288, 290 transhydrogenase and, 68 ubiquinone reductase and, 181, 182, 197 yeast NADH dehydrogenase and, 217 Cytochrome c oxidase biological role, 299-300 chemical and physical properties, 301302, 313-314 electronic spectroscopy, 315-319 electron paramagnetic resonance, 329-334 interaction with cytochrome c, 334335 kinetic studies, 335-337 ligand binding studies, 319-325 models, 314-315 potentiometry, 325-329 historical background, 300-301 lipids of, 312313 mechanisms, 337-344 metal components, 307-309 occurrence of, 300 preparation, 305-307 protein of, 309-312 Cytochrome c peroxidase cytochrome c interaction, 356-360 enzymatic activity, 352353 general comments, 380361 historical background, 345-347 preparation and molecular properties, 347-348 reaction mechanism, 353-356 structural aspects, 348-351 Cytochrome c reductase succinate dehydrogenase and, 245 transhydrogenase and, 55 Cytochrome cI choline dehydrogenase and, 262 ubiquinone reductase and, 205 Cytochrome cs, adenylyl sulfate reductase and, 281 Cytochrome P-450, monooxygenases and, a3 Cytochrome P-450 reductase, 165-166 catalytic activities, 167-169 cytochrome bs reductase and, 151-153 general properties, 166-167 mechanism, 169-173 substrates and components, 149-150

SUBJECT INDEX

Cytoplasm, cytochrome c oxidase polypeptides and, 311

D Deaminonicotinamide adenine dinucleotide glyceraldehyde3phosphate dehydrogenase and, 30 transhydrogenase and, 59 Dehydrogenase(s), characteristics of, 90-91 Demerol, ubiquinone reductase and, 204 Deoxycholate ubiquinone reductase and, 178 yeast NADH dehydrogenase and, 218 Deoxyribonucleic acid, blactate dehydrogenase and, 264 Dephospho coenzyme A, transhydrogenase and, 70, 71 Desulfotomaculum, adenylyl sulfate reductase in, 282 Desuljovibrio, adenylyl sulfate reductase in, 282 Desuljovibrio desuljuricans, sulfate reduction by, 281 Desuljovibrio vulgaris, adenylyl sulfate reductase of, 281, 282484,285 Detergent, see also specific compounds cytochrome bs reductase and, 154-156, 161, 163 cytochrome P-450 reductase and, 166 glyceraldehyde-3-phosphate dehydrogenase and, 26 transhydrogenaae and, 70 Deuterium, cytochrome b6 reductase and, 159 Deuterium oxide small NADH dehydrogenases and, 192-193 succinate dehydrogenase and, 229-230 transhydrogenase and, 70 Deuteroethanol, catalase and, 405 Deuteroformate, catalase and, 404 Diamide, glutathione and, 131 Diaphorase, transhydrogenase and, 59 Dibromoacetone, glyceraldehyde-3-phosphate dehydrogenase and, 23 Dichloroacetate, small NADH dehydrogenases and, 192

441

SUBJECT INDEX

Dichlorohydroquinone, cytochrome c oxidase and, 325 2,6-Dichlorophenolindophenol choline dehydrogenase and, 261 cytochrome b, and, 267 cytochrome P-450 reductase and, 167 a-glycerophosphate dehydrogenase and, 257 2-hydroxyacid dehydrogenase and, 272 small NADH dehydrogenases and, 196, 198, 199, 203, 206 succinate dehydrogenase and, 250 sulfite reductase and, 288 transhydrogenases and, 59 ubiquinone reductase and, 181, 182, 197 yeast NADH dehydrogenase and, 217 2,3-Dicyano-5,6-dichoro-l,4-benzoquinone, ubiquinone reductase and, 182 N,N'-Dicyclohexylcarbodiimide, transhydrogenase and, 72 1,l-Dideuteroethanol, catalase and, 404, 405 1,5-Difluoro-2,4-dinitrobenzene,glyceraldehyde-3-phosphate dehydrogenase and, 22 2,2-Difluorosuccinate, succinate dehydrogenase and, 237 Digitonin, transhydrogenase preparation and, 67 Dihydrolipoamide, physiological form, 93 Dihydroxyacetone phosphate, a-glycerophosphate dehydrogenase and, 256257 1,2-Dihydroxybenzene 3,5-disulfonate, see Tiron 2,3-Dime thoxy-5,6dimethylbenzoquinone, ubiquinone reductase and, 181 2,3-Dimethoxy-5-methylbenzoquinone, ubiquinone reductase and, 181 2,4-Dinitrophenol dehydrogenases and, 63 succinate dehydrogenase and, 247 Dioxygen, see Oxygen Diphosphatidylglycerol, cytochrome c oxidase and, 312 Diphosphoglycerate, glyceraldehyde-3phosphate dehydrogenase and, 35,37 38, 41, 42, 44, 47, 48, 49

Disulfide (s) lipoamide dehydrogenase and, 122 pyridine nucleotidedisulfide oxidoreductases and, 92, 98, 100, 105 Disulfide groups, thioredoxin reductase, 145-146

5,5'-Dithiobis(nitrobenzoate) glutathione reductase and, 132 glyceraldehyde-3-phosphate dehydrogenase and, 21, 36 lipoamide dehydrogenase and, 120,123 Dithionite adenylyl sulfate reductase and, 284 catalase and, 367,375, 382 cytochrome ba,., and, 239, 240 cytochrome c oxidase and, 335, 338 cytochrome P-450 reductase and, 171172 iron-sulfur centers and, 216 lipoamide dehydrogenase and, 113 small NADH dehydrogenase and, 193, 194 semiquinone formation and, 90 sulfite reductase and, 288 thioredoxin reductase and, 145 yeast NADH dehydrogenase and, 219 Dithiothreitol, succinate dehydrogenase and, 227,240, 242,248 Dodecyl sulfate cytochrome c oxidase and, 311 lipoamide dehydrogenase and, 123-124 succinate dehydrogenase and, 227-228, 230-231,234,244,245 ubiquinone reductase and, 180 DT-diaphorase, transhydrogenation and, 52 Duroquinone, ubiquinone reductase and, 182

E Echinocystis macrocarpa seeds, transhydrogenase of, 66 Electromechanochemical model, branshydrogenase, 75 Electron economy, cytochrome c oxidase, 325-326 Electron paramagnetic resonance adenylyl sulfate reductase and, 284

442 catalase, 370, 372, 381, 396 cytochrome c oxidase and, 308309, 314, 316, 317, 321, 322, 327, 335-336 copper and, 329-330 iron and, 331 ligand binding effects, 332-333 p-oxobishemin and, 333-334 valence state changes and, 331332 cytochrome c peroxidase, 350,351,353, 355 fumarate reductase, 256 succinate dehydrogenase, 235, 214, 253, 254 sulfite reductase, 288 ubiquinone reductase, 184-187 yeast NADH dehydrogenase, 219, 220 catalase and, 389,396, 397 sulfite reductase and, 292, 294-295 Electron transport choline dehydrogenase and, 261-263 cytochrome c oxidase and, 301-302 a-glycerophosphate dehydrogenase and, 258 microsomal, 148-149 cytochrome b, reductase system, 150151 cytochrome P-450 reductase system, 149-150 mixed function amine oxidase, 153154 synergism between systems, 151-153 mitochondrial, components of, 178-179 succinate dehydrogenase and, 223-224, 245, 250 transhydrogenase and, 72-74 Emasol, cytochrome c oxidase and, 313 Energy, conservation and coupling by complex I, 214-216, 296 Energy-coupling, transhydrogenase and, 71-75 Enoyl coenzyme A reductase, transhydrogenase and, 88 Entnmoeba histolytica, transhydrogenase of, 66 Erythrocytes glutathione reductase of, 93,94, 138 kinetics, 139-140 substrates, 132

SUBJECT INDEX

glyceraldehyde-3-phosphate dehydrogenase in, 48 methemoglobin reductase of, 164-165 Escherichia coli glutathione reductase of, 102 dissociation constants, 135 glyceraldehyde-3-phosphate dehydrogenase, hybrids of, 26 lipoamide dehydrogenase of, 102, 104, 105, 110, 114, 115, 126 nitrite reductase of, 276, 277 ribonucleotide reductase of, 142-143 sulfate reduction in, 281 sulfite reductase of, 287-290 thioredoxin amino acid composition, 102 partial sequence, 93 thioredoxin reductase general properties, 144-145 specificity, 144 transhydrogenase of, 65, 66 energy and, 73 function, 80-81 molecular properties, 69 reconstitution, 79 Esterase activity, glyceraldehyde-3-phosphate dehydrogenase, 45 EthanoI, catalase and, 392,399,401402, 403, 404, 405 Ethylenediaminetetraacetate cytochrome c oxidase and, 308 a-glycerophosphate dehydrogenase and, 258 Zhydroxyacid dehydrogenase and, 272273 n-lactate dehydrogenase and, 271 lipoamide dehydrogenase and, 126 NADH dehydrogenase and, 206 succinate dehydrogenase and, 247 thioredoxin reductase and, 147, 148 Ethyl hydrogen peroxide, catalase and, 391-392, 395 N-Ethylmaleimide cytochrome b, reductase and, 163, 164 glutathione reductase and, 141 small NADH dehydrogenase and, 203 succinate dehydrogenase and, 246, 249 Ethylmorphine, demethylation of, 150

443

SUBJECT INDEX

Ethylpyridylketone, transhydrogenase and, 59 Euglena gracilis, transhydrogenase of, 54

F Fatty acid(s) cholesterol side chain cleavage and, 85 desaturation of, 151 w-hydroxylation of, 150 synthesis, transhydrogenase and, 88 Ferredoxin sulfite reductases and, 295 transhydrogenase and, 54, 59 Ferric compounds, sulfite reductase and, 293 Ferric ion, cytochrome P-450 reductase and, 168-169 Ferricyanide adenylyl sulfate reductase and, 281,282, 283, 285 bacterial NADH dehydrogenase and, 221 choline dehydrogenase and, 261 cytochrome bl and, 267, 268 cytochrome ba reductase and, 156, 159, 160, 161, 162 cytochrome c oxidase and, 321, 327,337 cytochrome P-450 reductase and, 168, 171, 172 dehydrogenases and, 90 u-glycerophosphate dehydrogenase and, 257 high molecular weight NADH dehydrogenase and, 188, 201,203-204 2-hydroxyacid dehydrogenase and, 272 methemoglobin reductase and, 165 small NADH dehydrogenase and, 194195, 196, 198, 199, 203, 206 succinate dehydrogenase and, 223, 236, 237, 254 sulfite reductase and, 288, 290 thioredoxin reductase and, 148 transhydrogenase and, 59 ubiquinone reductase and, 181, 182, 183, 186, 197, 199,203,207 yeast NADH dehydrogenase and, 217, 218,219, 220, 221 Ferrocyanide, cytochrome c peroxidase and, 353, 355

Ferrocytochrome c, cytochrome c peroxidaae and, 345, 352, 353,354-355, 357, 361 Ferrous ions, succinate dehydrogenase and, 243 Ferrous iron, oxygen reduction and, 304305 Flavin adenylyl sulfate reductase and, 279, 285 choline dehydrogenase and, 260 cytochrome bz and, 268-269 a-glycerophosphate dehydrogenase and, 256, 257, 260 high molecular weight NADH dehydrogenase and, 188,190 Zhydroxyacid dehydrogenase and, 272, 273 succinate dehydrogenase and, 223-225, 226,232234,237,241,243,244,253, 254 sulfite reductases and, 287 Flavin adenine dinucleotide adenylyl sulfate reductase and, 282 283, 284, 297 choline dehydrogenase and, 261 fumarate reductase and, 256 a-glycerophosphate dehydrogenase and, 257, 260 2-hydroxyacid dehydrogenase and, 273 n-lactate dehydrogenase and, 270,272 lipoamide dehydrogenase and, 123,124, 125, 127 mitochondria1 protein and, 297 NADH dehydrogenase and, 217 nitrite reductase and, 275,276, 277,278 pyridine nucleotide-disulfide oxidoreductases and, 92, 94, 95, 97,99, 100, 101, 105 succinate dehydrogenase and, 234-235 sulfite reductase and, 288, 290,292,294 transhydrogenases and, 57-58 Flavin rnononucleotide bacterial NADH dehydrogenase and, 221 cytochrome bs reductase and, 162 2-hydroxyacid dehydrogenase and, 272 L-lactate dehydrogenase and, 264, 265 lipoamide dehydrogenase and, 124-125 low molecular weight NADH dehydrogenases and, 191, 193-194, 198

444 nitrite reductase and, 275, 277, 278 succinate dehydrogenase and, 238, 247, 249 sulfite reductase and, 288, 289, 292, 293-294 ubiquinone reductase, 179-180, 182, 186 yeast NADH dehydrogenase and, 218 Flavoprotein ( s ) functions of, 175-177 mitochondrial, 297 sulfite and, 282 Fluorescence cytochrome bs reductase and, 162, 163 glyceraldehyde-3-phosphate dehydrogenase, 32 lipoamide dehydrogenase, 117-118, 123, 124, 126 thioredoxin, 93 Fluoride catalase and, 377, 383,385 cytochrome c oxidase and, 320421,333 cytochrome c peroxidase and, 350, 353 p-Fluorodinitrobenzene, glyceraldehyde3-phosphate dehydrogenase and, 20 p-Fluoro-m,m'-dinitrophenylsulfone, glyceraldehyde-3-phosphate dehydrogenase and, 37 2-Fluoroethanol, catalase and, 401, 403 Formaldehyde, catalase and, 406 Formamide, catalase and, 386, 399 Formate catalase and, 374-375,383, 385, 386,404 succinate dehydrogenase and, 238, 247, 248 Formic acid, catalase and, 388,395,398, 399, 403, 405-406 Freezing, lipoamide dehydrogenase, 125126 Fructose-l,&diphosphatase, glutathione reductase and, 130 Fumarate choline dehydrogenase and, 262 a-glycerophosphate dehydrogenase and, 258 succinate dehydrogenase and, 238-239, 247, 253 Fumarate reductase anaerobes and, 254 yeast, 255-256

SUBJECT INDEX

p- (2-Furylacrylolyl phosphath, glyceraldehyde3phosphate dehydrogenase and, 35, 36

G Glucokinase, activity, liver and, 47 Gluconeogenesis, glyceraldehyde-3-phosphate dehydrogenase and, 45-49 Glucose-6-phosphate dehydrogenase, glutathione reductase and, 131 Glutamate, redox couple, 82 Glutamate dehydrogenase, transhydrogenases and, 52, 82, 85-86 Glutamate residues, glyceraldehyde-3phosphate dehydrogenase, 30 Glutamine residues, lipoamide dehydrogenase, 101 bis-N,N(y-Glutamyl)cystine,glutathione reductase and, 132 Glutathione nitrite reductase and, 275 reduced : oxidized ratios, 129-130 reduction of, 87 reoxidation of, 93 Glutathione-cystine oxidoreductase, 130 Glutathione-homocystine oxidoreductase, 130 Glutathione-insulin transhydrogenase, 130 Glutathione peroxidase, 130 Glutathione protein disulfide oxidoreductase, 130 Glutathione reductase amino acid composition, 102, 104, 105 cystine residues, 104 kinetic studies, 138-141 mechanism, 94,97-98,134 metabolic functions, 129-133 reaction catalyzed, 92 reduction of, 112, 113 specificity of, 92-93 coenzymes and, 91 thiol groups, 141-142 two-electron-reduced enzyme, properties, 133-138 Glyceraldehyde, glyceraldehyde-3-phosphate dehydrogenase and, 44 Glyceraldehyde-3-phosphatedehydrogenase catalytic properties,

SUBJECT INDEX

mechanism of action, 38-46 metabolic role, 45-49 pyridine nucleotide binding, 28-38 chemical modifications, 20-24 dissociation and hybridization, 24-27 distribution of, 3 historical background, 2-3 maximal activities in rat and human tissues, 47-48 metabolic role, 4549 molecular properties, isolation, 3-4 structure, 5-27 other activities of, 44-45 physiological activity, 38-44 preexisting asymmetry model, 35-38 pure, sources of, 2 reaction catalyzed, 1 ~-Glycerol-3-phosphatedehydrogenase, properties, 256-260 a-Glycerophosphate, succinate dehydrogenase and, 239, 250 a-Glycerophosphate cycle, function of, 259 Glycine catalase and, 387-388 oxidative decarboxylation of, 108 Glycine residues, glyceraldehyde-3-phosphate dehydrogenase, 12 Glycogen synthetase, glutathione reductase and, 130 Glycolate, succinate dehydrogenase and, 238 Glycolysis, glyceraldehyde-$phosphate dehydrogenase and, 4549 Glyoxylate, succinate dehydrogenase and, 238 Glyoxylate cycle, transhydrogenase and, 80 Guanidine, glyceraldehyde-3-phosphate dehydrogenase and, 24 Guanidinium chloride cytochrome bz and, 268 lipoamide dehydrogenase and, 123, 124 NADH dehydrogenases and, 199,207 succinate dehydrogenase and, 232 Guanosine triphosphate, oxidation of NAD+-linked substrates and, 110 Guiacol, cytochrome c peroxidase and, 353

H Halogenacetic acids, glyceraldehyde-3phosphate dehydrogenase and, 2 Hansenula anomala, L-lactate dehydrogenase of, 265, 268, 269 Heart cytochrome c oxidase of, 311 glyceraldehyde-3-phosphatedehydrogenase in, 47, 48 lipoamide dehydrogenase of, 102, 104 111-114, 116 transhydrogenase of, 62-63, 64, 65 kinetic constants, 76-77 molecular properties, 70-71 preparation, 67-68 Helix, glyceraldehyde-3-phosphate dehydrogenase, 13-14 Heme cytochrome bf and, 268-269 sulfite reductases and, 287, 288,289, 290, 292,293,294,295 Heme az, nitrite reductase and, 275,297 Heme A, cytochrome c oxidase and, 307, 309, 314-315 Heme G adenylyl sulfate reductase and, 285, 286 nitrite reductase and, 278 Heme oxygenase, cytochrome P-450 reductase and, 150 Hemoglobin, absorption spectra, 317-319 Hemoprotein(s), amino acid sequences, 371 Hepatoma, transhydrogenase in, 65 Hexokinase, activity in muscle, 47 Histidine residues catalase, 370-372,375,377-378,381,383, 395 cytochrome c peroxidase, 350 glyceraldehyde-3-phosphate dehydrogenase, 14, 44 modification of, 23-24 lipoamide dehydrogenase, 101 succinate dehydrogenase, 235 Homocystine, glutathione redubtasq and, 132 Horseradish peroxidase, reaction cycle, 356 Human, glyceraldehyde-3-phosphate dehydrogenase of, 9

446

SUBJECT INDEX

Hydrazoic acid, catalase and, 388 Hydrogen bonds, glyceraldehyde-3-phosphate dehydrogenase, 11, 14, 15,25, 43 Hydrogen peroxide catalase and, 388,395,398,403 cytochrome c peroxidase and, 345-346 Hydrogen sulfide, formation of, 279 Hydroperoxides catalase and, 398, 406 cytochrome c peroxidase and, 352,353, 354 Hydrophobic bonds, catalase and, 367 Hydroquinone, cytochrome c peroxidase and, 353 D-2-Hydroxyacid dehydrogenase, properties, 272-273 p-Hydroxybutyrate, redox couple, 82 ~-2-Hydroxybutyrate,D-laCtate dehydrogenase and, 270 25-Hydroxycholecalciferol,hydroxylation of, 83 Hydroxylamine catalase and, 388, 398 formation of, 153 nitrite reductase and, 275,276, 277,278, 279 sulfite reductase and, 288-289,290, 292, 293, 294, 295 p-Hydroxymercuribenzoate, transhydrogenase and, 58 cu-Hydroxymonocarboxylic acids, L-lactate dehydrogenase and, 267 5-Hydroxy-1,4-naphthoquinone,see Juglone 17-Hydroxyprogesterone, hydroxylation of, 150 Hyponitrite, nitrite reductase and, 275

I Imidazole, catalase and, 370 Infrared spectroscopy, cytochrome c oxidase and, 321,322, 323 Inosine diphosphate, succinate dehydrogenase and, 247, 248 Inosine triphosphate, succinate dehydrogenase and, 247, 248 Insulin glycolytic enzymes and, 47

phenylalanyl chain, glutathione reductase and, 130 Iodide, succinate dehydrogenase and, 247, 248 Iodine cytochrome ba reductase and, 163 glyceraldehyde-3-phosphate dehydrogenase and, 22-23, 24 Iodoacetamide glyceraldehyde-3-phosphate dehydrogenase and, 35, 36-37,38,42,43 lipoamide dehydrogenase and, 118-119, 122 Iodoacetate glyceraldehyde-3-phosphate dehydrogenase and, 20,28,35, 36-37,38, 39, 42-43, 45 lipoamide dehydrogenase and, 120, 123 a-Iodopropionamide, glyceraldehyde-3phosphate dehydrogenase and, 33 a-Iodopropionic acid, glyceraldehyde-3phosphate dehydrogenase and, 33 Iodosobenzoate, glyceraldehyde-3-phosphate dehydrogenase and, 20, 21 Ionic strength cytochrome P-450 reductase and, 167 glutathione reductase and, 139-140 glyceraldehyde-3-phosphate dehydrogenase and, 25, 26,43 Iron, see also Nonheme iron adenylyl sulfate reductase and, 279, 297 carbon monoxide and, 321-323 catalysis and, 363-364 cytochrome c oxidase and, 302,307309, 321 electron paramagnetic resonance, 331 deficiency, NADH dehydrogenases and, 219, 221 a-glycerophosphate dehydrogenase and, 256, 257, 260 high molecular weight NADH dehydrogenases and, 188, 190,201-202 low molecular weight NADH dehydrogenases and, 191, 193-194,198 mitochondria1 flavoprotein and, 297 nitrite reductase and, 275 sulfite reductase and, 288, 290, 292,293, 295 ubiquinone reductase and, 179, 180, 182, 183-184, 186-187, 204, 209-210

447

SUBJECT INDEX

Iron-copper coupling, cytochrome c oxidase, 326-327, 338-343 Iron sulfide, succinate dehydrogenase, 230-232,235,243,244, 245,246,253, 254, 296 Iron-sulfur centers bacterial NADH dehydrogenase and, 221 energy conservation and, 214-216, 296 yeast NADH dehydrogenase and, 217, 218, 219 Isocitrate dehydrogenase, transhydrogenase and, 52-53,81,84,86-88 Isoleucine residues, disulfide oxidoreductases, 104 Isonitriles, catalase and, 401 Isozymes, lipoamide dehydrogenase,. 109

J Juglone ubiquinone reductase and, 182 yeast NADH dehydrogenase and, 219, 220

K a-Keto acids decarboxylation of, 108 2-hydroxyacid dehydrogenase and, 272 a-Ketoglutarate, reductive carboxylation of, 87 Kidney transhydrogenase in, 64 function, 83 Kinetic studies, cytochrome c oxidase, 335-337

1 Lactate dehydrogenase(s1, types of, 263 n ( -)-Lactate dehydrogenase, 269-270 enzymic properties, 270-272 physical properties, 270 L(+)-Lactate dehydrogenase cytochrome b, core, 266-267 enzymic properties, 267-269 historical background, 263-264 physical properties, 264-266

Lactobacillus leichmannii, ribonucleotide reductase of, 142 Lecithin, transhydrogenase and, 70-71 Leucine residues disulfide oxidoreductases, 104 glyceraldehyde-%phosphate dehydrogenase, 11, 12 Leucomethylene blue, nitrite reductase and, 275 Leucomethyl viologen, 2-hydroxyacid dehydrogenase and, 272 Lipase, cytochrome P-450 reductase and, 167, 168,169,170-171, 172 Lipid cytochrome bs reductase and, 151, 158 cytochrome c oxidase and, 309, 311, 312-313 high molecular weight NADH dehydrogenase and, 188 peroxidation, cytochrome P-450 reductase and, 168,169 transhydrogenase and, 70 Lipoamide dehydrogenase(s1 amino acid composition, 102, 104,105 apoenzymes, 124 coenzyme specificity, 94 cystine residues, 104, 120-122 distribution, 106107 kinetic studies, 115-117 mechanism, 94-97, 126-129 mechanism of Massey and Veeger, review of, 111 metabolic functions, 107-1 10 reaction catalyzed, 92 role of NAD' as modifier, 117-120 structural studies, 120-126 two-electron-reduced enzyme, properties, 111-115 Lipoate glutathione reductase and, 93, 132, 140 ubiquinone reductas? and, 181 Lipoprotein, cytochrome c oxidase and, 302 Liver glyceraldehyde-3-phosphate dehydrogenase in, 47, 48 glycolysis, cytosolic redox state and, 46 lipoamide dehydrogenase of, 116-117 transhydrogenase in, 64

SUBJECT INDEX

Lobster muscle glyceraldehyde-3-phosphate dehydrogenase cysteine residues, 20,28 hybrids of, 26-27 lysine residues, 22 primary structure, 5, 6-8, 9, 18 pyridine nucleotide binding, 33 tyrosine residues, 22-23 Lubrol, NADH dehydrogenase and, 189 Lysine residues cytochrome bs reductase, 164 dihydrolipoate and, 108 glutathione reductase, 105, 119 glyceraldehyde-3-phosphatedehydrogenase, 11, 14, 44, 45 modification of, 21-22,3748 lipoamide dehydrogenase, 119, 126 thioredoxin, 119 Lysolecithin, transhydrogenase and, 68,71 Lysophosphatidylethaolamine, cytochrome P-450 reductase and, 150 Lysosomes, cytochrome bs reductase and, 154-156

M Magnesium choline dehydrogenase and, 263 transhydrogenase and, 70,77 Magnetic susceptibility, catalase, 382,396 Malate choline dehydrogenase and, 262 succinate dehydrogenase and, 237 Maleate, succinate dehydrogenase and, 238 “Malic” enzyme, steroid hydroxylations and, 83-84 Malonate choline dehydrogenase and, 263 succinate dehydrogenase and, 238, 247, 248 Manganese nitrite reductase and, 277 transhydrogenase and, 70 Membrane(s) integrity, glutathione and, 131 Menadione bacterial NADH dehydrogenase and, 221

cytochrome P-450 reductase and, 168, 169 2-hydroxyacid dehydrogenase and, 272 small NADH dehydrogenases and, 194195, 196, 206 sulfite reductase and, 288, 290 ubiquinone reductaae and, 181,182, 197, 207 yeast NADH dehydrogenase and, 218, 220 Menadione reductase, transhydrogenase and, 55 Mercaptoethanol cytochrome c oxidase and, 311 small NADH dehydrogenase and, 198 succinate dehydrogenase and, 243 Mercurials glyceraldehyde-3-phosphate dehydrogenase and, 21 large NADH dehydrogenase and, 203204 lipoamide dehydrogenase and, 114, 122 low molecular weight NADH dehydrogenases and, 191, 199, 203 nitrite reductase and, 276 succinate dehydrogenase and, 246 sulfite reductase and, 292 ubiquinone reductase and, 181, 186, 199, 203, 204 p-Mercuribenzoate choline dehydrogenase and, 261 cytochrome ba reductase and, 163 cytochrome c peroxidase and, 348 cytochrome P-450 reductase and, 168 L-lactate dehydrogenase and, 264 nitrite reductase and, 275, 276, 278 sulfite reductase and, 293 p-Mercuriphenyl sulfonate n-lactate dehydrogenase and, 27l’ sulfite reductase and, 288, 289 yeast NADH dehydrogenase and, 217 Mersalyl cytochrome bs reductase and, 163, 164 cytochrome P-450 reductase and, 168 small NADH dehydrogenase and, 193 succinate dehydrogenase and, 246 Metal(s) transition, oxygen reduction and, 303 Methanol, catalase and, 391, 392, 401, 403,404,405,406

SUBJECT INDEX

Methemoglobin reductase, cytochrome ba reductase and, 164-165 Methionine residues glyceraldehyde-3-phosphate dehydrogenase, 11 lipoamide dehydrogenase, 118 5-Methoxyindole-2-carboxylate, lipoamide dehydrogenase and, 110 Methylene blue choline dehydrogenase and, 261 cytochrome b, and, 267 a-glycerophosphate dehydrogenase and, 257 2-hydroxyacid dehydrogenase and, 272 nitrite reductase and, 275 succinate dehydrogenase and, 246 ubiquinone reductase and, 182 Methylene succinate, succinate dehydrogenase and, 238 3-Me thylflavin adenine dinucleotide, lipoamide dehydrogenase and, 125 Methyl hydrogen peroxide, catalase and, 390-391, 398, 405 Methyl hydroperoxide, catalase and, 392 N-Methyl hydroxylamine, catalase and, 399 o-Me thy1 hydroxylamine, catalase and, 399 2-Methylnaphthoquinone, see Menadione Methyl succinate, succinate dehydrogenase and, 237-238 Methyl viologen adenylyl sulfate reductases and, 281 nitrite reductase and, 276 sulfite reductases and, 287, 288-289, 290, 292, 293 Micrococcus denitrifians, transhydrogenase of, 73 Microorganism(s) , succinate dehydrogenase of, 254-256 Microsomes electron transport, 148-149 cytochrome b, reductase system, 150151 cytochrome P-450 reductase system, 149-150 mixed function amine oxidase, 153154 synergism between systems, 151-153 hydroxylation reactions in, 87

Mitochondria choline dehydrogenase and, 260-263 fatty acid synthesis in, 88 glutamate and isocitrate metabolism, 85-88 a-glycerophosphate dehydrogenase of, 256-259 membranes, cytochrome c oxidase and, 302 monooxygenase reactions, 83-85 nicotinamide adenine dinucleotide dehydrogenases, 177-178 energy conservation and, 214-216 high molecular weight, 187-189 inhibitors of, 203-207 low molecular weight, 189-198 relevance of low and high molecular weight dehydrogenases, 198-203 transhydrogenation and, 207-214 ubiquinone reductase (Complex I), 178-187 nicotinamide nucleotides, redox state, 81-82 transhydrogenase of, 62, 65 assay, 66-67 preparation, 67-68 Models, cytochrome c oxidase, 314-315 Molecular weight, succinate dehydrogenase, 232-234 Molybdenum bacterial NADH dehydrogenase and, 221 ubiquinone reductase and, 187 Monochloroacetate, small NADH dehydrogenases and, 192 Monofluorosuccinate, succinate dehydrogenase and, 237 Monooxygenase ( s ), transhydrogenases and, 83-85 Musca domestica, a-glycerophosphate dehydrogenases of, 258-259 Muscle glyceraldehyde-3-phosphate .dehydrogenase in, 47,48 transhydrogenase in, 64 Mycobacterium, 2-hydroxyacid dehydrogenase of, 273 Mycobacterium phlei, transhydrogenase of, 65-66

450

SUBJECT INDEX

Myoglobin, absorption spectra, 317-319 Myohematin, 300

N l,2-Naphthoquinone 4-sulfonate1 cytochrome b, and, 267 Neotetrazolium, cytochrome P-450 reductase and, 168 Nerve transmission, glutathione and, 131 Neurospora crassa cytochrome c oxidase of, 311 nitrite reductase of, 275-276 Nicotinamide adenine deoxydinucleotide, glyceraldehyde-3-phosphate dehydrogenase and, 30 Nicotinamide adenine dinucleotide choline dehydrogenase and, 261,262 cooperativity of binding, 30-35 cytochrome c oxidase and, 335 glyceraldehyde-3-phosphate dehydrogenase and, 3, 4, 26,42,45, 48 amino acid modification and, 21, 22 binding, 10-16, 28-38, 49 pK. and, 43 inhibition by, 206-207 lipoamide dehydrogenase and, 117-120, 125, 126, 128 nitrite reductase and, 275, 277, 278 oxidized : reduced ratio, glycolysis and, 46, 49 succinate dehydrogenase and, 252 Nicotinamide adenine dinucleotide dehydrogenase Azotobacter vinelandii, 221 high molecular weight, 187-189, 190 relevance to mitochondrial enzyme, 198-203 inhibitors of, 203-207 low molecular weight, 189-198, 295 relevance to mitochondrial enzyme, 198-203 transhydrogenase and, 78 yeast, 216-221 Nicotinamide adenine dinucleotide 3'phosphate, transhydrogenase and, 59, 61, 69 Nicotinamide adenine dinucleotide phosphate cytochrome bs reductase and, 156, 157, 160

glutathione reductase and, 134-137 large NADH dehydrogenase and, 202 nitrite reductase and, 275,276, 278 respiratory particles and, 200 sulfate reduction and, 281,283 sulfite reductases and, 287-295 transhydrogenation, complex I and, 207-214,296 ubiquinone reductase and, 181 Nicotinamide adenine dinucleotide-ubiquinone reductase, 178-179 activities, 180-183 composition, 179-180 spectral properties, 183-187 Nicotinamide bdeaminoadenine dinucleotide, glyceraldehyde-3-phosphate dehydrogenase and, 30 Nicotinic acid hydrazide adenine dinucleotide, glyceraldehyde-3-phosphate dehydrogenase and, 30 Nicotinamide mononucleotide, transhydrogenase and, 59 Nicotinamide nucleotide transhydrogenase AB-specific historical, 62-64 kinetics and reaction mechanism, 7578 molecular properties, 69-71 occurrence, 64-66 preparation and assay, 66-69 reconstitution, 78-79 relationship to energy-coupling system, 71-75 BB-specific historical, 52-53 molecular properties, 57-59 occurrence, 63-54 purification and away, 54-57 reaction mechanism and regulation, 59-62 definition, 5162 physiological roles, 79-81 fatty acid synthesis, 88 mitochondrial glutamate and isocitrate metabolism, 85-88 mitochondrial monooxygenase reactions, 83-85 redox state of mitochondrial nicotinamide nucleotides, 81-82

45 1

SUBJECT INDEX

Nicotinylhydroxamic acid adenine dinucleotide, glyceraldehyde-3-phosphate dehydrogenase and, 30 Nigericin, transhydrogenase and, 72 Nitrate reduction of, 273 succinate dehydrogenase and, 227, 247, 248 Nitric acid, ferrocytochrome c peroxidase complex, 350 Nitric oxide catalase and, 372,375,376 nitrite reductases and, 274,275 Nitrite catalase and, 391, 392,400 sulfite reductase and, 288-289,290,292, 293, 294, 297 Nitrite reductase (s) occurrence of, 274 properties of, 275-279, 297 Nitrogen, nitrite reductases and, 274 Nitrogen mustard, choline dehydrogenase and, 261 Nitrones, formation of, 153 p-Nitrophenylacetate, glyceraldehyde-3phosphate dehydrogenase and, 21,45 Nitrous acid, catalase and, 388,398 Nitrous oxide, catalase and, 400 Nonheme iron, see also Iron adenylyl sulfate reductase and, 282, 284, 285 choline dehydrogenase and, 260 succinate dehydrogenase and, 223-225, 226 sulfite reductases and, 287 Nuclear magnetic resonance, ferrocytochrome c, 357-359

0 Oleate, gluconeogenesis and, 46, 47 Oligomycin succinate dehydrogenase and, 249 transhydrogenase and, 67-68, 72 Optical rotatory dispersion glyceraldehyde-3-phosphate dehydrogenase, 14 transhydrogenase, 62 Ovary transhydrogenase, function, 83,85

Oxalate 2-hydroxyacid dehydrogenase and, 272 n-lactate dehydrogenase and, 271 Oxaloacetate choline dehydrogenase and, 262 D-lactate dehydrogenase and, 271 succinate dehydrogenase and, 237,238, 248-249 Oxidase(s), characteristics of, 90-91 Oxidative phosphorylation, transhydrogenase and, 73 Oximes, catalase and, 398 p-Oxobishemin A, cytochrome c oxidase and, 333-334 Oxygen adenylyl sulfate reductase and, 282,284 choline dehydrogenase and, 261, 262 cytochrome b, reductase and, 156 cytochrome c oxidase and, 323-325, 336-337, 338-343 cytochrome P-450 reductase and, 170 electronic structure, 303 a-glycerophosphate dehydrogenase and, 260 heme absorption spectra and, 317419 lactate dehydrogenases and, 269,270 microsomal mixed function oxidations and, 149, 152 reduction, chemistry of, 302-305 succinate dehydrogenase and, 236 sulfite reductase and, 288

P Palmitylcoenzyme A, transhydrogenase and, 70,78, 88, 212 Palmityldephosphocoenzyme A, transhydrogenase and, 70, 71 Pantothine, glutathione reductase and, 132 Peas, glutathione reductase of, 138 Pea seed, glyceraldehyde-3-phosphate dehydrogenase of, 40 Penicillium chrysogenum, glutathione reductase of, 138 Peptococcus glycinophilus glycine decarboxylation by, 108 lipoamide dehydrogenase of, 112 Peracetic acid, catalase and, 392-393, 395, 397

452 Perchlorate small NADH dehydrogenases and, 191, 192, 203 succinate dehydrogenase and, 227-228, 229,231,232, 247,248 Perfluoro-n-hexane, oxygen consumption and, 152-153 Peroxide, see also Hydrogen peroxide oxygen reduction and, 303,305 PH cytochrome bs reductase and, 163,164 cytochrome c peroxidase and, 350-351 cytochrome P-450 reductase and, 167, 168 glutathione reductase and, 134, 140, 141 glyceraldehyde-3-phosphate dehydrogenase and, 41,48,49 lipoamide dehydrogenase and, 116, 119, 125, 128 small NADH dehydrogenases and, 194195, 203 succinate dehydrogenase and, 247, 248 thioredoxin reductase and, 144 transhydrogenases and, 58, 76,208,210211, 213 o-Phenanthroline 2-hydroxyacid dehydrogenase and, 272 n-lactate dehydrogenase and, 271-272, 273 NADH dehydrogenases and, 206 succinate dehydrogenase and, 246 Phenazine methosulfate choline dehydrogenase and, 261, 262, 263 a-glycerophosphate dehydrogenase and, 257, 258 n-lactate dehydrogenase and, 270 succinate dehydrogenase and, 223,225, 227, 232,236,237, 238, 239,242, 243, 246, 249-250, 253, 254 Phenobarbital, cytochrome P-450 reductase and, 150 Phenols, catalase and, 398 Phenylalanine residues, glyceraldehyde-3phosphate dehydrogenase, 11, 33 Phenyl mercuric acetate, glutathione reductase and, 141 Phenylmethylsulfonyl fluoride, L-lactate dehydrogenase and, 265,266

SUBJECT INDEX

Phosphate choline dehydrogenase and, 263 glyceraldehyde-3-phosphate dehydrogenase and, 44,48 nitrite reductase and, 275 succinate dehydrogenase and, 247 Phosphatidylcholine cytochrome c oxidase and, 312, 313 cytochrome P-450 reductase and, 149 Phosphatidylethanolamine, cytochrome c oxidase and, 312, 313 Phosphatidylinositol, cytochrome c oxidase and, 312, 313 3'-Phosphoadenosine 5'-phosphosulfate, formation of, 279 Phosphocreatine, glyceraldehyde-3-phosphate dehydrogenase and, 48 Phosphofructokinase activity in adipose tissue, 47 glycolysis and, 49 3-Phosphoglycerate, a-glycerophosphate dehydrogenase and, 258 Phosphoglycerate kinase glycolysis rate and, 46,49 pyruvate kinase and, 47 Phospholipase (9) choline dehydrogenase and, 260 transhydrogenase and, 70 Phospholipase A cytochrome c oxidase and, 313 a-glycerophosphate dehydrogenase and, 257 n-lactate dehydrogenase and, 270,271 NADH dehydrogenase and, 187 Phospholipase C low molecular weight NADH dehydrogenase and, 191 yeast NADH dehydrogenase and, 218 Phospholipid amine oxidase and, 153 cytochrome bs reductase and, 161 succinate dehydrogenase and, 244,245, 247 synthesis, a-glycerophosphate dehydrogenase and, 259 ubiquinone reductase and, 179, 180, 182-183, 201 Photoirradiation, semiquinones, 90

453

SUBJECT INDEX

Photooxidation, glyceraldehyde-3-phosphate dehydrogenase and, 24 Photosynthesis, transhydrogenation and, 64, 66

Piericidin A respiratory particles and, 199 small NADH dehydrogenase and, 199 transhydrogenation and, 211-212 ubiquinone reductase and, 181,203, 204-206, 214

yeast NADH dehydrogenase and, 219, 220

Pig muscle, glyceraldehyde-3-phosphate dehydrogenase of, 6-8, 18, 24 hybrids of, 26-27 tyrosine residues, 23 Plants, transhydrogenases of, 54 Plasmalogen, biosynthesis, 151 Polyacrylamide gel electrophoresis, succinate dehydrogenase, 227-228,230, 245

Potassium bromide, cytochrome bs reductase and, 162 Potassium ions, transhydrogenase and, 72

Potentiometry cytochrome c oxidase and, 322 electron economy, 325-326 interpretation and summary, 328-329 iron-copper coupling, 326-327 ligand binding, 327-328 Progesterone, hydrdxylation of, 150 Proline residues, glyceraldehyde-3-phosphate dehydrogenase, 11 n-Propanol, catalase and, 401-402 Propargyl alcohol, catalase and, 401-402, 403

Propionibacterium arabinosum, a-glycerophosphate dehydrogenase of, 260 2-Propyn-1-01, catalase and, 404 Prostaglandin, biosynthesis, 168 Prosthetic groups, succinate dehydrogenase, 234-235 Protease, D-lactate dehydrogenase and, 270

Protein(s) catalase function and, 369-370 cytochrome c oxidase and, 309-312 synthesis, glutathione and, 131

Protein disulfide isomerase, glutathione and, 132 Proteus mirabilis, sulfate reduction by, 281

Proteus vulgaris, sulfate reduction by, 281 Protoheme, cytochrome c peroxidase and, 345,346, 348,349

Protons, transhydrogenase and, 77-78 Protoporphyrin IX, catalase and, 366 Pseudomonas transhydrogenase function, 80 molecular properties, 58-59 reaction mechanism and regulation, 59-61

Pseudomonas aeruginosa nitrite reductase of, 274, 275 transhydrogenase of, 53 molecular properties, 57 purification, 54, 56 Pseudomonas denitrificans, nitrite reductase of, 274275 Pseudomonas fluorescena transhydrogenase of, 52-53 purification, 54, 56 Pseudomonas hydrophila, sulfate reduction by, 281 Pseudomonas stutzeri, nitrite reductase of, 274 Pyocyanine, nitrite reductase and, 275 Pyridine adenine dinucleotide, transhydrogenase and, 59 Pyridine aldehyde adenine dinucleotide cytochrome bs reductase and, 156,158159

glyceraldehyde-3-phosphate dehydrogenase and, 30 transhydrogenase and, 59 Pyridine nucleotidedisulfide oxidoreductases mechanism, similarities and contrasts, 94-99

reaction catalyzed-chemical similarities and cross-reactivity, 92-94 structure, similarities and contrasts, 99-105

Pyridoxal phosphate, glyceraldehyde-3phosphate dehydrogenase and, 22 Pyrogallol, cytochrome c peroxidase and, 353

SUBJECT INDEX

Pyrophosphate lipoamide dehydrogenase and, 125 transhydrogenase and, 72 Pyruvate gluconeogenesis and, 47,48 oxidation, transhydrogenase and, 80 Pyruvate dehydrogenase lipoamide dehydrogenase and, 126 transhydrogenase and, 55 Pyruvate kinase, glyceraldehyde-3-phosphate dehydrogenase and, 47 Pythium ultimum, lipoamide dehydrogenase of, 112

0 Quinoline oxide, choline dehydrogenase and, 262 Quinones, sulfite reductase and, 293

R Rabbit muscle glyceraldehydea-phosphate dehydrogenase, 25 amino acid modification, 22,24 hybrids of, 26-27 pyridine nucleotide binding, 33-35 Rat muscle, glyceraldehyde-3-phosphate dehydrogenase of, 25 Redox potentials cytochrome c oxidase, ligand binding and, 327-328 cytochrome P-450 reductase and, 172 Reductase(s), characteristics of, 90-91 Respiration, transhydrogenation and, 64 65, 72 R hodopseudomonas palustris sulfate reduction by, 281 transhydrogenase of, 66 R hodopseudomonas spheroides sulfate reduction by, 281 transhydrogenase of, 66 molecular properties, 71 preparation, 68 reconstitution, 79 Rhodopseudomonas viridis, transhydrogenase of, 66 Rhodospirillum moliachianum, transhydrogenase of, 66

Rhodospiiillum rubrum succinate dehydrogenase of, 254-255 transhydrogenase of, 66,69, 213 energy and, 73,74 molecular properties, 69,71 preparation, 68 reconstitution and, 78 Riboflavin cytochrome ba reductase and, 162 deficiency, glutathione reductase and, 131 nitrite reductase and, 275 Ribonucleotide reductase, thioredoxin and, 142-143 Rose Bengal, cytochrome c peroxidase and, 350 Rotenone cholesterol side cleavage and, 85 choline dehydrogenase and, 262-263 small NADH dehydrogenases and, 196, 199 transhydrogenation and, 211 ubiquinone reductase and mitochondrial, 181,182,183,197,204206,214-215 yeast, 217,219

S Saccharomyces carlsbergensis, NADH dehydrogenase of, 216, 219-221 Saccharomyces cerevisiae, see also Yeast adenylyl sulfate reductase of, 283 lactate dehydrogenase of, 269-270 L-lactate dehydrogenase of, 264-266, 268, 269 NADH dehydrogenase of, 216,217, 219-221 succinate dehydrogenase of, 254 sulfite reductase of, 292-295 transhydrogenase of, 66 Salmonella typhimurium, sulfite reductase of, 290-291 Seconal ubiquinone reductase and mitochondrial, 204 yeast, 217 Semicarbazide catalase and, 379 choline dehydrogenase and, 262

SUBJECT INDEX

Semiquinone cytochrome P-450 reductase, 170, 172 dehydrogenases and, 90, 97-98 glutathione reductase and, 137-138 oxidases and, 90 thioredoxin reductase, 147-148 Serine residues, pyridine nucleotide-disulfide oxidoreductases, 101 Serum albumin, lipoamide dehydrogenase and, 126 j3-Sheet, glyceraldehyde-3-phosphate dehydrogenase, ll, 14 Soybean leaves, nitrite reductase of, 277-278 Spinach lipoamide dehydrogenase of, 112 transhydrogenase molecular properties, 59 purification, 54-55, 56 Spleen, glyceraldehyde-3-phosphate dehydrogenase in, 48 Starvation, cytosolic redox state and, 46 Steroid(s) metabolism, transhydrogenase ahd, 65, 83-84 Steroid 17,20-lyase, cytochrome P450 reductase and, 150 Streptococcus jnecalis, a-glycerophosphate dehydrogenase of, 260 Sturgeon muscle, glyceraldehyde-3-phosphate dehydrogenase of, 28, 33, 42 Subunits catalase, 366 cytochrome b?, 264-266 cytochrome c oxidase and, 311 glyceraldehyde-3-phosphate dehydrogenase, primary structure, 5 succinate dehydrogenase, 230-232 Succinate cholesterol side chain cleavage and, 85 succinate dehydrogenase preparation and, 223-224, 227,242-243, 248 Succinate dehydrogenase a-glycerophosphate dehydrogenase and, 258 mammalian, 222-223 enzymic properties, 236-245 inhibitors and modifiers, 245-247 mechanism, 251-254

455 molecular properties, 223-226, 296 regulatory properties, 247-251 microorganisms and, 254-256 Succinyl coenzyme A, succinate dehydrogenase and, 247 Sulfate nitrite reductase and, 275 reduction, pathways, 279,281 succinate dehydrogenase and, 247 Sulfhydryl groups, see also Thiol groups catalase, 368 glyceraldehyde-3-phosphate dehydrogenase and, 2 Sulfhydryl reagents, transhydrogenase and, 69-70 Sulfide acid-labile, ubiquinone reductase and, 179, 180, 182, 184, 186-187, 204, 209-210 adenylyl sulfate reductase and, 284285, 297 high molecular weight NADH dehydrogenases and, 90,201-202 low molecular weight NADH dehydrogenases and, 191, 193-194, 195,198, 206 mitochondria1 flavoprotein and, 297 succinate dehydrogenase and, 224, 226, 243 sulfite reductase and, 287, 288, 289, 290, 292, 293, 295 Sulfite adenylyl sulfate reductase and, 282, 284 lipoamide dehydrogenase and, 122 oxidases and, 90 Sulfite reductaseb) NADPH-dependent occurrence, 286,287-288 properties, 288-295 reduced methyl viologen-dependent, 295 types of, 286-287 Superoxide catalase and, 398 cytochrome P450 reductase and, 169 dehydrogenases and, 90 mixed function oxidations and, 153 oxygen reduction and, 303

456

SUBJECT INDEX

T Temperature glyceraldehyde-3-phosphate dehydrogenase and, 25 lipoamide dehydrogenase and, 116, 118, 123, 125 NADH dehydrogenases and, 203 succinate dehydrogenase and, 228-229, 231-232 Tetrathionate, glyceraldehyde-3-phosphate dehydrogenase and, 20, 21 2-Thenoyltrifluoroacetone, succinate dehydrogenase and, 246, 250, 255 Thermua aquaticus, glyceraldehyde-3phosphate dehydrogenase of, 2, 4,6, 20, 21, 22 Thiamine pyrophosphate, lipoamide dehydrogenase and, 126 Thiobacilli, adenylyl sulfate reductases in, 282 Thiobacillus denitrificans adenylyl sulfate reductase of, 283 sulfate reduction by, 281 Thiobacillus thiooxidans, sulfate reduction by, 281 Thiobacillus thioparus adenylyl sulfate reductase of, 283, 284 sulfate reduction by, 281 Thiocapsa roseopersicina, adenylyl sulfate reductase of, 282,283, 285-286 Thiocyanate catalase and, 387 small NADH dehydrogenases and, 192 succinate dehydrogenase and, 232 Thiol(s) adenylyl sulfate reductase and, 283 catalase and, 399-400, 401 cytochrome bs reductase and, 160-161, 162-164 lipoamide dehydrogenase, 118, 119, 120, 123 nitrite reductase and, 275, 277 Thiol groups, see also Sulfhydryl groups cytochrome c peroxidase and, 348 glutathione reductase, 141-142 NADH dehydrogenases, 203-204 succinate dehydrogenase, 245-246 ubiquinone reductase, 188, 203

Thionicotinamide adenine dmucleotide glyceraldehyde-3-phosphate dehydrogenase and, 30 transhydrogenase and, 57,59,60,6142, 69 Thioredoxin Escherichia coli, general properties, 144-145 nature of, 92, 93 Thioredoxin reductase amino acid composition, 102 cystine residues, 104 light-activated reduction-neutral semiquinone, 147-148 mechanism, 94, 98-99 metabolic functions, 142-144 reaction catalyzed, 92 reduced states, mechanism, 145-147 specificity of, 92-93 coenzymes and, 94 specificity of, 144 Threonine residues, lipoamide dehydrogenase, 105 Threose 2,4-diphosphate, glyceraldehyde3-phosphate dehydrogenase and, 44 Thyroid gland, a-glycerophosphate dehydrogenase and, 259 Tiron NADH dehydrogenase and, 206 succinate dehydrogenase and, 246 a-Tocopherylquinone, ubiquinone reductase and, 181-182 Torulopsis nitratophila, nitrite reductase of, 275, 276 Transacetylase dihydrolipoate and, 108 transhydrogenase and, 55 Transsuccinylase, dihydrolipoate and, 108 Tribromoacetate, small NADH dehydrogenases and, 192 Trichloroacetate small NADH dehydrogenases and, 192 succinate dehydrogenase and, 227,232, 234 Trideuteromethanol, catalase and, 404 Trifluoroacetate, small NADH dehydrogenases and, 192 2,2,2-Trifluoromethyl ethanol, catalase and, 401

457

SUBJECT INDEX

Triiodothyronine a-glycerophosphate dehydrogenase and, 259 transhydrogenase and, 70 Triton X-100 cytochrome c oxidase and, 313 o-lactate dehydrogenase and, 270-271 NADH dehydrogenase and, 187, 188189 succinate dehydrogenase and, 234 ubiquinone reductase and, 182-183 yeast NADH dehydrogenase and, 218 Trout, glyceraldehyde-3-phosphate dehydrogenase of, 26 Trypsin cytochrome b2 and, 266-267 cytochrome b5 reductase and, 164 cytochrome P-450 reductase and, 166, 167,169, 170-171, 172 transhydrogenases and, 58,65,71,212, 213, 296 yeast NADH dehydrogenase and, 217 Tryptophan residues cytochrome b, reductase, 164 cytochrome c peroxidase, 350,355 glyceraldehyde-3-phosphate dehydrogenase, 29 lipoamide dehydrogenase, 126 reductases and oxidases, 101 thioredoxin, 93 Turnover number, succinate dehydrogenase, 236-237 Tyrosine residues cytochrome b5 reductase, 163 cytochrome c peroxidase, 355 glyceraldehyde-3-phosphate dehydrogenase, 21 modification of, 22-23 reductases, 101

U Ubiquinone(s1 choline dehydrogenase and, 262 energy conservation and, 214 a-glycerophosphate dehydrogenase and, 258 large NADH dehydrogenase and, 201 small NADH dehydrogenases and, 194, 196, 206

solubility, ubiquinone reductase and, 180-181, 182 succinate dehydrogenase and, 224,225, 236,239,243,245,246,247,244-251 yeast NADH dehydrogenase and, 218 Ubiquinone reductase, see under Nicotinamide adenine dinucleotide Urea glyceraldehyde-3-phosphate dehydrogenase and, 21,24 L-lactate dehydrogenase and, 269 lipoamide dehydrogenase and, 122,124 nitrite reductase and, 279 small NADH dehydrogenase and, 203 succinate dehydrogenase and, 227 sulfite reductase and, 290,291 transhydrogenase and, 57

v Valence changes, cytochrome c oxidase, 331-332 Valine residues, disulfide oxidoreductases and, 104,105 Valinomycin, transhydrogenase and, 72 Venom, cytochrome b5 reductase and, 154 Vibro cholinicus, sulfate reduction by, 281 Vitamin K,, ubiquinone reductase and, 181, 182

W Water, catalase and, 374,385, 387

X Xanthine oxidase, cytochrome P-450 reductase and, 169 X-ray(s) glyceraldehyde-3-phosphate dehydrogenase apoenzyme structure, 19-20 holoenzyme structure, 9-19 Y

Yeast, see also Saccharomyces cytochrome c oxidase of, 311

458 cytochrome c peroxidase in, 347 glutathione reductase of, 94, 102, 138 diasociation constants, 135 kinetics, 140-141 substrates, 132 glyceraldehyde-3-phosphate dehydrogenase of, 6-8,18,26 amino acid modification, 22,23 half-site reactivity, 37 hybrids of, 26-27 pyridine nucleotide and, 28,31-33

SUBJECT INDEX

lipoamide dehydrogenase of, 112,117, 126 sulfate reduction in, 281 thioredoxin reductase of, 102

Z Zinc 2-hydroxyacid dehydrogenase and, 272-273 n-lactate dehydrogenase and, 270, 271-272

Topical Subject Index VOLUMES I-XI11 A

Acetamidyllysine residues, proteinase inhibitors, 111, 451452 Acetate: coenzyme A lipase catalytic properties cation requirements, X, 479-480 estimates of substrate affinity, X, 481 formation of enzyme-bound acetyl adenyla te, X, 478 selective modification of amino acid residues, X, 480-481 steady state kinetics and reation mechanism, X, 481-483 substrates and inhibitors, X, 477-478 molecular properties, X, 474-475 Acetoacetate decarboxylase historical background, VI, 255-256 inhibition studies borohydride, VI, 267-269 p-chloromercuriphenyl sulfonate, VI, 269-270

8-diketones, VI, 265-266 hydrogen cyanide, VI, 267 monovalent anions, VI, 266-267 2-oxopropane sulfonate, VI, 264 kinetic properties, VI, 263-264 mechanism, VI, 261-263 properties assay, VI, 256-257 latency, VI, 258-259 molecular weight, subunits and amino acid composition, VI, 260261

purification, VI, 257-258 stability, VI, 261

Acetylcholinesterase acceleration, V, 111-114 esteratic site, V, 95-97 historical background, V, 87-88 inhibitors anionic site, V, 98-100 esteratic site, V, 100-110 fluoride, V, 110-111 physical properties, V, 90-93 purification, V, 89 substrate binding, anionic site, V, 93-95 Acetyl coenzyme A-acyl carrier protein transacylase catalytic properties assays, VIII, 187 mechanism, VIII, 187-188 p H optimum and substrate specificity, VIII, 187 historical background, distribution and metabolic significance, VIII, 185186

molecular properties, VIII, 186-187 Acetyl coenzyme A carboxylase distribution, VI, 54-56 historical background and metabolic significance, VI, 53-54 molecular characteristics, VI, 58-59 reaction catalyzed, VI, 53 regulation of, VI, 79-82 substrate specificity, VI, 56-58 subunit structure and function active subunits in Escherichia coli, VI, 60-64 biotin carboxylase, VI, 70-71 biotin carboxyl carrier protein, VI, 64-70 459

460

TOPICAL SUBJECT INDEX

reconstitution and, VI, 72-78 structure of liver and wheat enzymes, VI, 78-79 transcarboxylase, VI, 71-72 N-Acetyl-n-glucosamine repimerase, properties, VI, 377-378 N-Acetyl-n-glucosamine 6-phosphate 2'epimerase, properties, VI, 377-378 N-Acetylglutamate-5-phosphotransferase allosteric inhibition kinetics, IX, 516-518 temperature effect, IX, 519-520 catalytic reaction assays, IX, 514-515 kinetics, IX, 515-516 pH optima and activating ions, IX, 514

stoichiometry, IX, 513-514 substrate specificity, IX, 514-515 historical background, IX, 511412 purification allosteric enzyme of Chlamydomonas, IX, 513 nonallosteric enzyme of Escherichia, I X , 513 N-Acetylneuraminate aldolase, properties, VII, 298-299 0-Acetylserine sulfhydrase, properties, VII, 54 Acidic nuclear protein kinases, properties, VIII, 580 Acid phosphatase(s) amebic, IV, 498 assay, IV, 457 problems, IV, 454 bone, IV, 496-497 distribution, IV, 450 Drosophila melanogaster, IV, 498 electrophoretic behavior, IV, 454455, 468-469

Escherichia coli, IV, 498 functional groups and group reagents iodination, IV, 469-471 sulfhydryl groups, IV, 469 tyrosine and tryptophan, IV, 471-472 Gaucher, IV, 496 general, IV, 455457 historical, IV, 450 kine tics fluoride inhibition, IV, 459-462

a-hydroxycarboxylic acid inhibition, IV, 462465 ion effects, IV, 466 pH and substrate effects, IV, 457-458 surface inactivation, IV, 459 liver bovine, IV, 491493 mouse, IV, 489-491 rat, IV, 484-489 Neurospora crassa, IV, 497 physical properties, IV, 476 plant, IV, 497 preparation, IV, 466-468 red cell general properties, IV, 477 purification and separation of genetic types, IV, 477-484 Saccharomyces, IV, 497 serum, IV, 495-496 specificity, alkaline phosphatase and, IV, 450-454 spleen, IV, 493495 staphylococcal, IV, 498 transphosphorylation, IV, 472473 use as reagent, IV, 473-476 Acid proteinase(s) pepsinlike chemical properties, 111, 728-730 distribution and isolation, 111, 724728

enzymic properties, 111, 734-740 physical properties, 111, 731-733 renninlike, 111, 740-741 distribution and isolation, 111, 741742

enzymic properties, 111, 743-744 physical and chemical properties, 111, 742-743 Aconitase catalytic properties assay, V, 423 equilibrium concentrations, V, 424 mechanism of action, V, 433-439 pH optima, V, 423-424 single vs. dual catalytic site, V, 432-433

specificity, V, 4-22 cofactors ferrous iron requirement, V, 422 role of reducing agent, V, 423

TOPICAL SUBJECT INDEX

function, V, 413414 historical background, V, 414-415 inhibitors fluorocitrate, V, 4 W 3 0 iron-binding agents, V, 428 other carboxylic acids, V, 430-431 other inhibitors, V, 431432 intracellular distribution, V, 416-417 kinetics, latent period, V, 424425 Michaelis constants, V, 425426 relative reaction rates, V, 425 scheme, V, 426-428 mechanism of, 11, 302-304 molecular properties factors affecting stability, V, 419420 physiochemical properties, V, 418-419 purification, V, 417418 occurrence, V, 415416 reactions catalyzed, stereospecificity, 11, 164-168 role of metals, 11, 516-518 Aconitate, cis-trans isomerization, VI, 394-395 Active site(s) alkaline phosphatase, IV, 404-406 amino acid decarboxylases, VI, 245-248 8-aminolevulinate dehydratase, VII, 331433 aspartate transcarbamylase, IX, 262268 carbonic anhydrase, V, 617422, 643646 carboxylesterases, V, 61434 catalase, XIII, 3694388 chemical modification, I, 194-196 a-chymotrypsin, 111, 1!%-202 y-chymotrypsin, 111, 202-204 chymotrypsinogen, 111, 179-182 creatine kinase, VIII, 439442 deoxyribonuclease I, IV, 297-299 enolase components, V, 532-534 mapping with substrate analogs, V, 526-529 number, V, 530-532 fumarase, affinity labeling, V, 563-564 p-galactosidase, VII, 6574358 glucose-6-phosphate isomerase, VI, 285-287

461 glutamine synthetase, X, 720-733 j3-hydroxydecanoyl thioester dehydrase, V, 453455 invertase, V, 300301 lipase, VII, 593-595 papain, 111, 496499 activation, 111, 511414 chemical modification, 111, 51.5-516 half-cystine content, 111, 509-511 location of thiol group, 111, 514 staphylococcal nuclease, IV, 195-196 streptococcal proteinase, 111, 626-627 subtilisin, 111, 553-560, 575-584 thiolase, VII, 404-405 triosephosphate isomerase, VI, 330-333 trypsin, 111, 260-262 urease, IV, 20-21 Active-site-directed reagents as adjuncts to physical methods crystallography, I, 143 other spectroscopic methods, I, 145 spin labels, I, 143-145 characterization of functional groups and, I, 142 chymotrypsin and chloromethyl ketone from tosylphenylalanine, I, 94-96 other alkylating agents, I, 97-99 other neutral proteases, I, 102-103 other studies, I, 99-102 consecutive covalent modifications displacement, I, 115117 elimination, I, 115-116 intramolecular alkylation, I, 113-114 photolysis, I, 114-115 heme proteins and, I, 142 hydrolytic enzymes and, amidases, I, 130-131 deaminases, 1, 127-128 esterases, I, 124-127 glycosidases, I, 128-129 nucleases, I, 130 proteolytic enzymes, I, 118-124 lyases and carbonic anhydrase, I, 139-140 3-deoxy-o-arabino-heptulosonate-7phosphate synthase, I, 139 fumarase, I, 140-141 6-hydroxydecanoyl thioester dehydrase, I, 141

TOPICAL SUBJECT INDEX

2-ke to-3-deoxy-6-phosphogluconic aldolase, I, 139 oxidoreductases and alcohol dehydrogenase, I, 136 glutamate dehydrogenase, I, 137 guanosine5’-phosphate reductase, I, 137 inosinic acid dehydrogenase, I, 137138 lactate dehydrogenase, I, 136 as selective enzyme inhibitors, I, 146 transferases and acyltransferases, I, 134-135 adenylyltransferase, I, 135 amidotransferase, I, 131-134 triosephosphate isomerase and, I, 138 trypsin and related enzymes, I, 103112 Acyl carrier protein distribution and intracellular Iocalization, VIII, 158-164 function in fatty acid biosynthesis, VIII, 164-165 molecular properties physical properties, VIII, 166 prosthetic group and primary sequence, VIII, 166-170 structure-activity relationships, VIII, 170-173 historical background, VIII, 155-158 prosthetic group synthesis and turnover, VIII, 173-176 Acyl coenzyme A ligases assays, VII, 411 general aspects of reaction, VII, 409410 reaction catalyzed, VII, 407-409 Acyl transfer carboxyl anions and, 11, 226-235 tertiary amino groups and, 11, 235-238 Acyltransferases, modification of, I, 134135 Adenine aminohydrolase, IV, 51-54 Adenine nucleotide aminohydrolase, IV, 75-76 Adenosine phosphodiester derivative biosynthesis adenyl cyclase, VIII, 26-27 adenylylation of glycoside antibiotics, VIII, 27-30

ribonucleic acid biosynthesis, VIII, 20-26 Adenosine aminohydrolase catalytic properties mechanism, IV, 5943 nature of active site, IV, 58-59 reaction parameters, IV, 5 6 5 7 molecular properties chemical and physical, IV, 55-56 purification, IV, 54-55 physiological function, IV, 63-64 Adenosine aminohydrolase (nonspecific), IV, 73-75 Adenosine diphosphate synthesis of derivatives adenine-myonic acid dinucleotide and adenylyl diphosphoglycerate, VIII, 33-35 adenosine diphosphate glucose synthesis, VIII, 3 2 3 3 general features, VIII, 3&32 Adenosine diphosphate sulfurylase, X, 663-665 Adenosine diphosphoryl glucose pyrophosphorylase activator, general effects, VIII, 77-78 Aeromonas formicam, VIII, 108-109 Chlorella pyrenoidosa, VIII, 90 classification, VIII, 75-77 enterobacteriaceae activator effects on kinetic parameters, VIII, 97-99 activator-inhibitor interaction, VIII, 99-100 energy charge and, VIII, 104-107 inhibitor effect on kinetic constants, VIII, 101-102 manganese effects, VIII, 102-104 Entner-Duodoroff pathway and, VIII, 78-81 Escherichia coli, VIII, 102-107, 109-110 mutantb) SG5 and CL1138, VIII, 110-114 mutantb) SG14, VIII, 115-117 general background, VIII, 73-75 nonchlorophyllous plant t i m e , VIII, 93-94 other leaves, VIII, 89-90

463

TOPICAL SUBJECT INDEX

3-phosphoglycerate activation and phosphate inhibition of, VIII, 9192

physical properties, VIII, 117-119 R hodospirillum rubrum, VIII, 81-83 reaction mechanism, VIII, 86 temperature and, VIII, 83-86 Serratia marcescens, VIII, 107-108 spinach leaf, VIII, -9 Adenosine kinase assay, IX, 51-52 distribution and purification, IX, 51 kinetic and molecular properties, IX, 52-53

substrate specificity, IX, 53-54 Adenosine monophosphate, fructose-l,& diphosphatase and, IV, 618-620,627628, 636-637

Adenosine triphosphatase(s) Alcaligenes jaecalis membrane catalytic properties, X, 429 molecular properties, X, 428 solubilization and purification, X, 428

Bacillus megaterium membrane catalytic properties, X, 427-428 purification, X, 426 reassembly, X, 428 release from membranes, X, 426 size, amino acid composition and morphology, X, 427 subunits, X, 427 Bacillus stearothermophilus membrane properties, X, 425-426 solubilization and purification, X, 425

bacterial membrane, X, 396-400 other bacteria, X, 416 Streptococcus faecalis, X, 400, 416429

chloroplast Euglena gracilis, X, 394 spinach, X, 389-394 Escherichia coli membrane deficient mutants, X, 419421 molecular weight and catalytic properties, X, 418 release from membranes, X, 416-418 subunit composition, X, 418-419 function of, X, 375-377

Micrococcus lysodeikticua membrane electron microscopy, X, 422 FMC and, X, 424425 interaction with antibodies, X, 423 localization, X, 422423 release from, X, 421-422 size and catalytic properties, X, 422 subunits, X, 423-424 mitochondria1 assay, X, 377 beef heart, X, 377-386 rat liver, X, 387-389 yeast, X, 386-387 Rhodopseudoomonas spheroides membrane, X, 429 sarcoplasmic membrane calcium-dependent, X, 445-450 calcium-independent, X, 444-445 historical background, X, 432-434 Streptococcus jaecalis membrane active transport and, X, 414-416 amino acid composition, X, 405-406 carbodiimide-resistant mutants, X, 413-414

dicyclohexylcarbodiimide inhibition, X, 411-413 electron microscopy, X, 405 isolation of membranes, X, 400 kinetics, X, 408-409 molecular weight, X, 404-405 nectin and, X, 410411 purification, X, 402404 reassembly, X, 409-410 release from membranes, X, 400-402 subunits, X, 406408 Adenosine triphosphate calcium-efflux dependent net formation, X, 457458 phosphate exchange, X, 458-459 metal complexes, electronic structure, 11, 479-481 Adenosine triphosphate citrate lyase, VII, 368-369 assay and isolation, VII, 369-370 catalytic properties control, VII, 372 equilibrium and kinetics, VII, 371372

specificity, VII, 371 stereospecificity, VII, 372-373

464

TOPICAL SUBJECT INDEX

molecular properties cofactors, VII, 370 inhibitors, VII, 370-371 molecular weight, VII, 370 stability, VII, 371 sulfhydryl groups, VII, 370 reaction mechanism adenosine diphosphate-adenosine triphosphate exchange, VII, 373 citryl coenzyme A aa intermediary, VII, 376 citryl-enzyme as intermediary, VII, 375-376

citryl phosphate as intermediary, VII, 374-375 oxygen transfer to orthophosphate, . VII, 373 phosphoryl-enzyme as intermediary, VII, 373-374 reaction scheme, VII, 377 Adenosine triphosphate sulfurylase general properties, X, 656-658 mechanism, X, 658-662 occurrence and purification, X, 652654

reactions and assay, X, 655-656 Adenosyltransferase(s) , general background, VIII, 121-123, 152-154 Adenylate energy charge, metabolic regulation and, I, 470-476 Adenylate kinase biological aspects distribution, VIII, 279-282 function, VIII, 285-288 genetics and disease, VIII, 282-284 catalytic properties assay, VIII, 300-301 equilibrium constants, VIII, 302 mechanism, VIII, 302-305 metal requirement, VIII, 297-298 nucleotide specificity, VIII, 298-300 molecular properties composition, VIII, 291-293 physical properties, VIII, 295-297 preparation and purity, VIII, 288291

reactive groups, VIII, 293-296 role of metals in mechanism, 11,502 Adenyl cyclase, properties, VIII, 26-27

5’-Adenylic acid aminohydrolase catalytic properties kinetics, IV, 66-70 mechanism, IV, 70-71 physiological function, IV, 71-73 specificity, IV, 66 molecular characteristics chemical and physical properties, IV, 65-66

purification and homogeneity, IV, 6465

Adenylosuccinase 5-amino-4-imidazole-N-succinocarboxamide ribonucleotide and, VII, 182-183

mechanism of action, VII, 196-197 Neurospora, VII, 191-192 assay procedures, VII, 192 catalytic properties, VII, 192 purification, VII, 192 subunit structure, VII, 192-193 sterospecificity of additional or elimiination, VII, 195 yeast, VII, 185-186 assay procedure, VII, 187 function and distribution, VII, 186187

pH optima and equilibrium, VII, 191 purification, VII, 187 substrate affinity and product inhibition, VII, 189-190 substrate specificity, VII, 187-188 sulfhydryl reagents and, VII, 188189

Adenylylsulfate kinase, X, 662-663 Adenylyl sulfate reductase(s), properties, XIII, 279-286 Adenylyltransferase, modification of, I, 135

Adenylyl transfer reactions, general background, VIII, 1-6 Adipose tissue, hormone sensitive lipase, VII, 609-610 Adrenal gland, glycogen synthetase of, IX, 353 Aerobacter aerogenes, pullulanase of, V, 195-201

Aerobacter cloacae, phage polysaccharide depolymerase, V, 398

465

TOPICAL SUBJECT INDEX

Aeromonas formicans, adenosine diphosphoryl glucose pyrophosphorylase of, VIII, 108-109 Affinity labeling, chemical modification in general and, 91-94 Agaricaceae, y-glutamyltransferase of, IV, 95-96 Alanine dehydrogenase, regulation of, I, 443444 Alanine racemase, spore germination and, VI, 506 Alcaligenes fueculis, membrane adenosine triphosphatase of, X, 428-429 Alcohol dehydrogenase(s1 chemical modifications, XI, 141-145 arginine residues, XI, 179 cobalt enzyme, XI, 180 cysteine residues, XI, 176177 denaturation, XI, 180-181 histidine residues, XI, 177-179 manganese enzyme, XI, 180 properties, XI, 179-180 uncharacterized, XI, 179 coenzyme analogs and, XI, 150-152 comparisons evolutionary aspects, XI, 140-141 structural and functional aspects, XI, 136-140 denaturation studies, XI, 147-148 fluorescence and phosphorescence, XI, 148-150

functional aspects, activity in ethanol metabolism, XI, 106

physiological substrates, XI, 105-106 gene duplication and, I, 309-311 horse liver multiple molecular forms, XI, 107108

primary structure, XI, 113-116 human liver, multiple molecular forms, XI, 109110

primary structure, XI, 116 inhibitor binding, binary complexes, XI, 152-157 ternary complex, XI, 158-160

inhibitor studies coenzyme competitive inhibitors, XI, 181-182 others, XI, 182-183 kinetic aspects coenzyme binding, XI, 160-163, 183184

dismutase reaction, XI, 166 half-site reactivity, XI, 166-167 ordered mechanism, XI, 165-166 reaction mechanism, XI, 185-186 substrate binding, XI, 163-165 substrate specificity; XI, 184-185 liver, XI, 5 6 5 7 kinetic studies, XI, 20-22 mechanism for catalysis, XI, 168-171 metal content changes, XI, 145-147 modification of, I, 136 other sources, XI, 186-187 bacterial, XI, 187-188 insect, XI, 189-190 plant, XI, 188-189 rat liver, multiple molecular forms, XI, 111112

primary structure, XI, 114-117 tertiary structure crystallization and preliminary Xray studies, XI, 117-118 electron density maps, XI, 118-119 three-dimensional structure, XI, 1% 136

yeast chemical modifications, XI, 176181 inhibitor studies, XI, 181-183 kinetic aspects, XI, 22-23, 183-186 primary structure, XI, 173-176 purification and molecular properties, XI, 171-173 Aldehyde oxidase, metal complexes and, 11, 533-534 Aldolase(s) historical review, VII, 213-215 properties in bacteria and fungi, VII, 215-216

Schiff bases and, 11, 359-380 yeast, role of metals, 11, 515-516 Aldolase(s) (Metallo) distribution and general properties catalytic properties, VII, 253-254

466

TOPICAL SUBJECT INDEX

multiple forms in microorganisms,

VII, 255-256 purification and metal content, VII, 254-255

reaction mechanism other functional groups, VII, 257258

possible molecular homology in yeast and muscle enzymes, VII, 258

role of metal ion, VII, 256-257 Aldolase(s) (Schiff base-forming) developmental aspects embryonic tissues and, VII, 249-251 modification of structure, VII, 252253

tumors and, VII, 251-252 mechanism of catalysis isotope exchange reactions, VII, 216-217

Shiff base formation, VII, 217-219 organ-specific molecular properties and subunit structure, VII, 221-223 nonidentical subunits and microheterogeneity, VII, 223-224 occurrence of isozymes in vertebrate tissues, VII, 220-221 specificity, VII, 219-220 phylogenetic studies comparison of active site peptides,

VII, 248-249 isolation from other species, VII, 24k248

rabbit liver and brain isolation and general properties, VII, 241-243

primary structure of active site, VII, 243-244

rabbit muscle active site sequence, VII, 236 amino acid composition, VII, 224 functional groups, VII, 224-232 isolation of crystalline enzyme, VII, 224

primary structure, VII, 236-239 reaction mechanism, VII, 232-236 X-ray crystallography and electron microscopy, VII, 239-241

Aldose-ketose isomerases general considerations, VI, 271-272 nonphosphorylated sugars and, VI, 340-354

Algae blue green, fructose-l,6diphosphatases of, IV, 640-642 Alkaline phosphatase active sites, number of, IV, 404-406 chelating agents and, IV,426-427 chemical modification, IV, 391-392 arginine, IV, 390 histidine, IV, 390 leucine, IV, 390 methionine and cystine, IV, 390391 phenylalanine, IV, 389-390 tryptophan, IV, 390 competitive inhibitors, IV, 394-396 composition, IV, 423-425 analysis, IV, 378-380 sequence work, IV, 380 crystal structure, IV, 389 distribution, IV, 374-376 histochemical and gel localization, IV, 433-434

historical background, IV, 373374 in vitro assay, IV, 432-433 isozymes, IV, 384387 kinetic studies, IV, 409-416 factors affecting, IV, 434-436 inhibition and, IV, 442443 Kim and 'Vim.,,IV, 436-439 metal ions and, IV, 440-441 phosphoryl enzyme formation and,

IV, 439 transferase activity, IV, 439-440 mammalian assay techniques, IV, 432-434 chemical modification, IV, 427-428 distribution, IV, 420-421 function, IV, 421-422 general survey, IV, 417-420 kinetic studies, IV, 434-443 mechanism, IV; 443-447 physical properties, IV, 423-427 purification procedures, IV, 422-423 reaction catalyzed, IV, 430-432 substrate specificity, IV, 428-430 mutations and, I, 251-254 phosphoryl enzyme, IV, 396-401

TOPICAL SUBJECT INDEX

physical properties, IV, 387-388 purification, IV, 377-378 specificity, IV, 392-394 stability, IV, 425426 subunits, IV, 380-384 transphosphorylation and, IV, 406-409 zinc and, IV, 401404 Alkaline proteinase(s) diisopropylfluorophosphate-sensitive, 111, 744-745 chemical properties, 111, 749-754 distribution and isolation, 111, 745749 enzymic properties, 111, 758-763 keratinase, 111, 763-765 physical properties, 111, 754-758 8-Alkyl-L-cysteine sulfoxide lyase, properties, VII, 52 Alkylsulfatase(s), V, 15 Allylases, proton shifts and, 11, 299302 D-Altronate dehydrase, properties, V, 579 Ameba, acid phosphatase of, IV, 498 Amidases, modification of, I, 130-131 Amidinotransferase(s), reactions catalyzed, IX, 497-498 Amidotransferases, modification of, I, 131-134 Amine oxidase (s) , active site, substrate interaction, XII, 524-525 catalytic mechanism, XII, 525-526 definition and classification, XII, 511513 inhibitor reactions, XII, 523-524 metabolic function, XII, 513 microsomal, properties of, XII, 228. 230 other copper-containing, XII, 526-527 prosthetic groups, XII, 519-523 purification, molecular weight and substrate specificity, XII, 513-518 spectral properties, XII, 518-519 Aminoacetone, formation of, VII, 355 Amino acid(s) activation, VIII, 6-11 acyl carrier protein sequence, VIII, 166-170 amylase composition, microbial, V, 239-244 analogs, mutations and, I, 262265

aromatic aspartate transcarbamylase, IX, 267-268 biosynthesis, I, 228-237 asparaginase composition, IV, 111-113 aspartokinase(s) composition, VIII, 520, 542, 543 carboxylesterase composition, V, 52-53 creatine kinase composition, VIII, 390-392 degradation, VI, 504-506 elastase sequence, 111, 341-343 fumarase and, V, 544-545 a-glucan phosphorylase composition, VII, 446-447 glucose-6-phosphate isomerase composition, VI, 27S279 guanidino kinase composition, VIII, 469-470 function, VIII, 477-482 hexokinase composition, IX, 41 inorganic pyrophosphatase and, IV, 512514 microbial proteinases and, 111, 749751, 770-772 pancreatic ribonuclease sequence, IV, 653-654 papain composition and sequence, IV, 507-509 pepsin composition, 111, 128-130 sequence, 111, 1W133 phospholipase A, content, V, 80-81 sequence, V, 81-82 prothrombin composition, 111, 313 pyridoxal reactions with, 11,339445 sequences, carboxypeptidase B, 111, 64-66 streptococcal proteinase active site, 111, 626-627 composition, 111, 624-625 N- and C-terminal, 11, 625-626 thrombin composition, 111, 285-286 sequences, 111, 287-290 Amino acid decarboxylases active site absorption spectra of pyridoxal phosphatedependent, VI, 245-247

468

TOPICAL SUBJECT INDEX

pyridoxal phosphate binding site, VI, 247-248

distribution and general properties, VI, 224-237

general considerations, VI, 217-219 mechanism of action glycine decarboxylase, VI, 240-241 pyridoxal phosphate a-decarboxylation, VI, 237-240 pyridoxal phosphate p-decarboxylation, VI, 241-243 pyruvate-containing, VI, 244 metabolic importance bacterial, VI, 219-221 mammalian and plant, VI, 221-224 subunit structure, VI, 248-253 n-Amino acid oxidase, molecular properties and kinetic mechanism, XII, 445-456

L-Amino acid oxidase, molecular properties and kinetic mechanism, XII, 456-461

Amino acid racemases assay methods coupling to L- or n-specific enzymes, VI, 489 polarimetric, VI, 489-490 cofactors flavin, VI, 496 metal ions, VI, 496-497 no pyridoxal phosphate, VI, 495 pyridoxal phosphate, VI, 494-495 status uncertain, VI, 495-496 equilibrium position, VI, 490-491 history and survey, VI, 481488 kinetic features, VI, 491494 mechanism of action aminoacyl complex and, VI, 501-502 nonpyridoxal enzymes, VI, 498-500 pyridoxal enzymes, VI, 497-498 physiological aspects n-amino acids in animal tissues, VI, 508-507

biosynthesis and degradation of free amino acids, VI, 504-506 cell wall biosynthesis, VI, 502-503 peptide antibiotic biosynthesis, VI, 503-504

spore germination, VI, 506 substrate specificity, VI, 490

Aminoacyl transfer ribonucleic acid, enzymic deacylation, X, 509-510 Aminoacyltransfer ribonucleic acid synthetases amino acid activation, X, 505-506 assay, X, 506 binding parameters, X , 507-508 reaction product, X, 506 substrate specificity, X, 506607 amino acid biosynthesis and, X, 536 chemical properties affinity labeling, X, 505 amino acid composition, X, 503 chemical modification, X, 505 proteolytic modification, X, 504 terminal amino acids and sequence analysis, X, 503-504 general considerations, X, 489492 genetics of, X, 529-534 mechanism of reaction general, X, 510-511, 517-515 isoleucyltransfer ribonucleic acid synthetase, X, 511-517 occurrence and distribution, X, 492-491 purification, X, 494-496 Crystallization, X, 496-497 regulation of biosynthesis, X, 534-535 size and subunit composition, X, 502503

multichain with dissimilar subunits, X, 502 multichain with similar subunits, X, 499-502

single chain, X, 497499 Amino group(s) aspartate transcarbamylase, IX, 265267

chemical modification, I, 175 elastase, 111, 386-373 ribonuclease, IV, 677-682 tertiary, acyltransfer to, 11, 235-238 trypsin, 111, 269-270 w-Amino group migrations, VI, 547-548 reaction mechanisms cobamide coenzyme as hydrogen carrier, VI, 560-561 general considerations, VI, 559-560 hydrogen transfer reaction, VI, 561 role in bacterial fermentations, VI, 562-563

TOPICAL SUBJECT INDEX

Amino group transfer basic chemical features congruent nonenzymic models, IX, 387-391

general characteristics of intermediate steps, IX, 391-392 formally similar processes, IX, 384387

historical background, IX, 379-381 other substrates and, IX, 462463 w-amino and 0-0x0 acids, IX, 473474

glutamate-oxoglutarate or aspartateoxalacetate, IX, 463473 noncarboxylic acids, IX, 475-476 as side reactions, IX, 47-80 two a-amino-a-oxomonocarboxylic acids, IX, 473 recent developments, IX, 381-384 5-Amino-4-imidazole carboxamide ribonucleotide transformylase, properties, IX, 204-205 5-Amino-4-imidarole-N-succinocarboxamide ribonucleotide cleaving activity, VII, 183-184 assay, VII, 184-185 enzymic properties, VII, 185 function and distribution, VII, 184 purification, VII, 185 8-Aminolevulinate dehydratase active site, nature of, VII, 331-333 catalytic properties assay, VII, 330 kinetics, VII, 330-331 pH optima, VII, 331 cation requirements, VII, 328-330 mechanism of porphobilinogen synthesis, VII, 333-337 molecular weight, VII, 324326 aggregation and, VII, 326-327 reaction catalyzed, VII, 323-324 Schiff base and, 11, 361-362 subunits, quaternary structure and, VII, 327-328 6-Aminolevulinate synthetase analogous reactions, VII, 355356 cofactors, substrate specificity and kinetic constants, VII, 348-349 historical background, VII, 339-340

inhibitors amino acids and derivatives, VII, 349-350

carboxylic acids, VII, 351 sulfhydryl reagents, metals and porphyrins, VII, 350-351 mechanism, VII, 351-355 metabolic significance, VII, 343-344 molecular properties assay, VII, 347 high and low activity forms, VII, 344-315

isolation, VII, 346-347 stability, VII, 344 occurrence, VII, 341-342 Aminopeptidase A, 111, 111-112 Aminopeptidase B, 111, 112-113 Aminopeptidase M, 111, 102-105 Aminopeptidase P, 111, 115-116 Aminotripeptidase, 111, 117-118 Ammonia elimination enzymic general considerations, VII, 79-88 nomenclature, VII, 77-79 free energy, VII, 88-90 interpretation, VII, 92-94 ionic species, VII, 90-91 standard states, VII, 91-92 models for steric course, carbanion intermediate, VII, 114-116 concerted elimination, VII, 110-114 stereochernistry, VII, 94-95 configurations of amino acid chiral centers, VII, 95-98 configurations and conformations of olefinic products, VII, 109-110 specificity toward amino acid prochiral centers, VII, 99-109 Amphibolic pathways input signals and, I, 476-479 interaction with biosynthetic pathways, I, 481482 Amylase (8) action pattern effects of chain length, V, 173-182 hydrolysis and condensation specificity, V, 149-152 multiple attack, V, 165-173 single chain or multichain attack, V. 161-165

470

TOPICAL SUBJECT INDEX

subsite model, V, 154-161 substrate structural requirements, V, 152-154

amino acid content, V, 127 assay methods, V, 117-120 biosynthesis, genetics and control, V, 182-189

chemical modification, V, 129-132 classification, V, 115-117 mechanism of action, 140-149 microbial amino acid composition, V, 239-243 assay, V, 265-266 calcium and, V, 247-250 carbohydrate components, V, 245-257 chemical modification, V, 261-263 denaturation and renaturation, V, 251-256

fragmentation, V, 257-258 mechanism, V, 268-271 mode of action, V, 266% molecular weights, V, 250-251 purification, V, 236-239 specificity, V, 263-265 sulfhydryl and disulfide groups, V, 244-245

terminal groups, V, 241-244 thermostable and acid stable, V, 258-261

origin, purification and molecular variants, V, 121-127 pH, temperature and salt effects, V, 132-140

size and shape, V, 128-129 subunits and multiple binding sites,

V, 127-128 Androstenolone sulfatase, V, 7-9 Anhydrochymotrypsin, elimination reactions and, I, 115-116 Animals, nuclear ribonucleic acid polymerase, X, 262-300 Anions, carbonic anhydrase and, V, 646-652, 658-660

Antibiotics amino glycoside, adenylylation, VIII, 27-30

Antithrombins, thrombin and, 111,305306

n-Arabinose isomerase, properties, VI, 346348

L-Arabinose isomerase, properties, VI, 348-349

Arabinose-bphosphate isomerase, properties, VI, 324-325 n-Arabonate dehydrases, properties, V, 582

L-Arabonate-n-fuconate dehydratases, properties, V, 581-582 Arene oxides, epoxidases and, VII, 211212

Arginine kinase, see also Guanidino kinases role of metals in mechanism, 11, 501502

Arginine monooxygenase, properties,

XII, 203-204 Arginine residues chemical modification, I, 174 chymotrypsinogen, 111, 176-179 ribonuclease, IV, 689490 Argininosuccinase (8) assay procedures, VII, 171 bovine kidney catalytic properties, VII, 179 purification, VII, 178-179 catalytic properties inhibitors, VII, 172 pH optimum and equilibrium, VII, 173

substrate specificity, VII, 172 function and distribution, VII, 170-171 historical background, VII, 169-170 mechanism of action, VII, 196 molecular structure subunit constitution, VII, 176-177 sulfhydryl groups and, VII, 177-178 Neurospora, catalytic and physical properties, VII, 180-181 number of binding sites, VII, 174-175 pea seeds, catalytic and physical properties, VII, 181-182 primary structure amino acid composition, VII, 179 antigenic properties, VII, 179 purification, VII, 171-172 regulation cooperative substrate effects, VII, 173-174

nucleotide stimulation, VII, 174

471

TOPICAL SUBJECT INDEX

reversible cold inactivation and subunit dissociation, control of, VII, 175 kinetics, VII, 175-176 thermodynamic constants, VII, 176 stereospecificity of addition or elimination, VII, 194 Argininosuccinate, synthesis, VIII, 38-39 Arthropoda, glycogen synthetase of, IX, 358-359

Arylsulfatase(s), V, 23 type I, V, 3 4 , 23-26 type 11, V, 4, 26-39 Asparaginase (s) amylase content, V, 127 Escherichia coli isolation, IV, 107-108 properties, IV, 109-116 guinea pig serum isolation, IV, 105-106 properties, IV, 106-107 occurrence, IV, 102-105 other, IV, 116-117 physiological properties, IV, 117-121 properties amino acid composition, IV, 111-113 general, IV, 109-110 structure, IV, 113-116 substrate specificity and inhibitor effects, IV, 110-111 Asparagine synthetase, X, 578-580 bacterial sources, X, 568-572 glutamine-dependent, X, 572578 historical background, X, 561-568 Aspartate metabolism, regulation of, I, 457459 pyruvate carboxylase and, VI, 31-33 Aspartate ammonia-lyase catalytic process, VII, 135-137 distribution, purification and kinetic properties, VII, 116-118 size and constitution, VII, 119-121 Aspartate :oxoglutarate aminotransferase(s1 isoenzymes and multiple subforms, IX, 393-398 pig heart and animal tissues coenzyme analogs and, IX, 429435 dynamic spatial aspects, IX, 465-462 kinetics, IX, 424-429

optical properties, IX, 407416 physical parameters and macromolecular structure, IX, 398-406 primary structure and functionally important groups, IX, 416424 stereochemistry and active site topography, IX, 451455 substrates, quasi substrates and inhibitors, IX, 435-451 Aspartate residues, chymotrypsin, 111, 235-236, 243

Aspartate transcarbamylase active site functional groups, IX, 262-263

amino groups, IX, 265-267 aromatic amino acids, IX, 267-268 histidine residues, IX, 267 sulfhydryl groups, IX, 263-265 allosteric effectors bindine site, IX, 270-273 properties in presence of effectors, IX, 269-270 bacterial, IX, 297-302 biosynthesis and genetics control of, IX, 295-297 location of genes, IX, 292293 two chains-single operon?, IX, 293-294

catalytic subunit primary structure, IX, 232-234 size and substructure, IX, 231-232 comparison of native enzyme with subunits, IX, 277-278 cooperative properties of modified enzyme modification with partial specificity, IX, 279-280 other modified enzymes, IX, 284-285 specifically modified, IX, 280-284 urea and p H effects, IX, 278 cooperative substrate binding, IX, 268-269

detailed subunit structure, IX, 239-243 fungal, IX, 302-306 induced conformational changes allosteric effectors, IX, 276-277 substrates and substrate analogs, IX, 275-276 isolation and characterization away procedures, IX, 228-230

TOPICAL SUBJECT INDEX

properties associated with regulation, IX, 227-228 purification, IX, 238 size and subunit composition, IX, 230-231

kinetics of ligand binding, I X , 285-287 mammalian, IX, 306-307 mechanism, IX, 243-282 mechanisms for cooperativity structural models, IX, 290-292 two-state major transition, IX, 287-290

plant, IX, 307-308 reconstitution importance of metals, IX, 237-238 metals other than zinc, IX, 238-239 methods, I X , 237 regulatory subunit metal binding site, IX, 236 primary structure, IX, 235 as regulatory protein, IX, 236-237 size and substructure, IX, 234-235 stoichiometry of ligand binding nucleotides, IX, 274-275 substrates and substrate analogs, IX, 273-274 Aspartokinase (s) assay of, VIII, 512-513 Escherichia coli, VIII, 513-544 historical background, VIII, 509-51 1 lysine-sensitive, adenylylation, VIII, 44-45

other coliform bacteria, VIII, 544 reaction catalyzed, VIII, 511-512 regulated by concerted feedback inhibition Bacillus polymyxa, VIII, 546-548 Bacillus stearothermophilus, VIII, 550

Bacillus subtilis, VIII, 548-550 other bacilli, YIII, 551 other genera, VIII, 552 other nonsulfur photosynthetic bacteria, VIII, 545446 pseudomonads, VIII, 551452 R hodopseudomonas capsulatus, VIII,

Aspartokinase I, VIII, 515-516 chemical properties, VIII, 520-522 conformational changes, VIII, 526-536 distribution of two activities on, VIII, 536-540

extinction coefficient, VIII, 517 kinetic parameters, VIII, 519-520 ligand binding, VIII, 523-525 molecular weight, VIII, 517-518 purification and criteria of homogeneity, VIII, 516 stability, VIII, 517 sulfhydryl titration effects, VIII, 525-526

tetrameric structure, VIII, 518-519 Aspartokinase I1 amino acid composition, VIII, 541-542 extinction coefficient, VIII, 541 kinetic parameters, VIII, 541 molecular weight, VIII, 541 purification and criteria of homogeneity, VIII, 540 stability, VIII, 540 subunit structure, VIII, 541 Aspartokinase I11 amino acid composition, VIII, $543 extinction coefficient, VIII, 542 inhibition, VIII, 544 kinetic parameters, VIII, 543 molecular weight, VIII, 543 purification and criteria of homogeneity, VIII, 542 Aspergillus, proteinases of, 111, 747, 769 Aspergillus nidulans molybdenum hydroxylases of, XII, 412414

Asymmetry molecular, notations of, 11, 129-134 Azotobacter phage polysaccharide depolymerase assay and purification, V, 397 properties, V, 397-398 Azotobacter agilis, glutaminase of, IV, 97-98

B

544-545

Rhodopseudomonas spheroides, VIII, 552-553

Saccharomyces cerevkiae, VIII, 553

Bacillus alkaline proteinases of, 111, 605-606 proteinases of, 111, 767-768

TOPICAL SUBJECT INDEX

Bacillus cereus, phospholipase C of, V, 83-84 Bacillus megalerium membrane adenosine triphosphatase, X, 426-428 phage G-induced lytic enzyme bound, V, 409-410 soluble, V, 408-409 Bacillus polymyxa, aspartokinase of, VIII, 546-548 Bacillus stearothermophilus aspartokinasc of, VIII, 550 membrane adenosine triphosphatase of, X, 425-426 phage lytic enzyme, V, 410411 Bacillus subtilis aspartokinase of, VIII, 548-550 deoxyribonucleic acid polymerases, physiological role, X, 143-144 extracellular ribonuclease of, IV, 239-240 intracellular ribonuclease of, IV, 240 phage-induced exonuclease, IV, 258-259 Bacteria, see also specific organisms alcohol dehydrogenase of, XI, 187-188 aldolases of, VII, 215-216 amino acid decarboxylases of, VI, 219-221 amylases catalytic properties, V, 263-271 molecular properties, V, 236-263 aspartate transcarbamylases of, IX, 297-302 deoxyribonucleic acid methyltransferases, IX, 190-192 deoxyribonucleic acid polymerases of, X, 119-144 elongation factors, X, 55-67 endonuclease of, IV, 259-270 exonucleases of, IV, 252-259 ferredoxins of, XII, 37-46 fructose-1,6-diphosphatasesof, IV, 639-640 glutamate dehydrogenases bacilli, XI, 332 Escherichia coli, XI, 332-333 others, XI, 333-334 hyaluronidases of, V, 313-314 neuraminidases of, V, 324-325

5‘-nucleotidase of, IV, 338-340 photosynthesis in, XI, 509-516 pro teases acid, 111, 723-744 diisopropylfluorophosphate sensitive, 111, 744-765 metal-chelator sensitive, 111, 765-786 other, 111, 786-795 respiratory chains denitrifiers, XI, 521426 inorganic reductants, XI, 519-521 less-mitochondrion-like, XI, 517-519 mitochondrion-like,’XI, 516-517 sulfate respiration and, XI, 526-534 ribonucleic acid polymerases background, X, 333-335 catalytic properties, X, 344-374 molecular properties, X, 335-344 Bacterial cell wall lytic protease Myxobacter, 111, 786-788 other, 111, 789-790 Sorangium, 111, 788-789 Bacteriophage A endolysin catalytic properties, V, 391-392 chemical properties, V, 388-391 physiochemical properties, V, 387-388 purification, V, 385-386 induced exonuclease and, IV, 253-254 Bacteriophage F series polysaccharide depolymerase catalytic properties and biological significance, V, 393 enzyme assay, V, 392 partial purification, V, 392-393 stability, V, 393 Bacteriophage G lytic enzyme bound, V, 409-410 soluble, V, 408-409 Bacteriophage N20F’ lytic enzyme catalytic properties, V, 384-385 chemical properties, V, 383-384 physiochemical properties, V, 382-383 purification, V, 382 Bacteriophage P1,lytic enzyme, V, 399-400

474 Bacteriophage P14, lytic enzyme V, 399400 Bacteriophage PAL,lytic enzyme, V, 4W-401 Bacteriophage SP-3, induced exonuclease, IV, 258-259 Bacteriophage T2 induced exonuclease, IV, 255 lysozyme catalytic properties, V, 381382 chemical properties, V, 380-381 physicochemical properties, V, 380 purification, V, 379 Bacteriophage T 4 induced endonucleases I1 and IV, IV, 266-269 induced exonuclease, IX, 255 lysozyme catalytic properties, V, 369-374 chemical properties, V, 366-369 enzyme assays, V, 361-364 physicochemical properties, V, 366 purification, V, 364-366 role in life cycle, V, 376379 Bacteriophage T5, induced deoxyribonuclease, IV, 261 Bacteriophage T7, induced endonuclease, IV, 266-2436 Bacteriophage TP-I, lytic enzyme of, V, 410-411 Biosynthetic pathways input signals and, I, 474-481 interaction with amphibolic pathways, I, 481482 Biotin acetyl coenzyme A carboxylase and, VI, 64-71 activation of, VIII, 18 pyruvate carboxylase and, VI, 3-5 Blood coagulation, proteolysis and, I, 416-417 Blood cells, glycogen synthetase of, IX, 354-355 Blood platelets, thrombin and, 111, 300-301 Bone, acid phosphatase, IV, 4-97 Bovine pancreatic ribonuclease, see Ribonuclease Brain aldolase, VII, 241-244

TOPICAL SUBJECT INDEX

creatine kinase of, VIII, 401-403 glycogen synthetase of, IX, 353-354 Bromelain, 111, 542545 y-Butyrobetaine hydroxylase catalytic properties, XII, 168-169 purification, XII, 167

C Calcium amylases and, V, 247-250 binding, staphylococcal nuclease and, IV, 163-171 hydrolases and, 11, 524-525 Calcium translocation and phosphoryl transfer backward reaction adenosine, phosphate and calcium binding, X, 462463 phosphoprotein formation, X, 463-465 forward reaction adenosine triphosphate-adenosine diphosphate exchange, X, 462 adenosine triphosphate and calcium binding, X, 459-460 phosphoprotein formation and, X , 460-462 Candida utilia fructose-1,6-diphosphatase inhibition by AMP, IV, 636-637 purification and properties, IV, 635-636 regulation, IV, 640 relation to SDPase, IV, 638 structure, IV, 637-638 Carbamate kinase assays, IX, 107-108 forward reaction, IX, 108-109 reverse reaction, IX, 109-110 distribution, IX, 101-102 function and metabolite control, IX, 115-119 historical background, IX, 97-100 molecular properties composition, size and subunit st’ructure, IX, 103-104 purification, IX, 102103 stability, IX, 104-105 sulfhydryl reagent effects, IX, 105

TOPICAL SUBJECT INDEX

reaction catalyzed, IX, 105-107 specificity and cofactors, IX, 107 thermodynamics, kinetics and mechanism, IX, 110-115 Carbohydrate amylase composition, V, 245-247 prothrombin composition, 111, 313 thrombin composition, 111, 286-286 Carbon-hydrogen fission electron delocalization and, 11, 287-290 other catalyses, 11, 318-320 Carbonic anhydrase active-site-directed chemical modifications anionic reagents, V, 658-660 sulfonamides, V, 661 assay methods, V, 630-632 catalytic mechanism, V, 661-662 catalytic reaction, V, 662885 substrate binding, V, 662 conformation in solution hydrogen exchange, V, 628-829 spectroscopy of native enzymes, V, 622-625 stability and denaturation, V, 626-628 titration and chemical modification, V, 625-626 crystal structure investigations, V, 608-611 active site region, V,617-622 human enzyme structure, V, 611 secondary structures, V,611416 side chain locations, V, 616617 distribution and physiological function, v, 5 w 5 9 3 historical outline, V, 588490 inhibitors anions, V, 646-052 other, V, 658 sulfonamides, V, 652-658 kinetic properties hydration of aldehydes, V, 639-640 hydrolytic reactions, V, 636-639 interconversion of CO, and HCOo-, V, 632-636 metal ion cofactor, V, 640-641 cobalt as probe of active site, V, 643-646 specificity, V, 642-643

475 thermodynamics and kinetics of zinc binding, V, 641-642 modification of, I, 139440 molecular properties composition, V, 601-603 immunological properties, V, 598-599 methods of isolation, V,593-595 physical properties, V,599-601 polymorphism and nomenclature, V, 595-598 primary structure, V, 603-607 reactions catalyzed, V, 629-630 role of metals, 11, 522-524 structure fluorescence spectroscopy, 11, 425-426 physical probes, 11, 401406 Carbon-nitrogen cleavage, general features, VII, 168-169 Carboxyl anions, acyl transfer to, 11, 2’26-235 Carboxylesterase (s) active site, organophosphorus compounds and, V, 61-64 amino acid composition, V, 52-53 assay procedures, V, 64-85 equivalent weight, V, 50 inhibitors, V, 64 kinetics, V, 60-61 molecular weight, V, 48-50 multiple molecular forms, V, 4546 pH optimum, V, 59-60 physiological and pharmacological significance, V, 65-67 preparations and criteria of purity, V, 4748 substrate specificity acyl group transfer, V, 59 aromatic amides, V, 57-59 carboxyl esters, V, 53-57 thioesters, V,57 subunit structure, V, 51-52 Carboxyl group activation of, VIII, 6-20 chemical modification, I, 174 ribonuclease, IV, 675-677 Carboxypeptidase(s) bovine, homologies, 111, 66-67 fungal, 111, 790-791 gene duplication and, I, 308-309 role of metals, 11, 519-522

476

TOPICAL SUBJECT INDEX

Carboxypeptidase A amino acid sequence, 111, 5 crystallography, 111, 17-18 description of structure, 111, 2146 determination of structure, 111, 18-21 esterase activity, 111, 7 kinetics, 111, 7-10 mechanism of action, 111, 14-17 ester cleavage, 111, 50-51 inhibition and activation, 111, 51-54 metal studies, 111, 54-56 peptide cleavage, 111, 46-50 metals and, 111, 11-12 side chain modification, 111, 1214 structure backbone conformation, 111, 29-31 correlation of sequence with, 111, 31-33

folding of chain, 111, 26-29 general features, 111, 21-23 helical segments, 111, 23-26 interpretation of substrate specificity, 111, 43-44 side chain conformation and interaction, 111, 33-36 pstructure, 111, 26 substrate binding changes, 111, 40-42 substrate and inhibitor complexes, 111, 3 7 4 0 success and failure of crystallography, 111, 44-46 substrate specificity, 111, 6-7 Carboxypeptidase B activation and inhibition, 111, 75-76 amino acid sequences and end groups, 111, 64-86

chemical composition, 111, 62-64 distribution, 111, 59-60 historical background, 111, 57-59 kinetics and competitive inhibition, 111, 71-75

mechanism, comment on, 111, 77 physical properties, 111, 61-62 purification and assay, 111,60431 specificity, 111, 69 esterase activity, 111, 71 peptidase activity, 111, 70-71 use in protein structural analysis and modification, 111, 77-79 Castor bean, lipase of, VII, 613-614

Catalase active site distal ligand identity, XIII, 376-385 ligand exchange reactions, XIII, 385-388

ligand identity at fifth and sixth coordination positions, XIII, 369376

apoprotein, selective modifications, XIII, 376-385 general properties, XIII, 366-369 historical background, XIII, 363-365 redox reactions, XIII, 388-389 nature of Compound I, XIII, 389-390 reaction mechanism, XIII, 390-408 Catalytic site, see Active site Cathepsin C, see Dipeptidyltransferase Catheptic endopeptidases cathepsin B, 111, 478-479 cathepsin D, 111, 476477 cathepsin E, 111, 477478 Catheptic exopeptidases cathepsin A, 111, 481482 cathepsin C, 111, 479-480 catheptic carboxypeptidases A and B, 111, 480 Cellobiose-2’-epimerase, properties, VI, 377

Cellulase (9) action on cellulose and related substrates, V, 287-289 applications, V, 289-290 assay and detection, V, 275-277 C1 factors and, V, 280-282 induction and repression, V, 277-278 physical and chemical properties, V, 282-285

production and isolation cultural conditions, V, 278 influence of cultural conditions on physical properties, V, 278-279 isolation methods, V, 279-280 significance and distribution, 8,274-275 substrate binding and catalytic properties, V, 285-287 Cellulose polysulfatase, V, 11-12 Cell wall structure and action of lytic enzymes chemical properties of cell walls, V, 353455

477

TOPICAL SUBJECT INDEX

mode of action of enzymes, V, 355 morphogenesis of cell wall and membrane, V, 352-353 Cerebroside sulfatase, V, 13-14 Chain extension, protein structure and,

I, 293-300 Chain shortening, protein structure and, I, 292-293 Charge-transfer forces, propinquity effects and, 11, 254-264 Chemical modification alkaline phosphatase bacterial, IV, 389-392 mammalian, IV, 427-428 amylases, V, 129-130, 261-263 chymotrypsin, 111, 234, 238-239 control of selectivity, reaction conditions and, I, 168-170 reagent choice, I, 168 determination of degrees and sites of analytical methods, I, 170-171 instability of modified residues, I, 171-172

in elucidation of noncovalent structure intermolecular reactions, I, 191-194 intramolecular reactions, I, 183-191 functional group reactivity determinants field or electrostatic effects, I, 154 hydrogen bonding, I, 152-154 matrix effects, I, 155 microenvironment polarity, I, 151-152

miscellaneous effects, I, 155-156 steric effects, I, 154-155 general principles, I, 166167 immunochemistry and, I, 199-200 inorganic pyrophosphatase, IV, 514-518 insoluble enzymes and antigens, I, 200-201

lysozyme, I, 207-211 mechanism and reactivity kinetic considerations, I, 159-162 protein functional group nucleophilicities, I, 162-164 superreactivity, I, 164-166 papain, I, 205-207 active site, 111, 515-516 other reactions, 111, 516518 pepsin, 111, 133-137

as primary structure probe chemical cleavage and, I, 181-182 location of modified residues, I, 182 proteolysis and, I, 178-181 as probe of function, accessory sites, I, 196-197 active site, I, 194-196 multifunctional enzymes, I, 197-198 proteinase inhibitors, 111, 443447 reagent reactivity determinants, catalytic factors, I, 158 electrostatic interaction, I, 157 local environment polarity, I, 158-159 selective adsorption, I, 156 steric factors, I, 157-158 reversible, I, 172-173 amino groups, I, 175 arginine, I, 174 carboxyl groups, I, 174 histidine, I, 173 methionine, I, 173 serine and threonine, I, 173 sulfhydryl groups, I, 174-175 tryptophan, I, 173 tyrosine, I, 174 ribonuclease, IV, 675-697 subtilisin BPN’, I, 203-205 subtilisins, 111, 596-602 transient, I, 176 trypsin, 111, 269-273 unsuspected, I, 176-178 X-ray crystallography and comparisons of protein solutions and crystals, I, 202-203 isomorphous heavy atom replacements, I, 201-202 Chemical reaction rate equations, derivation, 11, 61-63 Chlamydomonas reinhardti, N-acetylglutamate-5-phosphotransferase of, IX, 513 Chlorella pyrenoidosa, adenosine diphosphoryl glucose pyrophosphorylase of, VIII, 90 Chloroplast ( s ) adenosine triphosphatase Euglena gracilis, X, 394 spinach, X, 389-394 ribonucleic acid polymerase of, X, 329-330

478

TOPICAL SUBJECT INDEX

Choline dehydrogenase electron transport system and, XIII, 261-263

properties, XIII, 260-261 Choline sulfatase, V, 14 Chondroitin, hyaluronidase and, V, 310 Chondroitin sulfates, hyaluronidase and, V, 310 Chondrosulfatase, V, 12 Chymopapains, 111, 537-538 Chymotrypsin(s) active center amino acids, interaction, 111, 245-248 active-site-directed reagents and, I, 94103

individual amino acid function aspartic acid 102, 111, 235-236 aspartic acid 194, 111, 243 histidine 57, 111, 231-234 isoleucine 16, 111, 236-243 serine 195, 111, 217-231 ribonuclease and, IV, 674 structure physical probes, 11, 391-396 ultraviolet difference spectroscopy, 11, 414-417 fluorescence spectroscopy, 11, 421-424 substrate specificity acylamido interaction, 111, 208 amino acid side chain binding, 111, 205-208

locked substrates, 111, 209-212 stereo specificity, 111, 207-209 a-Chymotrypsin active center structure acyl enzyme, 111,20&202 enzyme-inhibitor complex, 111, 199200

enzyme-product complex, 111, 198199

native enzyme, 111, 190198 X-ray diffraction, 111, 190-195 7-Chymotrypsin, active center structure, 111, 202-204 8-Chymotrypsin, active center, 111,204 1.-Chymotrypsin, actiye center, 111, 204 Chymotrypsinogen activation, 111, 167-169 active site unblocking, 111, 244 X-ray crystallography, 111, 169-176 activation refolding, 111, 182-183

arginine 145, 111, 176-179 catalytic site, 111, 179-182 isoleucine 16, 111, 175-176 methionine 192, 111, 179 Chymotrypsinogen A, chemical structure, 111, 187-189 Chymotrypsinogens A and B, activation products, 111, 185-187 Circular dichroism protein structure and, 11, 381-382, 408 secondary, 11, 382386 tertiary, 11, 386-391 typical cases, 11, 381407 ribonuclease, IV, 719-723 Cis-trans isomerization about double bonds enzymic with bond migration, VI, 39@-395 without bond migration, VI, 381390 nonenzymic metals and metal ions, VI, 401406 nucleophilic catalysis, VI, 397-401 photoisomerization, VI, 395-397 reversible addition of radicals, VI, 406

Cistron, types of mutations and, I, 243245

Citramalate, cleavage and synthesis, VII, 431-432

Citrate lyase assay and isolation, VII, 378-379 catalytic properties control, VII, 387 equilibrium and kinetics, VII, 380381,386387

specificity, VII, 380, 385-386 stereospecificity, VII, 381, 387 molecular properties cofactors, VII, 379 inhibitors, VII, 380 molecular weight and subunits, VII, 379

sulfhydryl groups, VII, 379380 reaction mechanism, VII, 381382, 388389

Citrate synthase assay and isolation, VII, 358-360 catalytic properties control, VII, 362-383 equilibrium and kinetics, VII, 382

TOPICAL SUBJECT INDEX

specificity, VII, 361-362 stereospecificity, VII, 363-364 molecular properties activators, VII, 361 cofactors, VII, 360 inhibitors, VII, 361 molecular weight and subunits, VII, 360 sulfhydryl groups, VII, 360-361 proton transfer and, 11, 316-317 reaction mechanism citric anhydride as intermediary, VII, 367-368 citryl coenzyme A as intermediary, VII, 367 enolization of coenzyme A, VII, 365366 inversion of configuration of acetyl coenzyme A, VII, 366-367 keto or enol .form of oxalacetate, VII, 365 Citric acid cycle reactions stereospecificity fumarase, 11, 171-176 isocitrate dehydrogenase, 11, 168-171 succinate dehydrogenase, 11, 176-179 synthesis of citrate, 11, 164-168 Clostridium histolyticum collagenases, 111, 662-663 catalytic properties, 111, 670-689 chemical and biosynthetic modification, 111, 669-670 composition, 111, M.Wf37 culture of organism and enzyme purification, 111, 663-665 molecular size, 111, 667-668 physical properties, 111, 669 possible subunits, 111, 668-669 Clostridium pasteurianum, phosphofructokinase of, VIII, 256 Clostridium perfringem phospholipase C of, V, 84-85 polynucleotide phosphorylase of, VII, 571-572 Clostripain as collagenase contaminant, 111, 715716 definition, 111, 699-700 -aeneral features

479 pH optimum and ion efficts, 111, 706-707 sulfhydryl requirement, 111, 707-708 historical perspective, 111, 700-703 inhibitors affinity labeling, 111, 714-715 competitive, 111, 712-714 origin of active site specificity, 111, 717-71 9 purification and assay, 111, 704 specificity proteins and polypeptides, 111, 710712 synthetic esters and amides, 111, 708710 structural properties amino acid composition, 111, 706 physical constants, 111, 704-705 use in sequence analysis, 111, 716-717 Cobalt carbonic anhydrase and, V, 643646 Coenzyme A transferase (s) catalytic properties assay, IX, 487 mechanism and kinetics, IX, 488496 specificity, IX, 486-487 thermodynamics, IX, 487488 properties, IX, 485436 reactions catalyzed, IX, 483485 Coenzyme B12-dependent reactions, stereochemistry, 11, 204-214 Collagenase(s) catalytic properties activation by transition metals, 111, 683 assay, 111, 684-685, 692 cofactors, 111, 677-678 evidence for intrinsic metal component, 111, 678-683 mechanism, 111,688-689 reaction catalyzed, 111, 670, 691 specificity, 111, 670-677, 691 substrate interaction, 111, 688 thermodynamics and kinetics, 111, 685-688 zinc containing peptides, 111, 683-684 Clostridium histolytlcum, 111, 662-689 clostripain in. 111. 715-716 definition, 111, 652-653

480

TOPICAL SUBJECT INDEX

human synovial fluid, 111, 693-696 known enzymes and their functions, 111, 653-659 nature of substrate, 111,650-652 properties, generalizations, 111,659-660 R a m catesbiana, 111, 689-693 uses of, 111, 660-662 Compartmentalization, metabolic regulation and, I, 423426 Complementation, mutations and, I, 24925 1

Conformational states, frozen, I, 370-371 Control mechanisms compensatory antagonism of endproduct inhibition, I, 436437 metabolite activation, I, 434-436 precursor substrate activation, I, 438-439

Convergence, protein evolution and, I, 328-329

Cooperativity, molecular basis, I, 375379

Cooperativity index, enzyme regulation and, I, 358 Coordination schemes determination confirmatory techniques, 11, 468-470 provisional techniques, 11, 464-468 enzymes, metals and substrates, 11, 463464

experimentally determined, 11, 470476

techniques, 11, 464470 experimentally determined higher metal complexes, 11, 475476 higher substrate complexes, 11, 471475

ternary complexes, 11, 470-471 Copper-containing oxidase(s) blue biological distribution and function, XII, 558-559 history, XII, 557-558 oxidation-reduction properties, XII, 571-574

purification and molecular properties, XII, 560-563 catalytic properties reducing substrates, XII, 575-578

reduction of oxygen, XII, 578-579 specificity, inhibition and steadystate kinetics, XII, 574475 historical background, XII, 507-511 magnetic and spectroscopic properties definitions and distribution of copper forms, XII, 563-566 type 1 copper, XII, 566-567 type 2 copper, XII, 567-570 type 3 copper, XII, 570-571 reducing dioxygen to hydrogen peroxide amine oxidase, XII, 511-527 galactose oxidase, XII, 527-533 Cortisone sulfatase, V, 10 Creatine kinase active form of substrates guanidine, VIII, 414 nucleotide, VIII, 412413 adenosine polyphosphates as inhibitors, VIII, 422 anion effects, VIII, 423427 catalytic site, formation and topography, VIII, 439442 conformational changes, substrateinduced, VIII, 436-438 “essential” thiol group importance for catalytic activity, VIII, 443448 structural involvement, VIII, 44% 443

substrate effects, VIII, 448-451 equilibrium, VIII, 428-431 groups essential for activity cysteine residues, VIII, 431432 histidine, VIII, 434 lysine, VIII, 43S434 tyrosine, VIII, 434-436 historical background, VIII, 384485 hybrid, VIII, 403 mechanism, VIII, 451455 metal ions and, VIII, 409412 other brain-type, VIII, 402403 other muscle enzymes assay and specific activity, VIII, 4.p purification, VIII, 400 stability, VIII, 401 ox brain assay and specific activity, VIII, 401 purification, VIII, 401

TOPICAL SUBJECT INDEX

stability, VIII, 401-402 rabbit muscle assay and specific activity, VIII, 395-398

purification, VIII, 395 stability, VIII, 398-399 role of metals in mechanism, 11, 499500

structure amino acid composition, VIII, 390392

isoenzymes, interspecific hybrids, conformers and genetic variants, VIII, 386-390 molecular weight, VIII, 395 primary, VIII, 392-393 secondary and tertiary, VIII, 393-394 subunit shape and organization, VIII, 394-395 substrate binding, VIII, 414-420 substrate specificity guanidines and organization of creatine binding site, VIII, 403-407 nucleotides and related inhibitors, VIII, 407-409 temperature and, VIII, 420-422 Crosslinks intramolecular, ribonuclease, IV, 696697

Crolalus adamanleus venom, phospholipase AI of, V, 75-76 Crotalus atroz venom, phospholipase A2 of, V, 7.7-78 Crotonase, properties, V, 568-571 Cyanate, inorganic pyrophosphatase and, IV, 516-517 Cyanogen bromide, inorganic pyrophosphatase and, IV, 514 Cyclic adenosine monophosphate protein kinase dependent mechanism, VIII, 568-572 nomenclature, VIII, 566 properties, VIII, 572-578 purification, VIII, 568 tissue and subcellular distribution, 567-568

Cyclodiene insecticides, epoxidases and, VII, 210-211 jj’-Cystathionase, properties, VII, 51-52 y-Cystathionase, properties, VII, 57159

Cystathionine p-synthetase, properties, VII, 54-56 Cystathionine y-synthetase, properties, VII, 60-61 Cysteamine oxygenase, properties, XII, 148-149

Cysteine lyase, properties, VII, 56-57 Cysteine oxygenase, properties, XII, 149150

Cysteine residues creatine kinase, VIII, 431432, 442-451 guanidino kinases, VIII, 477-480 Cysteine synthetase, properties, VII, 54 Cystine-disulfide groups, ribonuclease, IV, 690-696 Cystine peptides, microbial proteinases, 111, 752-754 Cytidine diphosphate-D-glucose oxidoreductase, properties, V, 478479 Cytidine kinas6 see Uridine kinase Cytidine monophosphate kinase, see Uridine monophosphs te kinase Cytidine triphosphate synthetase allosteric control, X, 546-547 cooperative effects, X, 547-548 6-diazo-5-oxonorleucine and half-thesites reactivity, X, 549-550 historical, X, 540-541 purification, X, 541-542 reaction catalyzed, X, 539-540 covalent chemistry, X, 543-546 related enzymes, X, 552-558 role of nucleoside triphosphates, X, 550-552

structure, X, 542-543 Cytochrome (s), absorption spectra, XI, 398-400 bacterial common methodology in research, XI, 506-508 evolution, XI, 540-547 patterns in electron transport pathways, XI, 508-509 sequence and structure, XI, 534-540 type b, XI, 591-593 Cytochrome b, mammalian, purification of, XI, 563564

occurrence and function, XI, 550-551 plant, XI, 587-591

TOPICAL SUBJECT INDEX

reaction with substrates adenosine triphosphate-induced reduction, XI, 561 antimycin effect, XI, 562 kinetics, XI, 560-561 redox change, XI, 562-563 respiratory, XI, 551652 absorption spectra, XI, 554-558 different types, XI, 552-554 miscellaneous, XI, 563-564 oxidation-reduction potential, XI, 558-560 Cytochrome bl, occurrence of, XI, 579584 Cytochrome bl, preparation and properties, XI, 585-587 Cytochrome ba biological role, XI, 567 distribution, XI, 566 isolation, XI, 567-568 nomenclature, XI, 565-566 properties, chemical, XI, 668-569 physical, XI, 569 spectral, XI, 589-571 structure, XI, 571-572 amino acid sequence, XI, 572-573 ternary, XI, 573-576 Cytochrome bs reductase cytochrome P-450 reductase and, XIII, 151-153 mechanism, microsome bound, XIII, 161-162 mechanism of Strittmatter, review, XIII, 156-161 methemoglobin reductase and, XIII, 164-165 molecular properties, amphipathic and soluble forms, XIII, 154-156 structural studies, XIII, 162-164 Cytochrome b-662, distribution and preparation, XI, 584 properties and structure, XI, 584-585 Cytochrome(s) c, bacterial, XI, 497-506 evolution, XI, 64&547 function, XI, 506-509 photosynthesis, XI, 509316 respiratory chain, XI, 518634 structure, and sequence, XI, 534-540

eukaryotic, photosynthetic cytochromes f and CJSS,XI, 493497 respiratory cytochrome c, XI, 400489 respiratory cytochrome c,, XI, 489492 metal complexes and, 11, 534-538 principles of protein evolution and, I, 274-285 respiratory amino acid sequence, XI, 419429 evolution, XI, 429-450 molecular folding and structural integrity, XI, 450463 oxidation reduction mechanism, XI, 463489 structure, XI, 405-419 Cytochrome cl, respiratory, XI, 489-492 Cytochrome cm, photosynthetic, XI, 493497 Cytochrome c oxidase biological role, XIII, 299-300 chemical and physical properties, XIII, 301-302, 313-314 interaction with cytochrome c, XIII, 334-335 kinetic studies, XIII, 335-337 models, XIII, 314-315 chemistry of oxygen reduction, XIII, 302-307 electronic spectroscopy absorption spectra, XIII, 315-319 circular dichroic spectra, XIII, 319 electron paramagnetic resonance studies copper, XIII, 329-330 iron, XIII, 331 ligand binding effects, XIII, 332333 p-oxobishemin and, XIII, 333334 valence state changes and, XIII, 331-332 historical background, XIII, 300-301 ligand binding studies, XIII, 319-320 azide, fluoride and cyanide, XIII, 320-321 carbon monoxide, XIII, 321-323 dioxygen, XIII, 323-326 lipids of, XIII, 312-313 mechanisms, XIII, 337344

TOPICAL SUBJECT INDEX

metal components, XIII, 307-309 potent iome try electron economy, XIII, 325-326 iron-copper coupling, XIII, 326-327 ligand binding effects, XIII, 327-328 summary, XIII, 328329 protein of, XIII, 309-312 Cytochrome c peroxidase cytochrome c interaction, XIII, 356360 enzymic activity, XIII, 352-353 general comments, 360-361 historical background, XIII, 345-347 preparation and molecular properties, XIII, 347-348 reaction mechanism, XIII, 353356 structural aspects, XIII, 348-351 Cytochrome f, photosynthetic, XI, 493497 Cytochrome 0,occurrence and properties, XI, 592-593 Cytochrome P-450 reductase, XIII, 165166 catalytic activities, XIII, 167-169 general properties, XIII, 166-167 mechanism, XIII, 169-173 Cytophaga, isoamylase of, V, 204-206

D Deaminases, modification of, I, 127-128 Debranching enzyme (s) characterization of, V, 223-226 classes of, V, 192-194 direct, V, 194-208 indirect, V, 208-210 assay of glucosidase-transferase activity, V, 211-212 effect on glycogen structure, V, 217219 glucosidase-transferase in glycogen storage disease, V, 221-222 other enzymes, V, 222-223 pH dependence, V, 215-217 purification and physical properties, v, 210-211 reversion reactions, V, 219-220 specificity, V, 213-215 in vivo roles, V, 226-228 structure determination and, 228-229

arrangement of unit chains, V, 230233 average chain length, V, 229-230 enzymic action pattern, V, 233-234 Debye forces, propinquity effects and, 11, 254-264 3-Decynoyl-N-acetylcysteamine,isomerization of, V, 459-461 Dehydratases, miscellaneous, VII, 53-54 Dehydration ( 8 ) metal ion-assisted L-arabonate-n-fuconate, v, 581-582 n-arabonate and p-xylonate, V, 582 galactonate, V, 578479 gluconate, V, 575-578 hexarate, V, 579-581 n-mannonate and n-altronate, V, 579 Schiff base-assisted glucosaminate dehydrase, V, 586 2-keto-3-deoxy-~-arabonatedehydratase, V, 583-585 5-keto-4deoxyglucarate dehydratase, v, 585-586 Dehydrogenases the bigger family general XI, 94 members, XI, 94-99 primordial mononucleotide binding proteins, XI, 101 structural relationships, XI, 99 time scale, XI, 99-101 characteristics of, XIII, 90-91 comparison of three-dimensional structure known structures, XI, 64-65 malate and lactate to alcohol and glyceraldehyde-3-phosphatedehydrogenase, XI, 65-69 mononucleotide binding unit, XI, 69-70 NAD binding structure, XI, 70-73 recognition of similar structural domains, XI, 73-74 dissociation constants of enzyme-coenzyme compounds, cooperative effects, XI, 46-47 initial measurements and binding studies, XI, 38-44 pH effects and role of histidine, XI, 44-46

484 domain and subunit assembly, conservation of contacts, XI, 91 gene fusion, XI, 89-90 quaternary structure, XI, 91-93 quaternary structure-evolution, XI, 93 equilibrium and kinetics of enzymecoenzyme reactions dissociation constants, XI, 34-42 kinetics, XI, 4247 flavodoxin and, XI, 94-96 functional aspects of dinucleotide binding domains, XI, 83-84 adenosine monophosphate, XI, 87-88 nicotinamide adenine dinucleotide, XI, 84-87 nicotinamide mononucleotide, XI, 88-89 some generalizations, XI, 89 inhibition and activation analogs, XI, 30-34 product, XI, 34-35 substrate, XI, 25-30 initial rate equations ordered mechanism with isomeric enzyme-coenzyme compounds : conformation change, XI, 10-11 ordered and random mechanisms for three-substrate reactions, XI, 13-15 preferred pathway mechanism, XI, 12-13 rapid equilibrium mechanism, XI, 11-12 simple ordered mechanism, XI, 7-10 kinases and, XI, 96-98 kinetics of enzyme-coenzyme reactions, conformational changes, XI, 50-52 velocity constants, XI, 47-50 kinetics of transient phase, XI, 47-48 integrated rate equations, XI, 48-50 lactate dehydrogenases, XI, 52-53 liver alcohol dehydrogenase, XI, 5052 other enzymes, XI, 53-55 kinetic studies with alternative substrates liver alcohol dehydrogenase, XI, 2022 other enzymes, XI, 23-24

TOPICAL SUBJECT INDEX

yeast alcohol dehydrogenase, XI, 2223 nicotinamide adenine dinucleotidelinked, metal complexes and, 11, 525-528 preliminary generalizations, XI, 3-4 quaternary structure, XI, 91-92 alcohol dehydrogenase subunit association, XI, 93 evolution, XI, 93 P and R axis-cooperativity, XI, 9192 Q axis, XI, 91 rhodanese and, XI, 98 sequence comparisons based on structural alignments glutamate dehydrogenase comparisons, XI, 79 lactate, alcohol and glyceraldehyde3-phosphate dehydrogenases, XI, 77-79 statistics of comparisons, XI, 79-83 sequence comparisons in the absence of three-dimensional structural information significance, XI, 7677 suggested homologies or analogies, XI, 74-76 steady-state kinetics cooperative rate effects, XI, 31-34 inhibition and activation by substrates, substrate analogs and products, XI, 22-31 initial rate equations for ordered and random mechanisms, XI, 6-14 isotope exchange a t equilibrium, XI, 14-16 kinetic studies with alternate substrates, XI, 18-22 maximum rate and Haldane relations, XI, 16-18 phenomenological initial rate equations, XI, 4-6 subtilism and, XI, 98-99 Dehydroluciferin, adenylylation of, VIII, 19-20 Deletions, protein structure and, I, 293300 Deoxyadenosine kinase, properties, IX, 66-68

485

TOPICAL SUBJECT INDEX

Deoxyadenosine monophosphate kinase, properties, IX, 86-87, 95-96 3-Deoxy-~-arabino-heptulosonate7-phosphate synthase, modification of, I ,

assay, X, 239-240 isolation of covalent intermediates deoxyribonucleic acid-adenylate,

139

ligase-adenylate, X, 245 mechanism of phosphodiester bond synthesis, X, 244-252 physical homogeneity, X, 241 physical properties, amino acid analysis, X, 242 molecular weight, X, 241 stoichiometry, X, 242-243 purification, X, 240-241 reversal of, 246-248 role in vivo bacteriophage-induced, X, 252-254 Escherichia coli, X, 254-259 steady state kinetics overall reaction, X, 248-249 partial reactions, X, 249-252 Deoxyribonucleic acid methyltransferases biological significance, IX, 194-195 occurrence, IX, 190 properties bacterial, IX, 190-192 eukaryotic, IX, 192-193 regulation, IX, 193-194 Deoxyribonucleic acid polymerase(s) catalytic reactions basic features, X, 123-124 fidelity of replication, X, 127-129 implications of mechanism, X, 134-

Deoxycytidine kinase, properties, IX, 62-66

Deoxycytidine monophosphate kinase, properties, IX, 88-89, 95-96 Deoxycytidylate hydroxymethyltransferase, properties, IX, 209-210 Deoxyguanosine kinase, properties, IX, 68-69

Deoxyguanosine monophosphate kinase, IX, 94-96, see Guanosine monophosphate kinase Deoxyribonuclease (s) adenosine triphosphate-dependent, IV, 259, 261-262

catalytic properties inhibitors, IV, 281-283 phosphodiesterase activity, IV, 283 substrate concentration, pH and ions, IV, 280-281 classification of, IV, 251-252 spleen components, IV, 275 dimeric structure, IV, 275-276 distribution, localization and role, IV, 285-287 features of degradation, IV, 276-278 general catalytic properties, IV, 280283

135

multiple sites in active center, X,

isolation, IV, 272-273 mechanism, IV, 278-280 methods of investigation, IV, 278 physical and chemical properties, IV,

124-125

polymerization step, X, 125-126 pyrophosphorolysis and pyrophosphate exchange, X, 126-127 ribonucleotides and, X, 133-134 specialized functions, X, 135-137 synthesis without template, X, 132-

273-275

specificity, IV, 283-285 Deoxyribonuclease I active center, IV, 297-299 chemical nature, IV, 292-297 historical background, IV, 289-291 inhibitor, IV, 299-302 ions and, IV, 302-303 kinetics, IV, 303-308 physiological role, I, 310 specificity, IV, 308-310 Deoxyribonucleic acid ligase(s) adenylyl transfer functions, VIII, 45-48

X,

245-246

133

synthetic product, X, 131-132 template-primer, X, 129131 classification native and denatured templates, X, 182-183

6

polyribonucleotide templates, X, 184 size, X, 183 comparison of properties, X, 144

486

TOPICAL SUBJECT INDEX

definitions and measurements, X, 175176

exonucleases associated, IV, 255-258 fidelity of synthesis, X, 201-202 historical, X, 174-175 inhibitors and activators, X, 199-201 initiators, X, 203-204 intracellular distribution chloroplasts, X, 181 membranes and other structures, X, 181-182

mitochondria, X, 180-181 nuclei, X, 179-180 isolation and physicochemical properties, X, 120-123 kinetics extent of synthesis, X, 195-196 temperature and, X, 194-195 metal activators, X, 193-194 molecular properties antisera and, X, 192-193 homogeneity, X, 188-189 presence of nuclease, X, 189-190 sulfhydryl groups, X, 190-191 zinc, X, 191-192 occurrence, X, 176-179 pH and PI, X, 194 properties of purified viral enzyme, X, 218

size, X, 219-220 storage and stability, X, 219 proteolytic cleavage: two enzymes in one polypeptide, X, 138-139 purification chromatography, X, 185-186 extent, X, 186-188 stability, X, 185 subcellular fractionation, X, 184-185 substrates requirements, X, 198-199 specificity, X, 196-197 templates deoxyribonucleic acid, X, 204-205 ribonucleic acid, X, 206-207 variety of, X, 119-120 viral, purification, X, 216-218 solubilization, X, 215 virus purification and, X, 214-215

2-Deoxyribose-5-phosphate aldolase catalytic reaction activators and inhibitors, VII, 320 assay, VII, 319-320 equilibrium constant, VII, 320 pH optimum, VII, 320 Schiff base formation, VII, 321 substrate specificity, VII, 321 historical background, VII, 315 metabolic significance, VII, 316-317 molecular properties isolation, VII, 317-319 physical properties, VII, 319 occurrence, VII, 315-316 Deoxysugars synthesis, general considerations, V, 465467

Deoxy sugar aldolase (s) ,general, VII, 303-304

Deoxythymidine diphosphate-D-glucose oxido-reductase kinetic properties, V, 469-470 molecular properties, V, 467469 pyridine nucleotide and substrate release, V, 474476 subunit association, V, 476-478 reaction mechanism enzyme bound pyridine nucleotide,

V, 472-473 intramolecular hydrogen transfer, V, 470-472

isotope effects, V, 473-474 Deoxythymidine kinase distribution, purification and assay,

IX, 69-70 kinetic, molecular and allosteric properties, IX, 71-74 reaction mechanism; active site, IX, 74

substrate specificity, IX, 70 Deoxythymidine monophosphate kinase distribution and purification ; assay; stability, IX, 91-92 kinetic and molecular properties, IX, 93-96

substrate specificity, IX, 92-93 Deoxyuridylate hydroxymethyl transferase, properties, IX, 210 Dermatan sulfate, hyaluronidase and, V, 310-311

487

TOPICAL SUBJECT INDEX

2,5-Diaminohexanoate, lysine mutase and, VI, 554 Diazonium-1H-tetrazole, inorganic pyrophosphatase and, IV, 517-518 3,&Dideoxyhexoses, synthesis of, V, 479480 7,&Dihydro-2-amino-4-hydroxy-6-hydroxymethylpteridine pyrophosphokinase, X, 627-628 Diisopropylfluorophosphate, papain and, 111, 516-517 Dioxygenase(s) biological function and general properties double bond cleavage, XII, 123-125 double hydroxylation, XII, 125 miscellaneous, XII, 125-127 sulfur-containing compounds, XII, 125 classification, XII, 121-123 heme-containing indoleamine 2,3-dioxygenase, XII, 130-132 tryptophan dioxygenase, XII, 127130 history and definition, XI, 120-121 a-ketoglutarate, XII, 151-152 y-butyrobetaine hydroxylase, XII, 167-169 p-hydroxyphenylpyruvate hydroxylase, XII, 179-183 lysyl hydroxylase, XII, 165-167 mechanism, XII, 183-189 prolyl hydroxylase, XII, 152-165 pyrimidines and nucleosides, XII, 169-179 nonheme iron-containing, XII, 132-133 cysteamine oxygenase, XII, 148-149 cysteine oxygenase, XII, 149-150 phenolic, XII, 133-148 phenolic extradiol, XII, 140-144 intradiol, XII, 133-140 others, XII, 144-148 Dipeptidases, 111, 116-117 Dipeptidyl aminopeptidase 1, see Dipeptidy 1-transferase Dipeptidyltransferase, 111, 105-111, see also Cathepsin C

Diphosphoglycerate mutase, properties, VI, 476477 Disaccharide phosphorylases general background, VII, 515-518 substrate specificity, VII, 526-528 Disulfide bridges elastase, 111, 339341, 348-349 p-hydroxydecanoyl thioester dehydrase, V, 452453 prothrombin, 111, 313 thrombin, 111, 290-291 trypsin, 111, 271 Disulfide groups, amylases, V, 244-245 Disulfide loop, proteinase inhibitor reactive site and, 111,420-422 Divergence protein evolution, I, 314-316 factors influencing rate, I, 317-321 speciation of homologous proteins: genetic drift, I, 321-328 Dopamine p-monooxygenase, properties of, XII, 294-295 Double bonds isolated, sterospecificity of reactions, 111, 179-186 Double reciprocal plots enzyme regulation and, I, 356-357 nonlinear, 11, 56-59 Drosophila melanogaster acid phosphatase of, IV, 498 molybdenum hydroxylase genetics of, XII, 406412 E

Ehrlich ascites cells, 5'-nucleotidase of, IV, 34-49 Elastase, see also Tosyl elastase activation, 111, 244-245 amino acid sequence determination complete sequence, 111,341-343 disulfide bridged peptides, 111,339341 assay methods elastin and, 111, 325-326 synthetic substrates and, 111, 326327 criteria of purity, 111, 32-29 crystals, activity of, 365-366 enzymic activity

488

TOPICAL SUBJECT INDEX

inhibitors and activators, 111,337-338 irreversible inhibitors, 111, 338-339 proteins and, 111, 332-333 synthetic substrates and, 111, 333337

history and distribution, 111, 323-325 physicochemical properties, 111, 329331

proelastase and, 111, 331-332 purification methods, 111, 327-328 reactivity and pK. of amino groups,

111, 367472 competitive labeling technique, 111, 366-367

nitrous acid and, 111, 372 valine and aspartate residues, 111, 372-373

ribonuclease and, IV, 672-673 sequence homologies in serine proteinases activation peptides, 111, 343-344 B chains, 111, 344-348 disulfide bridges, 111, 348-349 hypothetical models, 111, 349-352 stability, 111, 332 X-ray crystallography, crystals and, 111, 353-354 Fourier synthesis, 111, 355-356 heavy atom derivatives, 111, 354-355 Electron delocaliration, enzymic-C-H fission and, 11, 287-290 Electron density maps, interpretation of,

I, 46-52 Electron microscopy, collagenase action and, 111, 693 Electron paramagnetic reaonance, ribonuclease, IV, 723-725 Electron-transferring flavoprotein catalytic properties, XII, 116-118 function, XII, 109-110 molecular properties oxidation-reduction, XII, 116 properties of chromophore, XII, 111-115

purification, molecular weight and amino acid composition, XII, 111 a, ,!) Elimination reactions aconitase, 11, 302-304 factors influencing, VII, 79-81 conjugation, VII, 81

delocaliration of ,!) charge, VII, 81-82

modification of X , VII, 82-84 solvent effects, VII, 85-86 stereoelectronic control, VII, 85 weak bases, VII, 86 fumarase, 11, 304-308 nonenrymic aspartate-fumarate interconversion, VII, 86-88 stereochemistry, 11, 309-312 ,!)-Elimination reactions mechanism, VII, 66-72 dehydratases, VII, 39-48 y-Elimination and replacement mechanism, VII, 72-73 pyridoxal phosphate and, VII, 59-62 y-Elimination reactions, pyridoxallinked, VII, 57-59 Elongation stringent response and, X, 78-79,82-83 effect of ppGpp, X, 81-82 synthesis of MSI and MSII, X, 79-81 Elongation factor(s) bacterial, X, 55-57 function of factor G, X, 64-67 function of factor Tu, X, 58-63 physical properties, X, 57-58 role of factor Ts, X, 63-64 Elongation factor G interaction with ribosomes, X, 66-67 protein synthesis and, X, 65-66 Elongation factor Tu complex interaction with ribosomes,

X, 62-63 guanine nucleotide binding, X, 58-59 ternqry complex, X, 59-62 Endolysin bacteriophage A catalytic properties, V, 391-392 chemical properties, V, 388391 physicochemical properties, V, 387388

purification, V, 385-386 Endonuclease (8) bacterial nonspecific, IV, 259-262 specific, IV, 262-270 Enolase active site components, V, 532-534

TOPICAL SUBJECT INDEX

number, V, 530-532 chemical properties amino acid composition, V, 503 end groups and terminal sequences, V, 503-506 immunochemistry, V, 507 polypeptide chain identity, V, 506507 criteria of purity, V, 502-503 general conisderations, V,499-501 kinetic parameters, V,523-524 magnesium and, V,524-526 mechanism of dehydration reaction evidence for carbanion intermediate, v, 537 isotope effects, V, 535-536 molecular properties, summary, V, 518-519 monomer-dirner activity relationships, v, 537-538 physical properties, V,507-508 electrophoretic mobility, V, 508-510 subunit structure, V, 510-518 properties of reaction catalyzed assay, V, 519-523 equilibrium, V, 523 substrates, V, 519 rabbit muscle, glycidol phosphate and, v, 534 role of metals, 11, 508-509 substrate specificity, active site mapping with analogs, V, 526-529 yeast carboxymethylation, V, 533 photooxidation, V, 533-534 Enterochrome-566, properties, XI, 592 Enzyme (s) bridge complexes, 11, 477478 carbonyl containing other, 11, 356358 pyridoxal phosphate, 11, 346-356 covalent modification glycogen phosphorylase, I, 413-415 glutamine synthetase, I, 409413 development of novel properties, I, 334-335 intrachain repetitions, I, 335 multichain enzymes, I, 335-337 , inactive, proteinase inhibitors and, 111, 454-457

loss of function, survival value, I, 332-334 metal linkage coordination geometry, 11, 492494 nature of ligands, 11, 490-492 role of metals in mechanism, 11,498499 hydrolases, 11, 519-525 lyases, 11, 508-519 phosphoenolpyruvate carboxylation, 11, 507-508 phosphoryl and nucleotidyl transfer, 11, 499-507 specificity, historical background, 11, 119-129 structure in solution circular dichroism and optical rotatory dichroism, 11, 381-408 fluorescence spectroscopy, 11, 418430 geometry and quaternary structure, 11, 440-442 infrared spectroscopy, 11, 374-379 ionizable groups, 11, 430440 secondary and tertiary structures, 11, 373-374 ultraviolet absorption of peptide groups, 11, 379-380 ultraviolet difference spectroscopy, 11, 408-417 regulation of concentration balance between synthesis and degradation, I, 402-403 catabolite repression, I, 400401 feedback repression of synthesis, I, 401402 substrate induction of synthesis, I, 399-400 Enzyme crystallography, state of the art, I, 86-87 Enzyme crystals differences from other crystals, I, 5-7 Fourier description structure analysis and phase problem, I, 29-32 as sum of waves, I, 26-29 growth of, batch, I, 19 equilibrium dialysis, I, 19-22 vapor diffusion, I, 19

TOPICAL SUBJECT INDEX

mounting and radiation damage, I, 22-23 structure and symmetry, I, 7-13 Enzyme regulation diagnostic tests, I, 356358 allosteric protein evaluation, I , 372375 cooperativity index, I, 358 double reciprocal plots, I , 356-357 equations of state, I, 386388 fitting saturation curves, I, 361365 frozen conformational states, I, 370-371 Hill plot, I, 358-359 minimal substrate technique, I, 372 Scatchard or Klotz plots, I, 359-361 Leelocity curves, I, 368369 Y X or N x versus log (X) plots, I, 357 gloasary, I, 395-396 molecular models, qualitative features, I, 344-348 quantitative molecular parameters derivation of general equation, I, 34a353 simple models, I, 353-355 Epimerase(s) definition and history, VI, 365-357 keto-enol rearrangement and, VI, 373-374 noncarbohydrate, VI, 378 oxidation-reduction and, VI, 369-371 proton shifts and, 11, 295-298 Epinephrine, glycogen synthetase and, IX, 338, 341 Epoxidase(s) general considerations, VII, 199-200 metabolic roles, VII, 200-201 Equations of state, enzyme regulation and, I, 365-368 Equilibria chemical, metabolic regulation and, I, 418-419 Erthrocytes, phosphofructokinase of, 257 Eecherichia coli acetyl coenzyme A carboxylase subunits, VI, 60-64 N-acetylglutamate-5-phosphotransferase of, IX, 513 acid phosphatase of, IV, 498

adenosine diphosphorylglucose pyrophosphorylase energy charge and, VIII, 104-107 manganese effect, VII, 102-104 mutants, VIII, 109-117 alkaline phosphatase, I, 251-254, IV, 373-415 asparaginase, IV, 107-116 aspartate metabolism, regulation of, I , 457-459 aspartate transcarbamylase of, IX, 225497 aspartokinases of, VIII, 513-544 adenylylation of, VIII, 44-45 cytochrome b, of, XI, 579, 580, 581 cytochrome b-562, properties and structure, X I , 584-585 deoxyribonuclease, ATP-dependent, IV, 259 deoxyribonucleic acid ligase isolation and physical properties, X, 239-243 mechanism, X , 244-252 role in vivo, X, 254-259 deoxyribonucleic acid polymerase I, physiological role, X, 139-141 deoxyribonucleic acid polymerase 11, physiological role, X, 142 deoxyribonucleic acid polymerase, 111, physiological role, X, 142-143 deoxyribonucleic acid polymerase, exonucleases and, IV, 255-258 endonuclease I, IV, 259-280 endonuclease 11, IV, 204-285 exonucleases I and 111, IV, 253 exonuclease I V of, IV, 254-255 3' + 5' exonuclease, IV, 256 5' + 3' exonuclease, IV, 256-258 fatty acid synthetase, 3decynoyl-Nacetyl-cysteamine and, V, 461-463 fructose-l,6-diphosphatase of, IV, 63% 639 P-galactosidase, VII, 624-625 8-galastosidase, mutations, I, 255-266 glutaminase, IV, 80-93 glutamine synthetase of, VIII, 40-44, X, 755-807 inorganic pyrophosphatase catalytic properties, IV, 518-526 molecular properties, IV, 501418

491

TOPICAL SUBJECT INDEX

membrane adenosine triphosphatase, X , 416-421 methyltransferase of, IX, 154-160 phage lytic enzymes, V, 355-361 F series polysaccharide depolymerase, V, 392-393 X endolysin, V, 385-392 N20F’ lytic enzyme, V, 382-385 T 2 lysozyme, V, 379-382 T4 lysozyme, V, 361-379 phosphofructokinase of, VIII, 256-257 polynucleotide phosphorylase of, VII, 548-570 mutant, VII, 571-572 restriction endonucleases, IV, 263-264 ribonucleases of, IV, 241-243 succinyl coenzyme A synthetase of, X, 582591 thioredoxin reductase, general properties, XIII, 144-145 tryptophan synthetase catalytic properties, VII, 22-30 molecular properties, VII, 8-21 vitamin BIZmethyltransferase of, IX, 122-154 Ester(s), subtilisin and, 111, 592-593 Esterase(s) classification and distribution, V, 43-45 modification of, I, 124-127 other, V, 67-89 Estrone sulfatase, V, 6-7 Ethanolamine deaminase activation by monovalent cations, VI, 545 assays, VI, 542 cobamide binding sites, VI, 543-544 inhibitors, cobamide, VI, 544-545 isolation, VI, 541-542 occurrence, VI, 540-541 other properties, VI, 546-547 physical properties, VI, 542-543 specificity of coenzyme requirement, VI, 544 substrate specificity and binding, VI, 545-546 a-Ethylmalate, synthesis, VII, 426 Etiocholanolone sulfatase, V, 9-10 Euglena gracilis, chloroplast adenosine triphosphatase, X, 394

Eukaryotes deoxyribonucleic acid polymerases of, X , 173-209 polypeptide chain initiation inhibitors, X, 43 initiation factors, X, 2943 initiator aminoacyl-transfer ribonucleic acid and, X, 28-29 messenger ribonucleic acid translation, X,43-44 ribonucleic acid polymerases, X,261331 Evolution allosteric proteins, I, 390393 principles, cytochrome c and, I, 274285 structure-function relationships in proteins, I, 267-274 Evolutionary factors expression in protein structure convergence, I, 328-329 development of novel properties, I, 334-337 divergence, I, 314-328 loss of function and survival value, I, 332-334 parallelism, I, 329-332 Exonuclease(s), bacterial, IV, 252-259 F Fast reaction techniques application of conformational changes, 11, 112-114 enzyme-substrate reactions, 11, 108112 Fatty acid(s) activation, VIII, 6-11 biosynthesis, acyl carrier protein and, VIII, 164-165 unsaturated, cis-trans isomerization, VI, 390-394 Fatty acid: coenzyme A ligases catalytic properties, general considerations, X, 475-477 Fatty acid synthetase(s), I, 226-228 3-decynoyl-N-acetylcysteamine and, V, 461-463 Fatty acyl coenzyme A synthetases distribution and isolation

TOPICAL SUBJECT INDEX

acetate : coenzyme A ligase, X, 470 long chain fatty acid: coenzyme A ligase, X, 471-472 medium chain fatty acid :I coenzyme A ligase, X, 470-471 medium-long chain fatty acid: coenzyme A ligase, x, 472-473 other related acid: coenzyme A ligases, X, 473-474 scope of chapter, X, 469-470 Feedback inhibition metabolic regulation and, I, 403404 allosteric concept, I, 404406 cooperative effect, I, 406408 terminology, I, 408 Feedback regulation multifunctional pathways, I, 444445 aspartate metabolism, I, 457-459 concerted inhibition, I, 449-450 cumulative inhibition, I, 452454 enzyme multiplicity, I, 445-447 heterogeneous metabolic pool inhibition, I, 454-455 multivalent repression, I, 455-457 sequential controls, I, 447-449 synergistic inhibition, I, 450452 Fermentation, reductive monocarboxylic acid cycle and, VI, 213-214 Ferredoxin( s) bacterial background, XII, 37-39 chemical properties, XII, 4 4 4 6 physical properties, XII, 39-44 chemical properties apoproteins and reconstitution, XII, 25-28

chemical modification, XII, 28-29 electron transfer reactions, XII, 29 exopeptidases and, XII, 28-31 two irons per center background, XII, 15-20 chemical properties, XII, 25-31 perturbants of EPR spectra, XII, 23-25

physical properties, XII, 20-23 Ferredoxin-linked carboxylations general considerations, VI, 193-196, 214-216

a-ketobutyrate synthase, VI, 203-204 a-ketoglutarate synthase, VI, 201-203

a-ketoisovalerate synthase, VI, 205 phenylpyruvate synthase, VI, 205-207 pyruvate synthase, VI, 197-201 reductive carboxylic acid cycle of bacterial photosynthesis, VI, 207213

reductive monocarboxylic acid cycle of fermentative metabolism, VI, 213-214

Fibrinogen derivatives, thrombin and, 111,298-299 thrombin and, 111, 295-298 Ficin, 111, 538-542 Fish, glycogen synthetase of, IX, 358 Flavin coenzyme(s), structure and chemistry, XII, 423-425 Flavodoxins composition, molecular weight and purification, XII, 59-60 discovery, nomenclature and distribution, XII, 58-59 flavin mononucleotide in, XII, 65-66 flavin-protein interactions apoprotein preparation and properties, XII, 82-83 modified flavins and, XII, 85-87 protein modification and, XII, 87-88 thermodynamics and kinetics, XII, 83-85

function, XII, 60-63 oxidation-reduction potentials, XII, 98-102

reactivity comproportionation, XII, 102-103 dithionite, XII, 103-104 oxygen and ferricyanide, XII, 105108

redox proteins and, XII, 108-109 regulation by iron, XII, 63-65 spectroscopic properties circular dichroism, XII, 94-95 fluorescence, XII, 93-94 magnetic resonance, XII, 95-98 optical absorption spectra, XII, 88-91, 99

single crystal absorbance, XII, 91-93 structure, XII, 66-67 determination and comparison of chemical sequences, XII, 67-70 three-dimensional, XII, 70-82

TOPICAL SUBJECT INDEX

Flavoprotein monooxygenase(s) external, XII, 204-206 bacterial luciferase, XII, 226-229 m-hydroxybenzoate-4-hydroxylase, XII, 225 m-hydroxybenzoate-6-hydroxylase, XII, 224-225 p-hydroxybenzoate hydroxylase, XII, 211-216 imidazoleacetate monooxygenase, XII, 225-226 kynurenine-3-hydroxylase,XII, 230-231 melilotate hydroxylase, XII, 21722I microsomal amine oxidase, XII, 229-230 orcinol hydroxylase, XII, 223-224 phenol hydroxylase, XII, 221-223 salicylate hydroxylase, XII, 206-211 internal, XII, 193-194 arginine, XII, 203-204 lactate, XII, 194-199 lysine, XII, 199-203 Flavoprotein oxidase ( s ) chemical mechanism flavin reduction, XII, 474-503 oxidation by oxygen, XII, 503-505 definition of, XII, 421-423 kinetics computer simulation, XII, 442-443 strategy, XII, 425-426 summary, XII, 443-445 mechanism confirmation, XII, 437442 correlation of steady-state and transient kinetics, XII, 435-437 molecular properties and kinetic mechanism n-amino acid oxidase, XII, 445-466 L-amino acid oxidase, XII, 456461 glucose oxidase, XII, 461-466 monoamine oxidase, XII, 466-471 old yellow enzyme, XII, 471-473 steady-state kinetics rate equation, XII, 429-432 velocity measurements, XII, 426429 transient-state kinetics oxidative half-reaction, XII, 435

493 reductive half-reaction, XII, 432431 Fluorescence, ribonuclease, IV, 718-719 Fluorescence spectroscopy protein structure, 11, 418421,429-430 typical cases, 11, 421-429 Fluorocitrate, aconitase and, V, 428-430 Formaldehyde (and congeners) transfer deoxycytidylate hydroxymethyltransferase, IX, 209-210 deoxyuridylate hydroxymethyltransferase, IX, 210 glycine decarboxylase, IX, 221-223 thymidylate synthetase, IX, 210-215 serine hydroxymethyltransferase, IX, 215-221 Formate (and congeners) transfer 5-amino-4-imidazole carboxamide ribonucleotide transformylase, IX, 204-205 formiminoglutamate formiminotransferase, IX, 206-207 formiminoglycine formiminotransferase, IX, 206 5-formiminotetrahydrofolate cyclodeaminase, IX, 202-203 N-formylglutamate transformylase, IX, 207-208 5-formyltetrahydrofolate cyclodehydrase, IX, 207-208 10-formyltetrahydrofolate deacylase, IX, 200 10-formyltetrahydrofolate synthetase, IX, 198-200 glycinamide ribonucleotide transformylase, IX, 203-204 5,lO-methenyltetrahydrofolate cyclohydrolase, IX, 201 methionyltransfer ribonucleic acid transformylase, IX, 208-209 Formiminoglutamate formiminotransferase, properties, IX, 206-207 Formiminoglycine formiminotransferase, properties, IX, 206 5-Formiminotetrahydrofolate cyclodeaminase, properties, IX, 202-203 N-Formylglutamate transformylase, properties, IX, 207-208

494 5-Formyltetrahydrofolate cyclodehydrase, properties, IX, 201-202 10-Formyltetrahydrofolate deacylase, properties, IX, 200 10-Formyltetrahydrofolate synthetase, properties, IX, 198-200 Frog muscle, glycogen synthetase of, IX, 357 Fructose-1,Bdiphosphatase(8) activation by disulfide exchange coenzyme A and acyl carrier protein, IV, 623-624 cystamine and, IV, 622-623 homocystine, IV, 624-625 as regulatory mechanism, IV, 625626 assay methods and mechanism of action, IV, 615-616 Candida utilis inhibition by AMP, IV, 636-637 purification and properties, IV, 635836 regulation, IV, 640 relation to SDPase, IV, 638 structure, IV, 637438 comparative properties, IV, 645646 Eschen'chia coli, IV, 638439 higher plants and blue-green algae physiological role, IV, 642-643 purification and properties, IV, 640642 regulation, IV, 643 historical review, IV, 612413 kidney purification and properties, IV, 629630 regulation, IV, 630431 molecular structure binding sites for divalent cation, IV, 628-629 binding sites for F D P and AMP, IV, 627-628 induced conformational changes, IV, 629 molecular weight and subunit structure, IV, 826-627 muscle evidence for presence. IV. 632 physiological ;ole, IV; 634-835

TOPICAL SUBJECT INDEX

purification and properties, IV, 632633 structure and relation to other enzymes, IV, 633-634 physiological role, IV, 613-615, 644645 proteolysis, changes induced, IV, 618 purification and properties, IV, 629630, 632-633 optimum pH and cation effects, IV, 617418 purification procedures, IV, 616-617 substrate specificity, IV, 618 regulation, I, 439-441,IV, 613-615, 630431 modification of tyrosine residues and, IV, 619-620 papain and, IV, 619 pH and, IV, 618419 pyridoxal phosphate and, IV, 620 Saccharomyces cerevisiae, IV, 640 slime molds, IV, 640 sulfhydryl groups, activation and, IV, 621422 L-Fucose isomerase, properties, VI, 346348 L-Fuculose 1-phosphate aldolase catalytic reaction assay, VII, 314 equilibrium constant, VII, 314 substrate specificity, VII, 314 properties, VII, 313-314 Fumarase catalytic properties active site affinity labeling, P,563564

catalytic site number, V, 562-563 kinetics, V, 552-557 substrate specificity, V, 557-562 general considerations, V, 539-540 historical development, V, 540-541 isotope exchange and, 11, 304-308 mechanism of action, V, 564-568 modification of, I, 140-141 molecular properties amino acid composition, end groups and peptide maps, V, 544-545 dissociation and recombination of subunits, V, 546-549 physical properties, V, 542-544

TOPICAL SUBJECT INDEX

subunit structure, V, 545-546 thiol groups, V, 549-552 preparation and assay, V, 541-542 stereospecificity, 11, 171-176 Fungi, see also Molds aldolase of, VII, 215-216 aspartate transcarbamylases of, IX, 302-306 carboxypeptidases of, 111, 790-791 glutamate dehydrogenases Neurospora crassa, XI, 323-329 others, XI, 329-332 nuclear ribonucleic acid polymerase, X, 310-311 ribonucleases of, IV, 208-239 G

n-Galactarate dehydrase, properties, V, 580-581 Galactonate dehydrase, properties, V, 578-579 Galactose oxidase chemical and physical properties, XII, 528-529 copper of, XII, 529-530 discovery and purification, XII, 527528 inhibitors, XII, 531 mechanism, XII, 532-533 molecular properties and kinetic mechanism, XII. 461-466 optical properties, XII, 530-531 specificity, XII, 531 p-Galac tosidase assay and standardization, VII, 620623 bacterial suspensions and animal material, VII, 624 standard assays and units, VII, 623 transferase assay, VII, 624 chemical properties, VII, 636-639 enzymic properties active site, VII, 657-658 condensation reactions, VII, 660 inhibition studies, VII, 660-661 kinetics, VII, 648-4351 mechanism, VII, 651-657 metal activation, VII, 645-648

other sources, VII, 661-663 pH dependence, VII, 644-645 specificity, VII, 641-644 transfer reaction, VII, 658-660 general background, VII, 618 immunological properties, VII, 839641 mutations and, I, 255-256 occurrence, VII, 619-620 physiochemical properties associated forms, VII, 633-634 denaturation and renaturation, VII, 634-636 size, shape and quaternary structure, VII, 627-633 purification animal tissues, VII, 626-627 Escherichia coli, VII, 624-625 other microorganisms, VII, 625426 Galactosyl transferase biological significance, IX, 377 catalytic properties kinetics, IX, 371-376 reaction catalyzed, IX, 369 substrate specificity, IX, 369-371 purification and properties, IX, 367369 Gastricin, gene duplication and, I, 308 Gastric lipase, properties, VII, 605-606 Gaucher’s disease, acid phosphatase and, IV, 496 Genes cistron and types of mutations, I, 243-245 duplication, I, 3 W 3 0 3 carboxypeptidases, I, 308-309 ethanol dehydrogenaaes, I, 309-311 gastricin, I, 308 a-lactalbumin, I, 311-314 lysozyme, I, 311-314 pancreatic proteinases, I, 303-307 pepsin, I, 308 rennin, I, 308 sulfhydryl proteinases, I, 307-308 Genetic drift, speciation of homologous proteins and, I, 321428 Glucagon, glycogen synthetase and, IX, 341 a-Glucan phosphorylases allosteric transitions, VII, 473-474

496

TOPICAL SUBJECT INDEX

desensitization, VII, 480-482 kinetic and structural basis, VII, 474-480 catalytic reaction nucleotide activation, VII, 439-441 substrate specificity, VII, 437-438 chemical properties amino acid analysis and end groups, VII, 446-447 primary structure, VII, 448-451 enzyme structure, physical properties, VII, 441-446 general considerations, VII, 435-437 mechanism of catalysis kinetics, VII, 460-463 protein functional groups, VII, 463466 vitamin Ba coenzyme, VII, 451-459 regulation associationdissociation phenomena, VII, 469-473 interconversion of a and b forms, VII, 466469 mechanism of allosteric transitions, VII, 473-482 o-Glucarate dehydrase, properties, V, 580 Gluconate dehydratase, properties, V, 578 Glucosamine-6-phosphate isomerase, properties, VI, 314-318 Glucosaminate dehydrase, properties, V, 586

D-c%ICOSeisomerase nature of, VI, 341-344 properties, VI, 349-354 Glucose8phosphatae.e distribution, IV, 600-610 catalytic properties, IV, 565-588 assay methods, IV, 566-567 control of phosphotransferase activity, IV, 592-595 kinetics and reaction mechanism, IV, 572-592 reactions catalyzed, IV, 567-571 thermodynamic considerations, IV, 571-572 detergents and, IV, 556-557 detergent-like effects, IV, 559-560 direct effects, IV, 557-559 modifying effects, IV, 560-562

historical phosphohydrolase activity, IV, 545546 phosphotransferase activity, IV, 546-547 kinetic studies, IV, 572-574 activators and inhibitors, IV, 578582 mechanism, IV, 582-592 pH and, IV, 574-576 substrate concentration, IV, 576-577 temperature, IV, 577-578 membranous nature, possible significance, IV, 562-564 metabolic roles and regulation, IV, 596-597 control in vivo, IV, 597-599 speculation, IV, 599 phospholipids and, IV, 554-556 reactions catalyzed multifunctional nature, IV, 567-568 substrate specificity, IV, 568-571 relation to other enzymes, IV, 552 solubilization and attempted purification, IV, 553-554 Glucose-6-phosphate distribution intracellular, IV, 548-551 phylogenetic and tissue, IV, 547-548 glycogen synthetase and, IX, 341-344 Glucose-6-phosphate isomerase catalytic properties active form of substrate, VI, 293-294 anomerase activity, VI, 294-296 assay, VI, 278-279 kinetic parameters, VI, 289-293 mechanism of action, VI, 296-301 molecular architecture active site amino acids, VI, 285-287 amino acid composition, VI, 278-279 conformational states, VI, 284-285 molecular weight and related properties, VI, 276-278 multiple forms, VI, 281-284 subunit structure, VI, 279-28E occurrence and function, VI, 272-276 Glucosidase-transferase assay of, V, 211-212 glycogen storage disease and, V, 221222

TOPICAL SUBJECT INDEX

Glucosulfatase, V, 11 Glutamate pyrrolidone carboxylate formation from, enzymic, IV, 133-139 nonenzymic, IV, 130-133 D-Glutamate cyclo transferase, pyrrolidone carboxylate formation and, IV, 133-136 L-Glutamate cyclotransferase, pyrrolidone carboxylate formation and, IV, 138 Glutamate dehydrogenase(s), cellular location, XI, 305-306 chemical modification, coenzyme site and specificity, XI, 352-354 cysteine residues, XI, 347-348 histidine residues, XI, 346-347 lysine residues, XI, 343-346 other modifications, XI, 348-349 spectrophotometric studies, XI, 349 substrate site, XI, 349-352 composition, XI, 320-322 distribution and coenzyme specificity, XI, 296-297 animals, XI, 300-305 bacteria, XI, 297-299 fungi, XI, 297 plants, XI, 299-300 electrophoretic and spectrophotometric properties, XI, 322-323 function and equilibrium, XI, 294-295 genetics and regulation of enzyme synthesis, bacteria, XI, 332-334 fungi, XI, 323-332 plants, XI, 334-335 historical background, XI, 295-296 kinetic studies, XI, 354-357 metamorphosis and, XI, 306 modification of, I, 137 oligomer structure, hexameric model, XI, 315-317 immunochemistry, XI, 317-318 molecular weight, XI, 314-315 trimer formation, XI, 317 polymerization, mechanism, XI, 308, 310-312 nucleotide effects, XI, 312-314 significance, XI, 307-308 primary structures, XI, 335-343

purification of, XI, 307 reaction mechanism, XI, 357-360 regulation of, I, 443-444, XI, 360-366 stability, denaturation and dissociation, XI, 319-320 tertiary structure, XI, 318-319 Glutamate mutase assay, VI, 524-525 catalytic properties conditions affecting activity, VI, 531-532 equilibrium, VI, 532 interaction of components and coenzyme, VI, 528-530 mechanism, VI, 532-534 substrate and coenzyme specificity, VI, 530-531 purification and molecular properties component E, VI, 525-526 component S, VI, 526-528 stereochemistry, 11, 206-210 Glutaminase(s) Azotobacter agik's, IV, 97-98 Escheiichia coli, IV, 80-81 acyl transferase reactions, IV, 84-85 assay, IV, 81 deuterium oxide and, IV, 90 6-diazo-5-oxonorleucine and, IV, 85-87 mechanism of action, IV, 90, 92-93 occurrence, IV, 81 other inhibitors, IV, 87 pH effects on kinetic parameters, IV, 88,89 purification, IV, 82 relationship to other acylases, IV, 90, 91 specificity, IV, 82-84 temperature and, IV, 88 survey, IV, 93-95 Glutamine occurrence and function of, X, 699704, 755-757 pyrrolidone carboxylate formation from enzymic, IV, 139-141 nonenzymic, IV, 130-133 L-Glutamine cyclotransferase, pyrrolidone carboxylate formation and, IV, 139-141

498

TOPICAL SUBJECT INDEX

Glutamine synthetase, X, 757-759 active site mapping, X, 720-733 adenylylation and deadenylylation, VIII, 4044 historical development of problem, X, 784-786 regulatory protein, X, 786-787 uridylylation and deuridylation of regulatory protein, X, 787-789 adenylylation effect biosynthetic activity, X, 794-796 y-glutamyltransferase activity, X, 796-800

adenylylation state determination enayme assays, X, 802-803 spectral analysis, X, 802 cascade control, X, 789-792 covalent modification of, I, 409-413 cumulative feedback inhibition, X, 776 mechanism, X, 777-780 divalent cation control adenosine triphosphate: cation ratio,

x,783-784

relaxation phenomena, X, 781 specificity, X, 781-783 general catalytic properties, X, 708-709 hybrid forms, X, 804-807 intermediate formation of y-glutamyl phosphate, X, 716-720 mechanism, X, 733-743, 759-763 multiple molecular forms, X, 800-801 determination of state of adenylylation, X, 802-803 heterologous interaction, X, 804-807 preparation of partially adenylylated form, X, 801-802 partially adenylylated direct isolation, X, 801 in vitro preparation, X, 801 mixing of forms, X, 802 subunit dissociation and reassociation, X, 802 partial reactions adenosine triphosphate formation,

X, 714-715 arsenolysis of glutamine, X, 713 cycloglutamyl phosphate synthesis, X, 715 exchange reactions, X, 709-710

methionine sulfoxime phosphate formation, X, 714 boxoproline formation, X, 713-714 physical and chemical properties, X, 704-708

physicochemical properties, X, 792-793 adenylylation site, X, 793-794 pyrrolidone carboxylate formation by, IV, 136-137 regulation of, X, 743-754, 775-792 repression of formation, X, 780-781 structure chemical composition, X, 769-771 physical characteristics, X, 763-769 taut, relaxed and tightened forms, X, 771-775

Glutaminyl peptides, pyrrolidone carboxylate formation from, IV, 139-141 y-Glutamyl amino acids, pyrrolidone carboxylate formation from, IV, 142-146

y-Glutamyl cyclotransferase, pyrrolidone carboxylate formation and, IV, 141-146

y-Glutamyl-cysteine synthetase general catalytic properties, X, 676-678 mechanism, X, 683-687 partial reactions adenosine triphosphate hydrolysis,

X, 678-679 enayme4activated glutamatelcomplex formation, X, 683 exchange reactions, X, 680-681 5-oxoproline formation, X, 679-680 phosphorylation of methionine sulfoxime, X, 681-683 pyrophosphate synthesis, X, 679 physical properties, X, 675-676 pyrrolidone carboxylate formation and, IV, 136-137 sources, purification and assay, X, 674-675

y-Glutamyl lactamase, see y-L-Glutamyl cyclotransf erase y-Glutamyltransferase agaricaceae, IV, 95-96 survey, IV, 93-95 y-Glutamyl transpeptidase kidney, IV, 96-97

499

TOPICAL SUBJECT INDEX

pyrrolidone carboxylate formation and, IV, 141 Glutathione S-epoxidetransferase, VII, 205-206

catalytic properties factors influencing, VII, 209-210 kinetics and specificity, VII, 209 measurement of reaction, VII, 208 notes on mechanism, VII, 210 products of reaction, VII, 208 molecular properties molecular weight, VII, 207 multiple forms, VII, 206-207 purification, VII, 206 stability, VII, 208 Glutathione reductase kinetic studies, XIII, 138-141 metabolic functions, XIII, 129-133 thiol groups, XIII, 141-142 two-electron-reduced enzyme, properties, XIII, 133-138 Glutathione synthetase acyl phosphate intermediate, X, 690-692

general catalytic properties, X, 689-690 historical background, X, 671-674 mechanism, X, 693-695 partial reactions, X, 693 physical properties, X, 688-689 Glyceraldehyde-&phosphate dehydrogenase, historical background, XIII, 2-3 isolation, XIII, 3-4 mechanism of action other activities, XIII, 44-49 physiological activity, XIII, 38-44 metabolic role, XIII, 45-49 pyridine nucleotide binding, XIII, 28-30

cooperativity of, XIII, 30-35 preexisting asymmetry model, XIII, 35-38

reaction catalyzed, XIII, 1 structure apoenzyme, XIII, 19-20 chemical modification, XIII, 20-24 dissociation and hybridization, XIII, 24-27 holoenzyme X-ray structure, XIII, 9-19

primary, XIII, 5-9 Glycerate kinases assay and distribution, VIII, 504-505 catalytic properties, VIII, 507-508 metabolic role, VIII, 505-506 molecular properties purification and state of purity, VIII, 506-507 stability, VIII, 507 Glycerol kinases assay and distribution, VIII, 488492 catalytic properties product inhibition, VIII, 501-502 substrate specificity and kinetics, VIII, 497-501 thermodynamics, VIII, 502 metabolic role, VIII, 492-493 molecular properties chemical modification, VIII, 496-497 composition, VIII, 494 purification and state of purity, VIII, 493494 size and subunit structure, VIII, 494-495

stability, VIII, 495-496 regulation mammals, VIII, 504 microorganisms, VIII, 502-503 L-Glycerol-3-phosphate dehydrogenase, properties, XIII, 256-260 Glycidol phosphate, enolase and, V, 534 Glycinamide ribonucleotide transformylase, properties, IX, 203-204 Glycine, oxidative decarboxylation of, IX, 221-223 Glycine amidotransferase biological distribution, IX, 498499 catalytic properties reaction mechanism, IX, 500-503 substrate specificity, IX, 499-500 regulation, IX, 503305 Glycogen structure determination, debranching enzymes and, V, 228-234 Glycogen phosphorylase, covalent modification of, I, 413415 regulation of, I, 441442 Glycogen storage disease, glucosidasetransferase and, V, 221-222

500

TOPICAL SUBJECT INDEX

Glycogen synthetase adrenal, IX, 353 assay, IX, 317418 association with glycogen, IX, 311-312 blood erythrocytes, IX, 355 leukocytes, IX, 354-355 platelets, IX, 355 brain, IX, 353-354 control of activity epinephrine effect, IX, 338, 341, 351-352

glucagon and, IX, 341, 351-352 glucose and, IX, 347-348 glycogen and, IX, 336, 340 insulin activation, IX, 336-338, 340-341, 348-351

muscle contraction events, IX, 338-340

donor specificity, IX, 318 general properties of a and b forms, IX, 324-325 glucosyl acceptor de novo synthesis, IX, 320-321 oligosaccharides, IX, 320 polysaccharides, IX, 319-320 heart, IX, 340-341 historical background, IX, 310-311 inactive form, IX, 330-331 kinase, IX, 326-327 liver, IX, 341-353 muscle, IX, 332340 nomenclature, IX, 322-324 nonmammalian organisms, IX, 357-361 phosphatase liver, IX, 328-330 muscle, IX, 327-328 physicochemical properties muscle, IX, 313-315 other tissues, IX, 316-316 properties of the two forms activity of, IX, 335-336 glucose 6-phosphate and, IX, 332-334, 341-344

inorganic phosphate and, IX, 334, 344-345

magnesium and other ligands, IX, 335, 345

nucleotides, IX, 334-335, 345 pH and, IX, 335, 346

physiological conditions and, IX, 348347

proteolytic inactivation, IX, 331 purification, IX, 312-313 reaction catalyzed, IX, 316-317 reaction mechanism, IX, 321-322 regulation of, I, 441-442 spleen, IX, 354 system of interconversion, IX, 325-326 tumors, IX, 356-357 tissues sensitive to sex hormones, IX, 355-356

Glycosidases, modification of, I, 128-129 Glycosulfatase (s) ccllulose polysulfatase, V, 11-12 cerebroside sulfatase, V, 13-14 chondrosulfatase, V, 12 glucosulfatase, V, 11 Glyoxylate condensing enzymes, general remarks, VII, 411412 Gout, molybdenum hydroxylase genetics and, XII, 400-402 Gram-negative bacteria phage lytic enzymes Aerobacter cloacae, V, 398 Azotobacter, V, 397498 Klebsiella pneumoniae, V, 394-395 Pseudomonas, V, 395-397 Salmonella, V, 398 Gram-positive bacteria phage lytic enzymes Bacillus megaterium, V, 408410 Bacillus stearothermophilus, V, 410-411

Micrococcus lysodeikticus, V, 401402 staphylococci, V, 398401 streptococci, V, 402-408 Guanidine hydrochloride, inorganic pyrophosphatase and, IV, 508-510 Guanidino kinase(s) bivalent metal ions and, VIII, 471473 catalytic reaction, VIII, 459 chemical stop assays forward reaction, VIII, 464 reverse reaction, VIII, 465 continuous recording assays potentiometric, VIII, 466 spectrophotometric, VIII, 465406 discovery and isolation, VIII, 459461 distribution and function, VIII, 461-464

TOPICAL SUBJECT INDEX

equilibrium, VIII, 485-486 function of amino acid residues cysteine, VIII, 477-480 others, VIII, 480-482 historical background, VIII, 457-459 isotopic assays, VIII, 465 mechanism, VIII, 482-485 molecular properties amino acid composition, VIII, 469470

immunological reactions, VIII, 470-471

molecular weight and subunit composition, VIII, 466-468 stability, VIII, 468-469 pH optimum, VIII, 476-477 substrate specificity general comments, VIII, 473 guanidines, VIII, 474-476 nucleotides, VIII, 476 Guanine aminohydrolaw, IV, 76-77 Guanosine aminohydrolase, IV, 77-78 Guanosine diphosphate-n-mannose oxidoreductase properties, V, 479 Guanosine kinase, see Inosine-guanosine kinase Guanosine monophosphate, synthesis, VIII, 39-40 Guanosine monophosphate kinase assay, IX, 84 distribution and purification, IX, 82-84 kinetic and molecular properties, IX, 84-85

substrate specificity; reaction mechanism, IX, 85-86 Guanosine 5'-phosphate reductase, modification of, I, 137 Guinea pig serum, asparaginase of, IV, 105-107 H

Heart aspartate transaminase of, IX, 398-462 glycogen synthetase of, IX, 340-341 mitochondria1 adenosine triphosphatase catalytic properties, X, 382-383 cold inactivation, X, 380 inhibitors and activators, X, 385-386 molecular properties, X, 378-380,381

501 nature of active site, X, 383 purification and stability, X, 377-378 5'-nucleotidase of, IV, 347-348 succinyl coenzyme A synthetase of, X, 591-594 Heavy atom derivatives globular macromolecules factors in derivative formation, I, 73-86

preparation and classification, I, 70-73

useful heavy atom compounds, I, 73 Helicorubin, properties, XI, 592 Heme proteins, modification of, I, 142 Hemoglobin allosteric properties, I, 388-390 mutations and, I, 257-259 Hemoprotein(s), cytochro,me M i k e , XI, 576-577 Hexarate dehydrase(s), properties, V, 579-581

Hexokinase (s) aggregation phenomenon, IX, 40-41 amino acid composition and essential groups, IX, 41 chemical studies comparison of native isoenzymes, IX, 10-12 nature of proteolytic modification, IX, 12-13 comparative aspects, IX, 46-48 isoelectric point, IX, 41 isoenzymes and modification by endogenous proteases, IX, 2-6 mechanism adenosine triphosphatase reaction, IX, 18-19 conformational changes, IX, 27-28 enzyme-substrate interaction and, IX, 43-44 equilibrium measurements, IX, 19-22 kinetic studies, IX, 13-17, 41-43 phosphoenzyme intermediate, IX, 23-24, 44

sulfhydryl groups and, IX, 25-27 modification by added proteases, IX, 6-7

molecular weight and subunit structure, IX, 39-40

TOPICAL SUBJECT INDEX

molecular weight in nondenaturing solvents, IX, 7-8 multiple forms, IX, 31-33 purification procedures, IX, 37-38 regulation, IX, 29-31, 4446 relation of soluble to insoluble forms, IX, 33-37 subunit size, IX, 8-10 Hill plots, enzyme regulation and, I, 358-359

Hirudin, thrombin and, 111, 304405 Histidine ammonia-lyase catalytic process function of prosthetic group, VII, 159-162

metal ion activation, VII, 162-163 prosthetic group, VII, 154-159 rate limiting step, VII, 164-166 reaction sequence, VII, 148-154 distribution, purification and kinetic properties, VII, 137-142 mechanism of action, VII, 195-196 size and constitution, VII, 146-148 Histidine deaminase, role of metals, 11, 509-510

Histidine decarboxylase, Schiff base and, 11, 356-357 Histidine residues aspartate transcarbamylase, IX, 267 chemical modification, I, 173 chymotrypsin catalytic reaction and, 111, 234 chemical modification, 111, 234 pH and, 111, 231-233 creatine kinase, VIII, 434 phosphofructokinase, VIII, 272 ribonuclease, IV, 685-689 subtilisin, 111, 580-584 Histone kinases, properties, VIII, 579-580 Homoarginine residues, proteinase inhibitors and, 111, 461-452 Homocitrate synthase, properties, VII, 425-426

Homocysteine synthetase(s1, properties, VII, 61-62 Homoserine dehydratase, properties, VII, 67-59 Homoserine dehydrogenase, see Aspartokinase

Hormones, phosphofructokinase and, VIII, 277-278 Hyaluronate-endo-p-glucuronidase, V, 313 Hyaluronate lyase, V, 313-314 Hyaluronidase (s) bacterial, V, 309 biological effects, V, 319-320 historical background, V, 307-308 kinetics, V, 318-319 leech, V, 313 lysosomal, V, 312 microbial, V, 313-314 preparation and assay, V, 315-317 properties, V, 318 submandibular, V, 312-313 substrates additional, V, 311 chondroitin, V, 310 chondroitin 4- and B-sulfate, V, 310 dermatan sulfate, V, 310-311 hyaluronate, V, 309 testicular, V, 311-312 testicular type, V, 308,311-313 Hydrogenase, proton transfer and, 11, 317-318

Hydrogen bonds, propinquity effects and, 11, 254-264 Hydrolase(s), role of metals in mechanism, 11, 519-525 D-2-Hydroxy acid dehydrogenase, properties, XIII, 272-273 erythro-p-Hydroxyaspartate dehydratase, properties, VII, 47-48 m-Hydroxybenzoa te-4-hydroxylase, properties, XII, 225 m-Hydroxybenroa te-6-hydroxylase, properties, XII, 224-225 p-Hydroxybenzoate hydroxylase, properties, XII, 211-216 8-Hydroxydecanoyl thioester dehydrase distribution, V, 464 function of, V, 441443 general considerations, V, 441443 inhibition by substrate analogs, V, 455-456

acetylenic inhibitor reaction site, V, 456-458

3-decynoyl-N-ace tylcysteamine and fatty acid synthetase, V, 461-463

TOPICAL SUBJECT INDEX

inhibitor specificity, V, 458459 isomeriration of 3-decynoyl-Nacetylcysteamine, V, 459-461 modification of, I, 141 mutants, V, 463 protein structure active site, V, 453-455 disulfide bridges, V, 452-453 nitration, V, 455 reaction mechanism isotope effects and labeled substrates, V, 449-450 kinetic studies, V, 446-449 trapping of intermediates, V, 451-452 substrate specificity, V, 444-445 optical and geometric, V, 446 thioester, V, 445-446 4-Hydroxy-2-ketoglutarate aldolase, properties, VII, 299-301 5-Hydroxymethyl deoxycytidine monophosphate kinase, properties, IX, 94-95 8-Hydroxy-8-methylglutarylcoenzyme A cleavage general, VII, 432-433 properties of beef liver enzyme, VII, 433-434 ,8-Hydroxy-8-methylglutaryl coenzyme A synthase distribution, VII, 429 general considerations, VII, 427428 properties of yeast enzyme, VII, 429-431 stereochemistry of product, VII, 428 p-Hydroxyphenylpyruvate hydroxylase, XII, 179-180 catalytic properties, XII, 180-183 purification and molecular properties, XII, 180 I

Imidazole rings, trypsin, 111, 269-270 Imidazolylacetate monooxygenase, properties, XII, 225-226 Imido adenylate derivatives synthesis, VIII, 37 argininosuccinate, VIII, 38-39 guanosine monophosphate, VIII, 39-40

503 Immunochemistry, chemical modification and, I, 199-200 Inclusion compounds, propinquity effects and, 11, 274-279 Indoleamine 2,3-dioxygenase historical, XII, 130-131 molecular and catalytic properties, XII, 131-132 Influenza virus neuraminidase isolation and purification, V, 328 purification of virus particles, V, 327-328 Infrared spectroscopy, enzyme structure and, 11, 374-379 Inhibition studies nomenclature and basics, 11, 18-21 prediction of patterns, 11, 21-25 types of experiments, 11, 25-43 Initial velocity studies bireactant, 11, 7-10 prediction of patterns, 11, 10-13 terreactant, 11, 13-18 Initiation factors discovery and function, X, 6-12 eukaryote miscellaneous, X, 42-43 ribosome associated, X, 29-33 supernatant, X, 33-42 initiation cycle and, X, 12-13 properties factor-1, X, 26-28 factor-2, X, 13-17 factor-3, X, 7-26 Inorganic ions metabolic regulation and divalent cations, I, 420-423 monovalent cations, I, 419-420 Inorganic pyrophosphatase catalytic properties assay, IV, 534 interaction with inhibitors, IV, 525-526 ion and inhibitor effects, IV, 518-519 kinetics, IV, 535-538 mechanism, IV, 538-539 nature and binding of active substrate and role of magnesium, IV, 522-525 pH effects, IV, 518

504

TOPICAL SUBJECT INDEX

reversal of reaction, IV, 519-520 substrate specificity and stoichiometry, IV, 520-522,534-535 chemical composition amino acid content, IV, 512 cyanogen bromide cleavage products, IV, 514 N- and C-terminal amino acids, IV, 512-514

tryptic digest maps, IV, 514 chemical modification, IV, 514-515 cyanate and diazonium-1H-tetrazole, IV, 516-518 2,4&trinitrobenzene sulfonate, IV, 515-516

molecular properties electron microscopy, IV, 506-508 homogeneity, IV, 502-503 metalloenzyme, IV, 532-534 physicochemical parameters, IV, 530-531

purification, IV, 501-502, 530 reconstitution from subunits, IV, 510-612

reversible divalent cation binding, IV, 531-532 size, IV, 504 other, IV, 539-541 physical properties optical properties, IV, 505-506 sedimentation and diffusion coefficients, IV, 504 viscosity, IV, 505 subunits in quanidine hydrochloride optical properties, IV, 510 sedimentation and diffusion coefficients and intrinsic viscosity, IV, 509

size, IV, 509 Inosamine phosphate amidinotransferase biological distribution, IX, 505-506 catalytic properties reaction mechanism, IX, 507-508 substrate specificity, IX, 506-507 regulation, IX, 508-509 Inosine-guanosine kinase, properties, IX, 54-56 Inosinic acid dehydrogenase, modification of, I, 137-138

Insects, alcohol dehydrogenase of, XI, 189-190

Insulin, glycogen synthetase and, IX, 336338, 340-341

Intestine monoglyceride lipase of, VII, 603-605 5'-nucleotidase of, IV, 345 Intramolecular reactions, miscellaneous, 11, 238-250 Invertase, V, 292-293 biosynthesis, V, 294-295 catalytic properties, V, 305 active site, V, 300-301 inhibitors, V, 301-302 kinetics, V, 302 mechanism, V, 302-303 determination, V, 292 localization and multiple forms, V, 293-294

properties, V, 298-300 purification, V, 295-298, 304-305 Invertebrates, cytochrome b-like pigments of, XI, 564 Ionizable groups, protein structure and, 11, 430-440 Iron-sulfur proteins, categories conjugated, XII, 3-4 ferredoxin, XII, 2 high potential, XII, 2-3 eight irons background, XII, 37-39 chemical properties, XII, 4 4 4 6 physical properties, XII, 3944 four irons per center background, XII, 31-33 high potential proteins, XII, 35-37 low potential proteins, XII, 34 Iron-sulfur enzymes, XII, 47-50 mitochondrial, XII, 56 model compounds, XII, 48-47 nitrogenase system, XII, 60-56 nomenclature, XII, 2 xanthine oxidase, XII, 56 Isoamylase (8) plant, V, 208 pseudomonad and Cytophaga, V, 204-206

yeast, V, 206-208

TOPICAL SUBJECT INDEX

Isocitrate dehydrogenase, stereospecificity, 11, 168-171 Isocitrate lyase, VII, 382-383 assay and isolation, VII, 383 molecular properties cofactors, VII, 384 inhibitors, VII, 385 molecular weight and subunits, VII,

proton transferase and, 11, 283-285 rate equations, 11, 63-65

Jack bean ureaae of, IV, 2-5

384

sulfhydryl groups, VII, 385 Isoleucine residues chymotrypsin chemical modification, 111, 238-239 conformation and, 111, 239-243 pH and, 111, 236-237 chymotrypsinogen, 111, 175-176 Isomerase(s) aldo-keto, proton shifts and, 11,290-295 Isopentylpyrophosphate isomerase assays, evaluation, VI, 567-568 function, VI, 566-567 mechanism, VI, 570-572 occurrence, VI, 565-566 properties inhibitors, VI, 569 metal ion and, 569 Michaelis constant, VI, 569 pH optimum, VI, 568 reversibility and position of equilibrium, VI, 569-570 synthesis of substrate and product, VI, 567

dsopropylmalate synthase properties, VII, 423 effect of leucine, VII, 424-425 Isotope effects primary, 11, 284-285 examples, 11, 325-329 origin and magnitude, 11, 322-324 pepsin and, 111, 180 secondary, 11, 285-287 examples, 11, 331-333 origin and magnitude, 11, 329-331 Isotopic exchange kinetics basic considerations, 11, 4 3 4 4 ping pong mechanism, 11, 48-52 sequential mechanism, 11, 44-48 lack of, 11, 287

K

Kallikreins, 111, 482-483 Keratinase, microbial, 111,763-765 p-Keto acid decarboxylase(s), Schiff bases and, 11, 359 p-Ketoacyl acyl carrier protein synthetase catalytic properties assays, VIII, 190 mechanism, VIII, 194-199 pH optimum, substrate specificity and kinetics, VIII, 190-194 historical background, distribution and metabolic significance, VIII, 188-189

molecular properties, VIII, 189-190 a-Ketobutyrate synthase, properties, VI, 203-204

2-Keto-3-deoxy-~-arabonate aldolase, properties, VII, 296 2-Keto3-deoxy-~-arabonate dehydratase, properties, V, 583-585 2-Keto-3-deoxy-o-fuconate aldolase, properties, VII, 296 2-Keto-3-deoxy-D-glucarate aldolase properties, VII, 297 5-Keto-4deoxyglucarate dehydratase, properties, V, 585-586 2-Keto3-deoxy-manno-octosonate aldolase, properties, VII, 298 2-Keto-3-deoxy-6-phosphogalactonic aldolase, properties, VII, 295-296 2-Keto3deoxy-6-phosphogluconic aldolase general properties Pseudomonas putida, VII, 283-285 Pseudomonas saccharophila, VII, 285 mechanism isotope exchanges, VII, 290-291 oxalacetate decarboxylation, VII, 291-292

TOPICAL SUBJECT INDEX

role of an additional base, VII, 292-295 role of Schiff base formation, VII, 289-290 modification of, I, 139 structure other characteristics, VII, 287-288 subunit composition and arrangements, VII, 285-287 3-Keto-dihydrosphingosine, formation of, VII, 355-356 a-Ketoglutarate dehydrogenase complexes, I, 224-225 a-Ketoglutarate synthase, properties, VI, 201-203 a-Ketoisovalerate synthase, properties, VI, 205 A'3-Ketosteroid isomerase animal tissue catalytic mechanism, VI, 617-618 distribution and properties, VI, 615-617 catalytic mechanism hydrogen isotopes and, VI, 605-607 proposal for, VI, 612-615 ultraviolet and fluorescence studies, VI, 607-612 catalytic properties active-sitedirected inhibitor, VI, 601 competitive inhibitors, VI, 6oo-gO1 kinetic studies, VI, 604 metal ions, chelators and urea, VI, 800 pH effects, VI, 602 reversibility, VI, 603-604 substrate specificity, VI, 59WOo thermodynamic parameters, VI, 602-603 historical, VI, 591-592 molecular properties chemical modification, VI, 595 induction, purification and crystallization, VI, 592-593 structure, VI, 598-599 ultracentrifugation, VI, 593-594 ultraviolet and fluorescence spectra, VI, 594-595 Kidney argininosuccinase of, VII, 178-179

fructose-1,6-diphosphatase purification and properties, IV, 629-630 regulation, IV, 630-631 7-glutamyl transpeptidase of, IV, 96-97 vitamin Ba methyltransferase of, IX, 164-165 Kineticist tools of, 11, 6-7 inhibition studies, 11, 18-43 initial velocity studies, 11, 7-18 isotope exchange, 11, 43-52 miscellaneous, 11, 52-61 Kinetics, basic theory, 11, 6 nomenclature and notation, 11, 2-5 Kinetic systems far from equilibrium computer solutions to rate equations, 11, 74-83 integrated rate expressions, 11, 71-74 single substrate-single product Michaelis-Menten mechanism, 11, 74-79 near equilibrium relaxation amplitudes, 11, 99-108 relaxation spectra, 11, 83-99 Kinin-destroying enzymes, 111, 483 Kinin-forming enzymes catheptic kininogenases, 111, 483 kallikreins, 111, 482483 Klebsiellu pneumoniae phage polysaccharide depolymerase enzyme properties and role, V, 394-395 preparation and assay, V, 394 Klotz plots, enzyme regulation and, I, 359-361 Kynurenine3hydroxylase, properties, XII, 230-231

1

a-Lactalbumin bovine, VII, 681-682 gene duplication and, I, 311-314 lactose synthetase and, IX, 366 8-Lactamase, see Penicillinase Lactate dehydrogenase, away, XI, 199-200

507

TOPICAL SUWECT INDEX

coenzyme binding oxidized, XI, 280-281 reduced, XI, 277-280, 281 fluorescence and spectroscopy coenzyme, XI, 268 protein, XI, 264-268 histidine-195, state of protonation, XI, 289 historical, XI, 192-195 isolation, XI, 200-202 isozymes, evolution of genes, XI, 198-199 general, XI, 195 known genes, XI, 195-198 D-lactate specific enzymes, XI, 199 kinetic studies, transient phase, XI, 57-59 mechanism, XI, 289-292 modification, I, 136 crystals, XI, 261 mild proteolysis, XI, 257-258 side chains, XI, 258-261 physical properties, XI, 261-264 reactions catalyzed cyanide addition, XI, 270 glyoxylate and XI, 269-270 ketopyruvate reduction, XI, 268-269 reviews, XI, 192 steady-state kinetics noninhibiting substrate concentrations, XI, 270-273 steady-state inhibitors, XI, 273-274 structure, amino acid sequence, XI, 202-205 composition, XI, 202, 203 oligomer, XI, 250-257 threedimensional structure, XI, 205-250

substrate binding, XI, 290-291 absence of enzyme-substrate compounds, XI, 281-282 active ternary complex, XI, 284-286 enzyme-anion complexes, XI, 282 enzyme :coenzyme :inhibitor complex, XI, 282-284 ternary complex transient kinetics, XI, 286-289 substrate inhibition lactate. XI. 275-276 pyruvate, XI, 274-275

significance of abortive complex, XI, 276-277

D(-)-Lactate

dehydrogenase, XIII,

269-270

enzymic properties, XIII, 270-272 physical properties, XIII, 270 L( +)-Lactate dehydrogenase cytochrome b, core, XIII, 266-267 enzymic properties, XIII, 267-269 historical background, XIII, 263-264 physical properties, XIII, 264-266 Lactate oxidase, properties, XII, 194-199 Lactic acid racemase, properties, VI, 380 Lactobacillus, histidine decarboxylase of, 11, 356-357 Lactose synthetase historical background, IX, 364-365 requirement for two proteins identification of a-lactalbumin, IX, 366

relationship to lysozyme, IX, 366-367 resolution of, IX, 365466 Leaves, adenosine diphosphoryl glucose pyrophosphorylase of, VIII, 86-90 Leech, hyaluronidase of, V, 313 Leucine aminopeptidase assay, 111, 83-84 chemical properties, 111, 88-89 enzymic properties inhibitors, 111, 96-98 mechanism of action, 111, 98-100 role of metal ion, 111, 89-91 substrate specificity and kinetics, 111, 91-96 historical background, 111, 82-83 physical properties, 111, 86-88 purification, 111, 84-86 use in sequence studies, 111, 101-102 Lipase(s1 activity, determination of, VII, 578680

castor bean, VII, 613-614 catalytic properties active site, VII, 593-595 chemical structure of substrate, VIZ, 595-599

colipase and, VII, 595 effectors and, VII, 594601 ester bond synthesis and, VII, 601602

508

TOPICAL SUBJECT INDEX

phosphoglycerides and, VII, 602-603 positional specificity, VII, 591-593 substrate physical state, VII, 586591

definition of, VII, 575-577 distribution animal, VII, 577 microorganisms, VII, 578 plant, VII, 577-578 gastrointestinal gastric, VII, 605-606 intestinal, VII, 603-605 microorganisms, VII, 614-616 milk, VII, 611-613 pancreatic, VII, 580-581 catalytic properties, VII, 586-603 molecular properties, VII, 583-586 purification, VII, 581-583 tiasue adipose tissue hormone sensitive, VII, 609-610 lipoprotein lipase, VII, 606-609 liver lipases, VII, 611 Lipoamide dehydrogenase distribution, XIII, 106-107 kinetic studies, VIII, 115-117 mechanism, XIII, 126-129 mechanism of Maasey and Veeger, review of, XIII, 111 metabolic functions, XIII, 107-110 role of NAD' as modifier, XIII, 117120

structural studies, XIII, 120-126 two-electron-reduced enzyme, properties, XIII, 111-115 Lipoate, activation of, 17-18 Lipoprotein lipase, properties, VII, 606609

Liver acetyl coenzyme A carboxylase of, VI, 78-79

acid phosphatases, IV, 484-493 alcohol dehydrogenase of, XI, 20-22, 56-57, 107-109, 111-117

aldolase, VII, 241-244 argininosuccinase of, VII, 171-178, 179 fructose-1,6-diphosphatase, IV, 618-633 glycogen synthetase of, IX, 341-353 lipases, VII, 611 mitochondria1 adenosine triphosphatase

catalytic properties, X, 388-389 molecular properties, X, 388 purification, X, 387-388 5'-nucleotidase of, IV, 343-345 sulfatase A of, V, 27-36 sulfatase B of, V, 37-38 sulfite oxidase of, XII, 414-419 vitamin Bn methyltransferase of, IX, 164-165

London forces, propinquity effects and, 11, 254-264 Long chain fatty acid:coenayme A ligase catalytic properties partial and exchange reactions, X, 486-487

substrates and inhibitors, X, 485-486 Luciferase, bacterial, properties, XII, 226-229

Luciferin, adenylylation of, VIII, 19-20 Lyase(s1, role of metals in mechanism, 11, 508-519 Lysine, biosynthesis, VI, 504 Lysine monooxygenase, properties, XII, 199-203, 293-294

n-cu-Lysine mutase, VI, 551-552 cofactor requirements, VI, 552 comparison to L-plysine mutase, VI, 554-555

cofactor requirements, VI, 555-559 2,5diaminohexanoate and, VI, 554 inhibitors, VI, 553 partial reactions, VI, 553-554 purification of complex, VI, 552 L-p-Lysine mutase assay, VI, 548-549 cofactor requirements, VI, 550 comparison to n-a-lysine mutase, VI, 554-555

cofactor requirements, VI, 555-559 inhibitors, VI, 551 occurrence, VI, 548 purification and physical properties, VI, 549-550 reversibility, VI, 551 Lysine residues acylation biotin activating enzyme, VIII, 18 lipoate activating enzyme, VIII, 1718

creatine kinase, VIII, 432-434

TOPICAL SUBJECT INDEX

ribonuclease, IV, 801 subtilisin, modification of, 111, 596-598 Lysosomes, hyaluronidase of, V, 312 Lysozyme, see abo Phage lysozyme activity biological role, VII, 669-670 muramidase and chitinase, VII, 667669 analysis of rate enhancement, VII, 864-865, 868 general acid catalysis, VII, 885 ion pairing and, VII, 867 substrate distortion, VII, 865-867 association with hydrogen ions, VII, 745-746 apparent ionization constants, VII, 736-739 enthalpy and volume changes, VII, 735-736 environment of ionizable groups, VII, 739-745 isionic and isoelectric pH, VII, 732 potentiometric titrations, VII, 732735 association with other ions and molecules, VII, 746-755 avian, VII, 677-4380 bacteriophage T2 catalytic properties, V, 381-382 chemical properties, V, 380-381 physicochemical properties, V, 380 purification, V, 379 bacteriophage T4 catalytic properties, V, 369-374 chemical properties, V, 366-369 enzyme assays, V, 361464 physicochemical properties, V, 366 purification, V, 364-366 role in life cycle, V, 375-379 bond rearrangement mechanism, VII, 855-856 chemical modification, I, 207-211 chemical modifications amino groups, VII, 780-785 arginine, VII, 785-786 carboxyl groups, VII, 786-788 cystine, VII, 789-790 histidine, VII, 791 methionine, VIJ, 791-792 other reactions, VII, 797-798

509 tryptophan, VII, 792-796 tyrosine, VII, 796-797 chromophore properties, VII, 799-802 absorbance, VII, 802-803 chromophore exposure, VII, 809 circular dichroism, VII, 807-809 fluorescence, VII, 803-806 solvent and saccharide perturbation, VII, 809 conformation of egg-white model main polypeptide chain, VII, 692699 side chains, VII, 699-707 water structure, VII, 707 crystallographic studiea of inhibitor complexes, VII, 707-708 a- and p-N-acetylglucosamine, VII, 708 enzyme-substrate complex structure, VII, 712-714 supplementary binding studies, VII, 714-717 tri-N-acetylchitotriose, VII, 708-711 denaturation, VII, 760-766 guanidine hydrochloride, VII, 774776 nonaqueous solvents, VII, 776-777 nuclear magnetic resonance, VII, 777-780 properties, VII, 770-772 stability, VII, 766-770 thermal, VII, 772-774 urea, VII, 776 early history, VII, 666-667 fluorescence spectroscopy, 11, 426-429 gene duplication and, I, 311-314 hen egg-white amino acid sequence, VII, 672-676 composition, VII, 672 preparation and purification, VII, 670-672 synthesis, VIT, 67M77 human, VII, 680-881 hydrodynamic measurements, shape and solvation, VII, 717-725 hydrogen exchange, VII, 730-732 large substrates bacterial cells and cell walls, VII, 836-841 chitin and derivatives, VII, 841-843

TOPICAL SUBJECT INDEX

low molecular weight substrates, VII, 846-847

bond cleaved, VII, 847 chain length and, VII, 847-848 character of substrate, VII, 848-849 Hammett constant, VII, 851-852 isotope effect, VII, 850 kinetic constants, VII, 850-851 kinetics of saccharide binding, VII, 852-855

pH and, VII, 849-850 rate enhancement, VII, 850 temperature and, VII, 849 mechanism, VII, 856-857 acetamido participation, VII, 863864

saccharide binding, VII, 808-835 substrates, kinetics and mechanism, VII, 836-868 X-ray studies, VII, 682-717 X-ray studies, I, 67-69 analysis of structure, VII, 6824392 conformation of egg-white model, VII, 692-707 crystallography of inhibitor complexes, VII, 707-717 Lysyl hydroxylase catalytic properties, XII, 166-167 purification and molecular properties, XII, 165 o-Lyxose isomerase, properties, VI, 344345

environment, VII, 863 general acid catalysis, VII, 857-860 glycosyl carbonium ion and, VII, 862-863

location of catalytic center, VII, 857 substrate distortion, VII, 860-862 optical properties, VII, 725-730 relationship to a-lactalbumin, IX, 366367

saccharide binding, VII, 809-810 acceptor and substrate reactivity, VII, 822-825 free energies of association, VII, 810-820

multiple modes of binding, VII, 833835 pH dependence, VII, 832-833 separation of group contributions, VII, 829-832 separation of site contributions, VII, 825-829 thermodynamics of association, VII, 820-822

self-association, VII, 756-757 structure in crystal and solution, VII, 757-760

transfer reactions, VII, 843-846 vertebrate chemical modifications, VII, 780-809 denaturation, VII, 760-780 general considerations, VII, 666-670 physical properties, VII, 717-760 preparation, composition and sequence, VII, 670-682

M

Macromolecular complexes, metabolic regulation and, I, 426-428 Maanesium, enolase and, V, 524526 Malate, cleavage of, VII, 431 Malate dehydrogenase, catalytic properties, active site structure, X I , 390-395 kinetic analyses, XI, 385-390 distribution and preparation, X I , 370373

mitochondrial, environment of, XI, 395-396

molecular properties, amino acid composition, XI, 375376, 377

molecular weight, XI, 373-374 nature of subforms, XI, 376, 378 subunit structure, XI, 374375 structure of NAD'dependent cytoplasmic interaction with coenzyme, X I , 382385

pig heart crystal structure, X I , 379382

types of, XI, 369-370 Malate synthase condensation mechanism, VII, 420-422 occurrence, VII, 412-414 proton transfer and, 11, 316-317 purification and properties, VII, 415 reversibility, VII, 417

511

TOPICAL SUBJECT INDEX

specificity, VII, 415-417 stereochemistry, VII, 417419 Maleate isomerase, properties, VI, 38% 385

Maleic anhydride, pyruvate carboxylase and, VI, 22-23 Maleyl isomerase(s), properties, VI, 385390

Malonyl coenzyme A-acyl carrier protein transacy lase catalytic properties assays, VIII, 179-180 mechanism, VIII, 180-185 pH optimum, substrate specificity and kinetics, VIII, 180 historical background, distribution and metabolic significance, VIII, 176178

molecular properties, VIII, 178-179 Mammals amino acid decarboxylases of, VI, 221224

glutamine synthetase of, X, 699-754 hexokinase mechanism, IX, 4144 occurrence of multiple forms, IX, 31-33

purification and molecular properties, IX, 37-41 regulation, IX, 44-46 relation of solubie to insoluble forms, IX, 33-37 neuraminidase of, V, 325-327 Mandelic acid racemase, properties, VI. 379-380

D-Mannonate dehydrase, properties, V, 579

D-Mannose isomerase, properties, VI, 344-345

L-Mannose isomerase, properties, VI, 345-346

Mannosed-phosphate isomerase catalytic properties general kinetic parameters, VI, 307309 mechanism of action, VI, 310-314 zinc effect on nonemymic isomerization, VI, 309-310 characterization as zinc metalloenzyme, VI, 305-307

history, occurrence and function, VI, 302-304

molecular properties of yeast enzyme, VI, 304-305 Medium chain fatty acid:coenzyme A ligases catalytic properties formation of butyryl adenylate, X, 484

steady state kinetics and reaction mechanism, X, 484-485 substrates and inhibitors, X , 483 Melilotate hydroxylase, properties, XII, 217-221

Metabolic functions regulation of balance allosteric determination of catalytic function, I, 442444 compensatory control mechanisms, I, 434-439

metabolite interconversion systems. I, 43@-434 oppositely directed exergonic reactions, I, 439442 Metabolic regulation adenylate energy charge and, I, 470476

covalently bonded modifiers and, I, 484

operational response curves, I, 482-483 principles chemical equilibria and, I, 418-419 compartmentalization and, I, 423426 covalent enzyme modification, I, 409-415

enzyme concentration, I, 399403 feedback inhibition, I, 403-408 inorganic ions, I, 419-423 macromolecular complexes, I, 426428

proteolysis, I, 416-417 unidirection of reversible reactions, I, 428-430 Metabolic systems, analogy with electronic systems, I, 463-465 Metal ( s ) catalysis by, VI, 401406 kinetics and, 11, 59-61 microbial proteinases and, 111, 772-773

512

TOPICAL SUBJECT INDEX

Metal bridge complexes binary complex formation, 11, 494495 development of concept, 11, 485-489 enzyme-metal linkage, 11, 490494 reactions within coordination sphere, 11, 497-498 ternary complex formation, 11, 495497

Metal complexes reaction chelation mechanisms, 11, 453-455 coordinated ligand reactions, 11, 455463

ligand substitution mechanisms, 11, 450-453

Metal ions properties relevant to catalysis general, 11, 448-450 reactions of metal complexes, 11, 450-463

Metalloenzymes, enzymic properties, 111, 76-77

5,lO-Methenyltetrahydrofolate cyclohydrolase, properties, IX, 201 Methionine adenosyltransferase catalytic properties activators and pH effects, VIII, 133135

assay, VIII, 129-130 energetics, VIII, 139-141 kinetics, VIII, 137-139 reversibility, partial reactions and mechanistic considerations, VIII, 130-133

substrate specificities and inhibition by substrate analogs, VIII, 135-137 net reaction, VIII, 125-127 purification and physical properties, VIII, 127-129 regulation and genetics mammals, VIII, 142-143 microorganisms, VIII, 141-142 significance and distribution, VIII, 123-125

Methionine residues chemical modification, I, 173 chymotrypsinogen, 111, 179 ribonuclease, IV, 682483 subtilisin, modification of, 111, 598-599

Methionyl transfer ribonucleic acid transformylase, properties, IX, 208209

4-Methoxybenzoate O-demethyl-monooxygenase, properties, XII, 285-287 3-Methylaspartate ammonia-lyase catalytic procesa evidence for carbanion mechanism, VII, 127-135 substrate and activator binding, VII, 121-127

distribution, purification and kinetic properties, VII, 118-119 size and constitution, VII, 119-121 p-Methylcrotonyl coenzyme A carboxylase distribution, VI, 4 1 historical background and metabolic significance, VI, 39-40 mechanism of action, VI, 4 2 4 5 molecular characteristics, VI, 42 reaction catalyzed, VI, 38-39 substrate specificity, VI, 4 1 4 2 a-Methyleneglutarak mutase, VI, 534535

assay, VI, 535 catalytic properties coenzymes, W,536 equilibrium, VI, 536-537 mechanism, VI, 537 substrates and inhibitors, VI, 536 purification and molecular properties, IV, 535-536 Methyl groups asymmetrical, stereospecificity of malate synthesis, 11, 157-164 7-Methyl-y-hydroxy-a-ketoglutaric aldolase, properties, VII, 301-302 Methylmalonyl coenzyme A epimerase, stereochemistry, 11, 206-210 Methylmalonyl coenzyme A mutase, VI, 511-512

assays, VI, 51%513 catalytic properties coenzymes, VI, 517-519 equilibrium, VI, 519 mechanism, VI, 519-524 pH and, VI, 519 substrates, VI, 517 distribution, VI, 513

TOPICAL SUBJECT INDEX

purification and molecular properties animal tissue, VI, 515-517 Propionibacterium shermanii, VI, 514-515 Methylmalonyl coenzyme A racemase, properties, VI, 378-379 N5-Methyltetrahydrofolate-homocysteine methyltransferases, see Methyltransferase, Vitamin Bn methyltransferase Methyltransferase assay and purification, IX, 154-155 catalytic properties and folate binding, IX, 156-158 folate substrates, IX, 161-162 occurrence, IX, 160-161 physical properties, IX, 155-156 reactions catalyzed, IX, 121-122 repression of synthesis, IX, 158-160 Mevaldate reductase, stereospecificity, 11, 186-189 Mevalonate labeled, preparation and use, 11, 192204 Mevalonate kinase, stereospecificity, 11, 186-189 Micelles, propinquity effects and, 11, 264-274 Michaelis-Menten mechanism single substrate-single product, 11, 7475 early phase solutions, 11, 77 general solutions to one intermediate mechanism, 11, 75-77 steady state solutions, 11, 77-79 Microbial proteinase(s) chemical properties, 111, 728-730, 742743, 749-754, 770-773 distribution and isolation, 111, 724-728, 741-742, 745-749, 767-770 enzymic properties, 111, 734-740, 742743, 758-763,776-786 inhibitors, 111, 762-763, 777-778 kinetics and mechanism, 111, 763, 778-786 optimum pH, 111, 758-759, 776-777 substrate specificity, 111, 759-762, 778-786 physical properties, 111, 731-733, 742743

513 conformation studies, 111, 755-758, 775-776 molecular weight, physical constants and isoelectric point, 111, 754-755, 773 stability, 111, 755, 773-775 Micrococcus luteus endonuclease, ATPdependent, IV, 261-262 polynucleotide phosphorylase of, VII, 548-570 ultraviolet repair enzymes, IV, 269-270 Micrococcus lysodeikticus adenosine triphosphatase of, X, 421-425 phage-induced lytic enzyme catalytic properties, V, 401-402 purification, V, 401 stability, V, 402 Microorganisms, see also Bacteria elc. aldolases of, VII, 255-256 amylase of, V, 239-271 lipases of, VII, 614-616 3’-nucleotidases of, IV, 354 ribonucleoside 2’,3’-cyclic phosphate diesterase of, IV, 356-363 succinate dehydrogenase of, XIII, 254256 type b cytochromes in, XI, 577-584 Microsomes electron transport, XIII, 148-149 cytochrome b, reductase system, XIII, 150-151 cytochrome P-450 reductase system, XIII, 149-150 mixed function amine oxide, XIII, 153-154 synergism between systems, XIII, 151-153 Milk, lipases, VII, 611-613 Milk xanthine oxidase catalytic properties mechanism of action, XII, 365-388 reactions catalyzed, XII, 344-365 historical background, XII, 303-304 molecular properties chemical modification, XII, 317-326 composition and physical properties, XII, 310-317 magnetic interactions of redox groups, XII, 342-344

514

TOPICAL SUBJECT INDEX

purification, XII, 304-310 redox group magnetic and optical properties, XII, 326-342 Mitochondria adenosine triphosphatase assay, X, 377 beef heart, X, 377-386 rat liver, X, 387-389 yeast, X, 386-387 nicotinamide adenine dinucleotide dehydrogenases, XIII, 177-178 energy conservation and, XIII, 214216

high molecular weight, XIII, 187-189 inhibitors of, XIII, 203-207 low molecular weight, XIII, 189-198 relevance of low and high molecular weight dehydrogenases, XIII, 198203

transhydrogenation and, XIII, 207214

ubiquinone reductase (Complex I), XIII, 178-187 ribonucleic acid polymerase of, X, 318-328

type b cytochromes in, XI, 564-565 Molds, see abo Fungi amylases catalytic properties, V, 263-271 molecular properties, V, 236-263 glycogen synthetase of, IX, 361 proteases acid, 111, 723-744 diisopropylfluorophosphate sensitive, 111, 744-765 metal-chelator-sensitive, 111, 765-786 other, 111. 786-795 Mblybdenum iron-sulfurflavin hydroxylase catalytic properties mechanism of reduction, XII, 394397

oxidizing substrates, XII, 397400 distribution and biological importance, XII, 301-303 enzymes to be considered, XII, 300-301 genetic studies Aspergillus nidulans, XII, 412414 Drosophila melanogaster, XII, 406412

nitrate reductase and, XII; 402406 xanthinuria and gout in man, XII, 400-402

molecular properties, XII, 389-394 other, XII, 388-389 Monoamine oxidase, molecular properties, and kinetic mechanism, XII, 466-471

Monooxygenase (s) copper-containing dopamine, XII, 294-295 phenol, XII, 295-297 flavin and pterin-linked, background, XII, 191-193 heme containing flavoproteins and, XII, 269-280 general mechanisms, XII, 280-285 iron-sulfur proteins as electron donors, XII, 259-269 iron and copper-containing functions, XII, 257-258 historical aspects, XII, 253-255 nomenclature, XII, 256-257 role in oxygen activation, XII, 256 iron-containing, heme, XII, 258-285 nonheme, XII, 285-294 model studies and possible mechanisms, XII, 241-262 nonheme iron-containing lysine, XII, 293-294 4methoxybenzoate 0-demethyl, XII, 285-287 phenylalanine, XII, 287-289 proline, XII, 292-293 tryptophan, XII, 291-292 tyrosine, XII, 290-291 pterin-linked, XII, 231 Multienzyme complexes biological significance, I, 237-240 biosynthesis of aromatic amino acids, I, 228-237 fatty acid synthetases, I, 226-228 a-ketoglutarate dehydrogenase, I, 224225

pyruvate dehydrogenase, composition and organization, I, 215220

regulatory features, I, 220-224

"OhCAL SUBJECT INDEX

Multiple isomorphous replacement how heavy an atom, I, 37-39 location of heavy atoms, I, 32-35 outline of method, I, 32 phase determination and error assessment, I, 35-37 Mung bean, 3'-nucleotidase of, IV, 353 Muscle aldolase, VII, 224-241, 258 creatine kinases of, VIII, 395-401 enolase, glycidol phosphate and, V, 534 fructose-l,6-diphosphatase evidence for presence, IV, 632 physiological role, IV, 634-635 purification and properties, IV, 632633

structure and relation to other enzymes, IV, 633-634 glycogen synthetase of, IX, 332-340 phosphofructokinase of, VIII, 254-256 phosphorylase kinase of, VIII, 557-564 Mutants advantages and limitations of, I, 265266

isolation of, I, 245-249 Mutations alkaline phosphatase and, I, 251-254 amino acid analogs and, I, 262-265 p-galactosidase and, I, 255-256 hemoglobin and,, I, 257-259 other enzymes and proteins, I, 259-262 reversion, suppression and complementation, I, 249-251 tryptophan synthetase, I, 256-257 types, cistron and, I, 243-245 Myelin, ribonucleoside 2',3'-cyclic phosphate diesterase in, IV, 364-365 Myokinase, see Adenylate kinase Myrosulfatase, V, 15-17 Myxobacter, protease of, 111, 786-788 N

Negative charge, proteinase inhibitors and, 111, 422-423 Nervous tissue 5'-nucleotidase of, IV, 346-347 ribonucleoside 2',3'-cyclic phosphate diesterase of, IV, 363-364

intracellular localization, IV, 364-365 physiological role, IV, 365 properties and substrate specificity, IV, 364 Neuraminidase (s) assay method, V, 339-341 biological significance, V, 341-342 historical background, V, 321-323 inhibitors, V, 339 kinetic data, IV, 337-339 occurrence, V, 323 bacterial, V, 324-325 mammalian, V, 325-327 viral, V, 324 properties, 329-331 purification influenza virus, V, 327-328 Vibrio cholerae, V, 328-329 substrate specificity configurational, V, 331-332 esterification of neuraminic acid, V, 336-337

N-substitution of neuraminic acid,

v, 335

0-substitution of N-acetylneuraminic acid, V, 336 position of glycoside linkage, V, 334335

steric hindrance in natural substrates, V, 332-333 Neurospora adenylosuccinase of, VII, 191-193 argininosuccinase of, VII, 180-181 Neurospora crassa acid phosphatase, IV, 497 aspartate transcarbamylase, IX, 302306

argininosuccinase of, VII, 180-181 invertase, V, 303-304 catalytic properties, V, 305 purified preparations, V, 304-305 Neutral proteinase(s) metal-chelator-sensitive, 111, 765-766 chemical properties, 111, 770-773 distribution and isolation, 111,767770

enzymic properties, 111, 776-786 physical properties, 111, 773-776 Nicotinamide adenine dinucleotide, pyruvate carboxylase and, VI, 33-34

516

TOPICAL SUBJECT INDEX

Nicotinamide adenine dinucleotide dehydrogenase Azotobacter vinelandii, XIII, 221 mammalian mitochondria, XIII, 177178

energy conservation and, XIII, 214216

high molecular weight, XIII, 187-189 inhibitors of, XIII, 203-207 low molecular weight, XIII, 189-198 relevance of low and high molecular weight, XIII, 198-203 transhydrogenation and, XIII, 207214

ubiquinone reductase (Complex I), 178-187

yeast, XIII, 216-221 Nicotinamide adenine dinucleotide kinase assay, IX, 77-78 distribution, purification and stability, IX, 76-77 kinetic and molecular properties, IX, 78-79

reaction mechanism, IX, 79-80 substrate specificity, IX, 80-82 Nicotinamide nucleotide transhydrogenase

AB-specific historical, XIII, 62-64 kinetics and reaction mechanism, XIII, 75-78 molecular properties, XIII, 69-71 occurrence, XIII, 64-66 preparation and assay, XIII, 66-69 reconstitution, XIII, 78-79 relationship to energy-coupling system, XIII, 71-75 BB-specific historical, XIII, 52-53 molecular properties, XIII, 57-59 occurrence, XIII, 53-54 purification and assay, XIII, 54-57 reaction mechanism and regulation, XIII, 59-62 definition, XIII, 51-52 physiological roles, XIII, 79-81 fatty acid synthesis, XIII, 88 mitochondrial glutamate and isocitrate metabolism, XIII, 85-88

mitochondrial monooxygenase reactions, XIII, 83-85 redox state of mitochondrial nicotinamide nucleotides, XIII, 81-82 Nitrate reductase, molybdenum hydroxlase “common cofactor,” XII, 402406

Nitrite reductase(s), properties of, XIII, 273-279

Nitrogenase, properties of, XII, 50-56 Nonheme iron proteinb), metal complexes and, 11, 531-533 Nuclear magnetic resonance, ribonuclease, IV, 723-725 Nuclear preparations rat liver, pyrrolidone carboxylate formation by, IV, 138-139 Nucleases, see abo Deoxyribo- and Ribonucleases modification of, I, 130 X-ray diffraction studies, I, 67-69 Nucleophilic catalysis, isomerization and, VI, 397-401 Nucleoside 3’,5’-cyclic phosphate diesterase, IV, 365-366 distribution, IV, 366 inhibitors and activators, IV, 368-370 intracellular localBation, IV, 367-368 metal ions, pH and substrate affinity, IV, 368 physiological function, IV, 370-371 possibility of other diesterases, IV, 370 substrate specificity, IV, 366-367 Nucleoside diphosphokinase(s) assay methods, coupled, VIII, 321-325 isotopic, VIII, 325 staining procedure, VIII, 325 catalytic properties conformational changes, VIII, 331 metal requirements, VIII, 329-330 reaction catalyzed, VIII, 320 specificity, VIII, 320-321 sulfhydryl groups, VIII, 330-331, distribution, VIII, 309-313 function in the cell, VIII, 331-333 historical development, VIII, 307-309 kinetics and catalytic mechanism pH and temperature effects, VIII, 328329

517

TOPICAL SUBJECT INDEX

substrate concentration effect, VIII, 326-328

molecular properties occurrence of isozymes, VIII, 313314

phosphorylated enzyme, VIII, 315320

physical properties, VIII, 315 purification procedures, VIII, 314315

Nucleotide kinases, reaction catalyzed, I X , 49-50 3’-Nucleotidase microorganisms, IV, 354 mung bean, IV, 353 rye grass, IV, 353 wheat seedling, IV, 353-354 5’-Nucleotidase (s) bacterial, IV, 338-340 bull seminal plasma, IV, 342-343 cardiac tissue, IV, 347-348 comparison of, IV, 349-352 Ehrlich ascites cells, IV, 348-349 intestinal, IV, 345 liver, IV, 343-345 nervous tissue, IV, 346-347 other vertebrate tissues, IV, 348 pituitary gland, IV, 346 potatoes, IV, 349 snake venom, IV, 342 yeast, IV, 341342 Nucleotide kinases, reaction catalyzed, I X , 50 Nucleotidyl transferring enzymes, role of metals in mechanism, 11, 502-504 Nucleus ribonucleic acid polymerases animal, X, 262-300 higher plant, X, 311-318 yeast and fungi, X, 300-311

0 Old yellow enzyme, molecular properties and kinetic mechanism, XII, 471-473 Oleate formation from stearate, 11, 179-184 hydration, stereospecificity, 11, 184-186 Oligonucleotides, polynucleotide phosphorylase and, XII, 549-552

Optical rotatory dispersion protein structure and, 11, 381-382, 408 secondary, 11, 382-386 tertiary, 11, 386-391 typical cases, 11, 391-407 ribonuclease, IV, 719-723 Orcinol hydroxylase, properties, XII, 223-224

Ornithine mutase, properties, VI, 559 Oxalacetate formation from phosphoenolpyruvate, general considerations, VI, 117119, 165-168

Oxidation-reduction reactions between metal complexes, 11,529-530 cytochrome c, 11,534-538 mechanistic principles, 11, 530-531 nonheme iron proteins, 11, 531-533 xanthine and aldehyde oxidases, 11, 533-534

within metal complexes mechanistic principles, 11, 525 xficotinamide adenine dinucleotidelinked dehydrogenases, 11, 525-528 vitamin BIZmechanisms, 11, 628-529 Oxidoreductase (8) stereospecificity absolute coenzyme configuration, 11, 144-147

historical background, 11, 134-137 hydrogen transfer and, 11, 137-144 significance of A- and B-side specificity, 11, 154-157 substrates and, 11, 147-151 without hydrogen transfer, 11, 151154

Oxygenases, see Dioxygenases, Monooxygenases Oxygen binding proteins, X-ray diffraction studies, I, 53-63 P

Pancreas pig, phospholipase A, of, V, 76-77 Pancreatic elastase, see Elastase Papain active site chemical modification, 111, 515-516 comparison with other proteinases, 111, 498-499

TOPICAL SUBJECT INDEX

geometry of, 111, 496498 location of thiol group, 111, 514 amino acid composition and sequence, 111, 507-509 assay methods, 111, 518-519 chemical modifications, I, 205-207 active site, 111, 515-516 other reactions, 111, 516-518 crystallization of, 111, 486-488 fructose-1,6-diphosphatase and, IV, 619 heavy atom derivatives, 111, 488-490 kinetic studies acyl enzyme intermediate, 111, 525532 deacylation, 111, 533-635 kinetic constants and pH, 111, 535537 pH, ionic strength and temperature effects, 111, 525 partial reduction, 111, 617 physical properties hydrodynamic properties, 111, 503 immunochemical studies, 111, 606 spectrophotometric and fluorescence properties, 111, 503-505 stability, 111, 505-506 preparation and crystallization, 111, 502-503 specificity esterase and thiolesterase activity, 111, 523-524 nucleophile binding, 111, 524-525 peptide and amide bond hydrolysis, 111, 519-523 transamidation and transesterification, 111, 524 three-dimensional structure description, 111, 491-496 electron density map, 111,490-491 water-insoluble derivatives, 111, 117518 Papaya latex, other proteolytic enzymes, 111, 537-538 Papaya peptidase A, 111,538 Parallelism, protein evolution and, I, 329332 Pasteur effect, phosphofructakinase and, VIII, 274-276 Pea seeds, argininosuccinase of, VII, 181182

Pentacovalency, pseudorotation and, VIII, 214-219 Penicillinase assay methods, IV, 35-39 background, IV, 23-25 catalytic reaction, IV, 27 conformation and function nonspecific transitions, IV, 44-15 specific transitions, IV, 45-46 definitions and specificity, IV, 25-26 factors affecting activity activators and inhibitors, IV, 43-44 pH and temperature, IV, 42-43 immunological studies, IV, 46 kinetics and substrate specificity, IV, 39-40 molecular properties composition and sequence analysis, IV, 31-35 purification and physical properties, IV, 2731 occurrence, IV, 26 structural modification of, IV, 41-42 substrate structural modifications, IV, 40-41 Pepsin action esterase activity, 111, 151 organic sulfite cleavage, 111, 151-152 Specificity, 111, 142-151 theories of, 111, 160-164 amino acids composition, 111, 128-130 sequence, 111, 130-133 assay, 111, 124-125 autolysis of, 111, 139-140 chemical modification, 111, 133-137 condensation reactions, 111, 156 denaturation of, 111, 137-138 electrophoretic mobility, 111, 127 formation from pepsinogen, 111, 138139 gene duplication and, I, 308 historical background, 111, 120 inhibition of, 111, 154-156 isotope effects, 111, 160 molecular weight and shape, 111, 126127 occurrence

519

TOPICAL SUBJECT INDEX

classification and nomenclature, 111, 121-123

other pepsinlike enzymes, 111, 123 optical properties, 111, 127-128 pH dependence, 111, 152-154 protein cleavage by, 111, 140-142 purification of, 111, 124 ribonuclease and, IV, 673 side chain specificity, 111, 145-147 stereochemical specificity, 111, 147-151 transpeptidation reactions, 111, 157-160 Pepsinogen, included with Pepsin Peptide(s) synthesis, nucleic acid independent, VIII, 11-17 Peptide bonds cleavage, proteinase inhibitors and, 111, 452-454 Peptide chain elongation, see also Elongation scheme of, X , 54 Peptide groups, ultraviolet absorption, 11, 379-380 PH chymotrypsin and, 111, 231-233, 236237

p-galactosidase and, VII, 635, 644-645 guanidino kinases and, VIII, 476-477 kinetic parameters and, 11, 52-56 methylmalonyl coenzyme A mutase and, VI, 519 phosphoglucomutase and, VI, 455 pyruvate carboxylase and, VI, 21 Phage, see also Bacteriophage Phage lysozyme, see also Lysozyme application to other biologically important problems, V, 350-352 early history, V, 344-345 recent developments, V, 345-349 Phenol o-monooxygenase, properties of, XII, 221-223, 296-297 Phenylalanine ammonia-lyase catalytic process function of prosthetic group, VII, 159-162

metal ion activation, VII, 162-163 prosthetic group, VII, 154-159 rate limiting step, VII, 164-166 reaction sequence, VII, 148-154

distribution, purification and kinetic properties, VII, 142-146 mechanism of action, VII, 195-196, size and constitution, VII, 146-148 Phenylalanine hydroxylase, properties, XII, 232-238 Phenylalanine 4-monooxygenase, properties, XII, 287-289 Phenylpyruvate synthase, properties, VI, 205-207 Phosphagen kinase, see Guanidino kinase(s) Phosphate, adenosine diphosphoryl glucose pyrophosphorylase and, VIII, 91-92 Phosphate esters hydrolysis of acyclic di- and triesters, VIII, 207-208 metaphosphate mechanism for monoesters, VIII, 202-206 Phosphoenolpyruvate, enzymatic synthesis, X, 631-633 Phosphoenolpyruvate carboxykinase discovery, distribution and physiological role, VI, 136-138 physical properties, VI, 143-148 reactions catalyzed and their properties metal requirement, VI, 141-142 nucleoside phosphate specificity, VI, 140-141

oxalacetate or phosphoenolpyruvate formation, VI, 139 pH optimum, VI, 142-143 pyruvate formation, VI, 139-140 thiol reagents and, VI, 143 regulation activity, VI, 151-154 concentration, VI, 150-151 intracellular location, VI, 148-150 Phosphoenolpyruvate carboxylase discovery, distribution and physiological role, VI, 119-122 kinetic and regulatory properties, VI, 126-133

mechanism of action, VI, 133-136 nature of reaction catalyzed, VI, 122124

role of metals in mechanism, 11, 507508

520

TOPICAL SUBJECT INDEX

structural studies, VI, 124-126 Phosphoenolpyruvate carboxytransphosphorylase characteristics and mechanism of catalyzed reaction, VI, 157-161 different forms, structure and catalytic activities, VI, 163-164 discovery, distribution and physiological role, VI, 154-157 kinetic parameters, effectors and inhibitors, VI, 162 Phosphoenolpyruvate synthetase catalytic properties kinetic studies, X, 643-645 mechanism, X, 638441 metal ion requirements, X, 645 pH and equilibrium, X, 645-646 regulation, X, 648-649 specificity, X, 646-647 stoichiometry, X, 637-638 molecular properties bound divalent metal ion, X, 637 molecular interconversions, X, 635636

phosphoryl and pyrophosphoryl enzyme, X, 636-637 purification, X, 633-634 stability, X, 634-635 sulfhydryl groups, X, 635 Phosphofructokinase assay of, VIII, 243-244 catalytic properties cation requirement, VIII, 247-248 isotopic exchange, VIII, 252 kinetic studies, VIII, 248-252 phosphoryl acceptor specificity, VIII, 244-245

phosphoryl donor specificity, VIII, 24&247

control of glycolysis hormones and, VIII, 277-278 Pasteur effect, VIII, 274-276 pyridine nucleotide oscillations, VIII, 276 purification, VIII, 241-243 reaction catalyzed, VIII, 240-241 regulation of, I, 439-441, VIII, 261-269 role of specific groups histidine, VIII, 272 other functional groups, VIII, 272-274

thiols, VIII, 269-272 structural properties Clostridium pasteurianum, VIII, 256 dilution effects, VIII, 259-260 Escherichia coli, VIII, 255257 isozymes, VIII, 257 molecular weight, VIII, 253-254 phosphorylation, VIII, 260-261 rabbit erythrocyte, VIII, 257-258 rabbit muscle, VIII, 254-256 Phosphoglucomutase activation, VI, 439442 assay, VI, 417-418 all-or-none, VI, 420421 colorimetric, VI, 418419 coupled, VI, 419 radiometric, VI, 419-420 catalytic reaction central complex structural differences, VI, 432 isomeric forms of phosphoenzyme, VI, 430-432 isotope exchange reactions, VI, 429430

kinetics, VI, 426-429 reaction sequence, VI, 421-424 roles of glucose diphosphate, VI, 424426

inhibition anions, VI, 442444 cations, VI, 444-446 chemical modification, VI, 446-447 miscellaneous, VI, 447 poor substrates and analogs, VI, 444 metal ion effects activation, VI, 448-449 addition and release of magnesium, VI, 449-451 addition and release of other metals, VI, 451-452 complexes in vivo, VI, 454 metal-binding site, VI, 453454 structural changes, VI, 452-453 pH and temperature effecta, VI, 455 physical and chemical properties, VI, 413-415

polymorphism, VI, 416-417 purity, VI, 412-413 stability, VI, 416

TOPICAL SUFLJECT INDEX

preparation chromatography, VI, 409410 dephospho-enzyme, VI, 411412 isolation, VI, 408-409 phosphate-labeled, VI, 410-411 specificity, VI, 436-439 structural studies active site phosphate group, VI, 455-456 active site phosphopeptide, VI, 456-457 conformational studies, VI, 457-458 thermodynamics hydrolysiv of phospho-enzyme, VI, 434-436 overall reaction, VI, 433 phosphate transfer to glucose phosphate, VI, 433-434 6-Phosphogluconate dehydrase distribution, V, 573-574 properties, V, 575-578 3-Phosphoglycerate, adenosine diphosphoryl glucose pyrophosphorylase and, VIII, 91-92 3-Phosphoglycerate kinase biological behavior occurrence, VIII, 337-338 reaction catalyzed, VIII, 338-337 species variation and genetics, VIII, 338-340 historical background, VIII, 335-336 molecular properties molecular weight, VIII, 342 primary structure, VIII, 342343 secondary structure, VIII, 344 tertiary structure, VIII, 344-346 purification procedures, VIII, 340-341 reaction kinetics backward and forward reactions, VIII, 346-348 metal ion specificity, VIII, 349 nucleotide specificity, VIII, 348-349 postulated mechanism, VIII, 349-351 Phosphoglycerate mutase assay, VI, 462-464 inorganic and simple organic compounds, effects, VI, 474-475 kinetics, VI, 469-470 pH and temperature effects, VI, 475

521 phosphate transfer to water, VI, 472474 physical and chemical properties, VI, 460-462 preparation and purity, VI, 459-460 reaction sequence, VI, 464-468 Specificity, VI, 471 stability and storage, VI, 462 thermodynamics, VI, 470-471 Phosphoglycolate, triosephosphate isomerase and, VI, 335-336 Phospholipase(s), general, V, 71-73 Phospholipase A,, V, 73 isolation and purification, V, 74-75 Crotalus adamanteus venom, V, 75-76 Crotalus atrox venom, V, 77-78 pig pancreas, V, 76-77 physical and chemical characteristics amino acid content, V, 80-81 amino acid sequence, V, 81-82 molecular weight, V, 78-80 sources, V, 74 substrates and mode of attack, V, 74 Phospholipase C isolation and purification Bacillus cereus, V, 83-84 Clostridium perfringens, V, 84-85 sources, V, 82-83 Phospholipids, glucose-6-phosphatase and, IV, 554-556 Phosphomutase(s), other sugars, VI, 458-459 Phosphoribosylpyrophosphate synthetase catalytic properties conditions affecting activity, X, 617 equilibrium constant, X, 617 mechanism, X, 618-621 substrates and activators, X, 614-617 inhibition by metabolites bacterial enzyme, X, 621-622 mammalian enzyme, X, 622-623 physiological significance, A, 623 occurrence and purification, X, 611-612 other regulatory aspects, X, 623-624 physical and chemical properties, X, 612-614 reaction catalyzed and assay methods, X, 608-611 related enzymes, X, 607-608

TOPICAL SUBJECT INDEX

Phosphorus acyclic, nucleophilic reactions and, VIII, 208-214 Phosphorylase kinase heart muscle, VIII, 564 liver and other mammalian tissues, VIII, 565 nonmammalian sources, VIII, 565 skeletal muscle, VIII, 557-564 0-Phosphorylethanolamine phospholyase, properties, VII, 5 2 5 3 Phosphoryl transfer catalysis intramolecular, VIII, 219-227 metal ion, VIII, 227-231 enzymic mechanisms, VIII, 232-233 bimolecular or aeaociative, VIII, 235-238

metaphosphate, VIII, 233-235 Phosphotransferase, glucose-bphosphatase and, IV, 592-595 Phosvitin kinase, properties, VIII, 5& 581

Photoisomerization, related problems and, VI, 395-397 Photosynthesis bacterial, XI, 509-510 green sulfur, XI, 514-516 purple nonsulfur, XI, 610-512 purple sulfur, XI, 512-514 reductive carboxylic acid cycle of, VI, 207-213 Physarum polycephalum, ribonuclease PPI of, IV, 241 Physiological function design requirements, I, 465-467, 484488

adenylate energy charge, I, 470-476 interactions between input signals, I, 476-484 kinetic properties, I, 467470 Pituitary gland, 5’-nncleotidase of, IV, 346 Plants acid phosphatase of, IV, 497 adenosine diphosphoryl glucose pyrophosphorylase algae, VIII, 90 leaves, VIII, 86-90 nonchlorophyllous tissue, VIII, 93-

94

alcohol dehydrogenase of, XI, 188-189 amino acid decarboxylases of, VI, 221-224

aspartate transcarbamylase of, IX, 307-308

cytochromes b, microsomes, XI, 591 mitochondria, XI, 589-591 photosynthetic systems, XI, 587-589 fructose-l,6diphosphatase

physiological role, IV, 642-643 purification and properties, IV, 640-642

regulation, IV, 643 isoamylases of, V, 208 nuclear ribonucleic acid polymerase, X, 311-318 pullulanase of, V, 202-204 viral ribonucleic acids, terminal sequences, X , 85-86 Pneumococci, endonucleases, IV, 260-261 Point mutations, protein structure and, I, 286-292 Polymerization statistics of, X , 157-158 oligomer synthesis, X, 159-180 polymer synthesis, X, 158-159 Polynucleotide ( 8 ) enzymic methylation, general considerations, IX, 167-168 polynucleotide phosphorylase and, VII, 552-557 staphylococcal nuclease and kinetic measurements, IV, 186-187 specificity, IV, 185-186 Polynucleotide phosphorylase catalytic reactions, VII, 545-546 Clostridium and mutant Escherichia enzymes, VII, 571-572 enzyme-polynucleotide complex, VII, 570 exchange reactions, VII, 570-571 other practical uses, VII, 572-574 phosphorolysis, VII, 548-557 polymerization, VII, 557-570 specificity, VII, 546-548 degradation of, VII, 540-542 general background, VII, 533-535 in vivo

TOPICAL SUBJECT INDEX

control of synthesis and activity, VII, 538-539 distribution and localization, VII, 536-538

physiological role, VII, 538 molecular weight and subunits, VII, 543-545

polymerization by kinetics, VII, 557-566 mechanism, VII, 566-570 purification, VII, 539-540 thermal stability, VII, 542-543 Polypeptide chain initiation eukaryotes inhibitors, X, 43 initiation factors, X, 29-43 initiator aminoacyl-transfer ribonucleic acid and, X, 28-29 messenger ribonucleic acid transla: tion, X, 43-44 prokaryotes general, X, 2-5 initiation factors, X,6-28 initiator aminoacyl-transfer ribonucleic acid and, X, 5-6 regulation interference factors, X, 44-45 messenger recognition, X, 46-51 Polypeptide chain termination events of protein synthesis and, X, 114-117

mechanism interaction of release factors with ribosomes, X, 101-108 in vitro assay, X, 100-101 peptidyl-transfer ribonucleic acid hydrolysis, X, 108-114 role of guanine nucleotides, X, 106108

requirements soluble protein factors, X, 95-100 terminator codons, X, 88-95 Polyprenyl biosynthesis stereospecificity 3-hydroxy3-methylglutaryl coenzyme A synthesis, 11, 186-189 preparation and use of labeled mevalonates, 11, 192-204 squalene biosynthesis, 11, 190-192

Polysaccharide depolymerase Aerobacter phages, V, 398 Azotobacter phage, V, 397-398 bacteriophage F series catalytic properties and biological significance, V, 393 enzyme assay, V, 392 partial purification, V, 392-393 stability, V, 393 Klebsiella phage enzyme properties and role, V, 394-395

preparation and assay, V, 394 Pseudomanas phages, V, 395-397 Porphobilinogen synthesis, mechanism of, VII, 333-337 Potatoes, 5’-nucleotidase of, IV, 349 Procarboxypeptidase B, physical and chemical properties, 111, 67-68 Proelastase, activation of, 111, 331-332 Prokaryotes polypeptide chain initiation general, X, 2-5 initiation factors, X, 6-28 initiator aminoacyl-transfer ribonucleic acid and, X, 5-6 Proline iminopeptidase, 111, 115 Proline 4-monooxygenase, properties, XII, 292-293 Proline reductase, Schiff base and, 11, 358

Proline residues, proteinase inhibitors and, 111, 423 Prolyl hydroxylase, XII, 152-154 assays, XII, 161 catalytic properties, XII, 156-160 nonvertebrate, XII, 163-165 purification and molecular properties, XII, 154-156 regulation, XII, 161-163 Propane-l,2-diol dehydrase, stereochemistry, 11, 210-214 Propinquity effects activation parameters and kinetic order, 11,250-254 approximation through noncovalent forces charge-transfer, 11, 254-264 Debye, 11, 254-264 hydrogen bonds, 11, 254-264

524 inclusion compounds, 11, 274-279 London, 11, 254-264 micelles, 11, 264-274 evaluation of concepts, 11, 220-226 Propionyl coenzyme A carboxylase distribution, VI, 48 historical background and metabolic significance, VI, 46-48 mechanism of action, VI, 51-53 molecular characteristics, biotin binding site, VI, 50-51 reaction catalyzed, VI, 46 stereochemistry, 11, 208-210 substrate specificity, VI, 49 a-(n-Propyl)malate, synthesis, VII, 426-427 Prostate gland acid phosphatase assay, IV, 457 electrophoresis, IV, 468-469 functional groups, IV, 469-472 general, IV, 455-457 kinetics, IV, 457466 physical properties, IV, 476 preparation, IV, 466-468 transphosphorylation, IV, 472-473 use as a reagant, IV, 473-476 Protease(s), see also .Proteinases activation, proteolysis and, I, 416 gene duplication and pancreatic, I, 303-307 dfhydryl, I, 307-308 hexokinases and, IX, 2-7 modification of, I, 118-124 serine, sequence homologies, 111, 343 352 X-ray diffraction studies, 1, 63-67 Protein (8) allosteric alternative approaches, I, 385-388 evaluation of, I, 372-375 evolutionary considerations, I, 390393 hemoglobin, I, 388-390 molecular basis of cooperativity, I, 375-379 protein design, I, 379-381 status of simple models, I, 381-385 chain termination and eukaryotic, X, 99-100

TOPICAL SUBJECT INDEX

prokaryotic, X, 95-99 cleavage by pepsin, 111, 140-142 functional group adenylylation, VIII, 40-49 genetic phenomena in chain shortening, I, 292-293 deletion, addition, chain extension, I, 293-300 gene duplication, I, 3W314 point mutations, I, 286-292 homologous, speciation of, I, 321-328 structure, common characteristics, I, 87-89 structure-function relationships, evolution and, I, 267-274 Proteinase(s), see also Protease(s) acid, see Acid proteinases alkaline, see also Alkaline proteinases bacterial, 111, 605-606 inhibitor association with, 111, 428-433 inhibitor complexes, dissociation of, 111, 434-436 microbial, see Microbial proteinases neutral, see Neutral proteinases thiol, microorganisms and, 111, 791-795 Proteinase inhibitors assays active site titrants, 111,393-396 competitive enzyme assays, 111, 396 errors in, 111, 396-400 association constants enzyme titration methods, 111, 402403 physicochemical methods, 111,400402 potentiometric method, 111, 403-406 competitive inhibition and, 111, 391393 complex formation, changes in conformation-sensitive parameters, 111, 406-410 crystalline inhibitors and complexes, 111, 383-384 definition, 111, 378 disulfide loop, reactive site and, 111, 422-423 enzyme-susceptible bond in reactive site, 111, 419 historical background, 111, 376-378

TOPICAL SUBJECT INDEX

identities and nomenclature, 111, 378379

kallikrein inhibitor and pancreatic inhibitors, 111, 379-380 serum inhibitors, 111, 382 soybean inhibitors, 111, 380-382 inhibition a t same reactive site, 111, 470-473

molecular differences analogies and homologies, 111,457463

specificity toward other enzymes,

111, 463473 nonoverlapping reactive sites, 111, 466-468

overlapping independent reactive sites, 111, 468470 physical properties, 111, 384-388 reactive site model chemical model of inhibitor, 111, 443-447

control dissociation of complex, 111, 437439

detection of reactive sites, 111, 412-418

equilibria of hydrolysis, 111, 423428 general properties, 111, 418-423 kinetics of interaction, 111,428437 nature of stable complex, 111, 450451

objections to, 111, 451457 overshoot of complex, 111,410-412 residue replacement a t reactive site,

111, 439-441 sites for other enzymes, 111, 441443 temporary inhibition, 111, 447-449 special purification techniques, 111, 389-391

N-terminal residue in reactive site,

111, 419-420 virgin and modified, interconversion,

111, 436437 Protein kinase(s) cyclic nucleotide-regulated, VIII, 566-578

historical background, VIII, 555-557 nonclassified, VIII, 578379 acidic nuclear protein kinases, VIII, 580

histone kinases, VIII, 579-580

phosvitin kinases, VIII, 580-581 substrate-specific phosphorylase kinase, VIII, 557-565 pyruvic dehydrogenase kinase, VIII, 565-566

Pro teolysis metabolic regulation and blood coagulation, I, 416417 proteolytic enzyme activation, I, 416 polynucleotide phosphorylase and,

VII, 557 Prothrombin activating enzyme, 111, 315-317 amino acid and carbohydrate composition, 111,313 aasays of, 111, 308 disulfide bridges, 111, 313 isolation and purity, 111, 308-311 metabolism biosynthesis, 111, 320-321 turnover rate, 111,320 number of polypeptide chains, 111, 314-315

physical properties, 111, 311412 structural aspects models, 111, 318-319 proteolysis, 111, 317-319 secondary proteolysis, 111, 319 N- and C-terminal analysis, 111,313314

Proton dissociation-replacement reactions, 11, 312-313 citrate synthase, 11, 316-317 hydrogenase, 11, 317-318 malate synthase, 11, 316-317 stereochemistry, 11, 313-315 1,l-Proton shifts, epimerases and, 11, 295-298

l,2-Proton shifts, aldo-keto isomerases,

11, 290-295 l,3-Proton shifts, allylases and, 11,299302

Protozoa, glycogen synthetase of, IX, 359 Pseudomonads aspartokinases of, VIII, 551-552 isoamylase of, V, 204-206 Pseudomonus, proteinases, of, 111, 769770

Pseudomonas aeruginosa, phage polysaccharide depolymerase, V, 395-397

TOPICAL SUBJECT INDEX

Pseudomonas putida, phage polysaccharide depolymerase, V, 397 Pseudomonas testosteroni As-3-ketosteroid isomerase catalytic properties, VI, 5 9 W mechanism, VI, 605-615 molecular properties, VI, 592699 Pseudouridine kinase, properties, IX, 62 Pullulanase Aerobacter aerogenes preparation and physical properties, V, 195-197 reversion reactions, V, 201 substrate specificity and action pattern, V, 197-201 plant, V, 202204 Purine aminohydrolase (s) assay methods, IV, 51 distribution, IV, 49-51 historical background, IV, 48 Purine nucleoside phosphorylase assays direct spectrophotometry, VII, 505 inorganic orthophosphate estimation, VII, 504 isotopic assays, VII, 505 pentose estimation, VII, 503-504 spectrophotometry coupled with xanthine oxidase, VII, 504-505 catalytic mechanism equilibrium studies, VII, 511 reaction mechanism, VII, 512 reaction sequence, VII, 511-512 distribution in nature, VII, 485-490 historical development, VII, 483485 kinetics substrate concentration and, VII, 505-510

temperature and pH effects, VII, 510-511

metabolic functions bacterial metabolism, VII, 494 chemotherapy, VII, 493-494 erythrocyte metabolism, VII, 4 9 2 493

fish skin, VII, 494495 nucleoside metabolism, VII, 490492 properties, VII, 495-496 purification, VII, 495 reactions catalyzed, VII, 496-500

specificity carbohydrates, VII, 502-503 purines, VII, 500-502 subunit structure, VII, 514 sulfhydryl groups, VII, 513-514 Pyridine nucleotide ( s ) deoxythymidine diphosphate-D-glucose oxidoreductase and, V, 474478 phosphofructokinase and, VIII, 276 Pyridine nucleotide-disulfide oxidoreductases mechanism, similarities and contrasts, XIII, 94-99 reaction catalyzed-chemical similarities and crom-reactivity, XIII, 92-94 structure, similarities and contrasts, XIII, 99-105 Pyridoxal, reactions with amino acids, 11, 339-345 Pyridoxal-linked reactions absorption spectra and, VII, 62-65 amino acids and, VII, 33-39 mechanisms, VII, 65-66 @-elimination and, VII, 66-72 7-elimination and replacement, VII, 72-73

stereochemistry, VII, 73-74 Pyridoxal phosphate, fructose-1,6diphosphatase and, IV, 620 Pyridoxal phosphate enzymes apoenzymes and, 11, 346-348 classification, 11, 367-368 comparative characteristics, 11, 363-364 mechanism of reaction, 11, 349-356 peptide sequences, 11, 366 structural and spectral properties, 11, 348-349

Pyrimidines, dioxygenase reactions of, XII, 169-179 Pyrimidine deoxyribonucleoside 2'-hydroxylase, catalytic properties, XII, 176178

Pyrimidine nucleoside monophosphokinases, IX, 87-88 Pyrocatechase structure, physical probes, 11, 406-407 Pyrophosphokinase(s), other, X, 624-628 Pyrrolidone carboxylate derivatives, enzymic formation of, IV, 146147

TOPICAL SUBJECT INDEX detection and determination, IV, 125127

enzymic formation from glutamate D-glUtaInate cyclotransferase, IV, 133-136

L-glutamate cyclotransferase, IV, 138 glutamine synthetase, IV, 136-137 yglutamylcysteine synthetase, IV, 136-137

rat liver nuclear preparations, IV, 138-139

enzymic formation from glutamine and glutaminyl peptides L-glutamine cyclotransferase, IV, 139-141

y-ghtamyl cyclotransferase, IV, 141 y-glutamyl transpeptidase, IV, 141 enzymic formation from y-glutamyl amino acids y-L-glutamyl cyclotransferase and, IV, 142-146 historical background, IV, 124-125 metabolism, IV, 149-151 natural occurrence, IV, 127-130 nonenzymic formation, IV, 130-133 Pyrrolidone carboxylyl peptidase, IV, 147-149

Pyrrolidonyl peptidase, 111, 113-114 Pyruvate carboxylase acyl coenzyme A derivatives and general properties, VI, 24-27 parameters reflecting enzyme conformation, VI, 29-31 specificity of activation, VI, 27-29 aspartate and, VI, 31-33 first partial reaction, VI, 9-10 immunochemical studies, VI, 23 general properties, VI, 2-3 generalized minimal mechanism, VI, 6-7

monovalent cation effects, VI, 6 partial reactions, VI, 3-5 presence of bound biotin, VI, 3 requirements, VI, 5-6 historical background, VI, 1-2 mild denaturation and chemical modification maleic anhydride, VI, 22-23 other reagents, VI, 23 sulfhydryl reagents, VI, 21-22

temperature, VI, 19-20 urea and pH, VI, 20-21 molecular parameters and quaternary structure, VI, 16-18 nicotinamide adenine dinucleotide and, VI, 33-34 phosphoenolpyruvate and, VI, 34 product inhibition-two-site mechanism, VI, 7-8 regulation of synthesis, VI, 34 role of metals, 11, 511-515 second partial reaction nature of bound metal ion, VI, 10-11 role of bound metal ion, VI, 11-15 Pyruvate dehydrogenase complexes composition and organization, I, 215220

regulatory features, I, 220-224 Pyruvate kinase assay, VIII, 371 catalytic mechanism, VIII, 379-382 control, VIII, 378-379 historical background, VIII, 353-355 kinetics inhibitors, VIII, 375-377 substrates and activators, VIII, 372-375

molecular properties chemical modification, VIII, 360-361 composition, VIII, 358 ronformationnl change, VIII, 361364

purification, VIII, 355-358 structure, VIII, 358-359 muscle, role of metal in mechanism, 11, 504-506 stoichiometry and specificity cofactors, VIII, 366-370 number of active sites, VIII, 370371

reaction catalyzed, VIII, 364 substrate specificity, VIII, 364-366 thermodynamics, VIII, 371-372 yeast, role of metals in mechanism, 11, 506-507

Pyruvate, phosphate dikinase catalytic properties kinetic studies, X, 643, 644-645 mechanism, X, 641-642 metal ion requirements, X, 645

TOPICAL SUBJECT INDEX

regulation, X, 648-649 specificity, X,646 stoichiometry, X,638 molecular interconversions, X, 635636 phosphoryl and pyrophosphoryl enzyme, X, 636-637 purification, X, 633-634 stability, X, 634 sulfhydryl groups, X, 635 Pyruvate synthase, properties, VI, 197201 Pyruvic dehydrogenase kinase, properties, VIII, 565-566

Q Qj3 replicase, elongation factors and, X,

83-85

R Rana cateebiana collagenase, 111, 689-690 catalytic properties, 111, 691-693 preparation, 111, 69M91 Rate equations derivation chemical reaction, 11, 61-63 isotopic exchange, 11, 63-65 Reactions, reversible, unidirection of, 428-430 Red cell acid phosphatase general properties, IV, 477 purification and separation of genetic types, IV, 477-484 Reduced nicotinamide adenine dinucleotide kinase, properties, IX, 82 Reduviin, thrombin and, 111, 304-305 Relaxation amplitudes kinetic studies near equilibrium calculation of amplitudes, 11, 10510s thermodynamic effects of chemical reactions, 11, 101-105 transformation to normal concentration variables, 11, 99-101 Relaxation spectra kinetic studies near equilibrium

alternative treatment of multistep mechanisms, 11, 91-93 analysis and interpretation, 11,95-99 multistep mechanisms, 11, 89-91 one-step mechanisms, 11, 83-84 thermodynamically dependent reactions, 11, 93-95 transient and stationary solutions of rate equations, 11, 84-87 two-step mechanisms, 11, 87-89 Rennin, gene duplication and, I, 308 R-enzyme, see Pullulanase j3-Replacement reactions, pyridoxallinked, VII, 54-57 Retinal isomerase, pigment regeneration and, VI, 587-589 Reverse transcriptase biological role noninfectious murine sarcoma virus, X, 232 other inhibitors, X, 233 rifamycins, X, 233 Rous sarcoma virus a, X, 231-232 temperature sensitive Rous sarcoma virus, X, 232-233 comparison to other polymerases, X, 233-235 inhibitors rifamycins, X, 231 sulfhydryl reagents, X, 230-231 nuclease activity deoxyribonuclease, X, 220-221 ribonuclease, X,221 ribonuclease H,X, 221-222 primer and direction of synthesis, X, 225-226 problems, X, 229-230 properties, X, 218 sire, X, 219-220 storage and stability, X, 219 properties of catalytic reaction deoxyribonucleoside triphosphate, X,224 divalent cations, X, 224-225 other conditions, X, 225 purification, X, 216-218 serological relationships avian leukosis viruses, X, 223 general considerations, X,222

TOPICAL SUBJECT INDEX

mammalian C-type viruses, X, 223224 other viruses, X,224 solubilization, X, 215 template fidelity of synthesis, X, 228-229 preferences, X,226-228 requirements, X, 226 size, X,229 virus purification and, X, 214-215 Reversions, mutations and, I, 249-251 L-Rhamnose isomerase, properties, VI, 345-346 L-Rhamnulose 1-phosphate aldolaae catalytic reaction assay, VII, 308-309 equilibrium constant, VII, 310-311 metal ions and, VII, 309 pH optimum, VII, 309 substrate binding and reaction sequence, VII, 311-313 turnover number, VII, 309 historical background, VII, 304 metabolic significance, VII, 305 molecular properties isolation, VII, 305 physical properties, VII, 306 structure, VII, 306-308 occurrence, VII, 304305 Rhodopseudomonas capsulatus, aspartokinase of, VIII, 544-545 R hodopseudomonua spheroides 8-aminolevulinate synthetase of, VII, 344-345 aspartokinase of, VIII, 552-553 membrane adenosine triphosphatase,

x,429

Rhodospirillum rubrum, adenosine diphosphoryl glucose pyrophosphorylase of, VIII, 81-86 Riboflavin kinase, properties, IX, 74-75 Ribonuclease (s) aggregation of, IV, 744-746 assays, IV, 747-750 rhain conformation and solvent-induced changes, IV, 725-726 added electrolytes, IV, 735-737 organic solvents, IV, 733-735 thermal and acid transitions, IV,

726-731

thermodynamics, IV, 740-744 transitions in derivatives, IV, 738740 urea and, IV, 731-733 chemical modification of functional groups amino groups, IV, 677-682 arginine, IV, 689-690 carboxyl groups, IV, 675-677 cystine-disulfide groups, IV, 690-696 histidine, IV, 685-689 intramolecular crosslinks, IV, 696697 methionine, IV, 682-683 other reagents, IV, 697 chemical modification of functional groups serine and threonine, IV, 696 tyrosine, IV, 684-685 chemical synthesis and S-peptide summary, IV, 697-705 classification of, IV, 205-207 discussion of mechanism and stability lysine 41 and, IV, 801 opposite vs. adjacent attack, IV, 791-794 pH and, IV, 801-806 proton transfer and rate-limiting step, IV, 795-796 role of oxygen, IV, 79-01 stabilization of intermediates, IV, 794-795 structure, IV, 785-788 transphosphorylation and hydrolysis, IV, 788-791 enzymic cleavage of main chain chymotrypsin, IV, 674 elastase, IV, 672-673 pepsin, IV, 673 subtilisin, IV, 669-672 trypsin, IV, 673-674 fungal, general survey, IV, 208-211 historical background, IV, 647-649 isolation and chromatography, IV, 649-653 macromolecular inhibitors, IV, 758-759 mechanism of catalysis Mathias and Rabin et al., IV, 780781 Roberts et al., IV, 784

TOPICAL SUBJECT INDEX

Usher, IV, 783-784 Wang, IV, 782-783 Witzel, IV, 781-782 microbial, of interest, IV, 239-243 list of, IV, 243-249 physical parameters diffusion coefficient, IV, 708-709 electrophoretic mobility, IV, 710-711 fluorescence, IV, 718-719 hydration and axial ratio, IV, 709710

hydrogen ion equilibrium, IV, 711712

hydrogen exchange, IV, 712-714 molecular weight, IV, 709 physical parameters nuclear magnetic resonance and electron paramagnetic resonance, IV, 723-725 optical rotatory dispersion and circular dichroism, IV, 719-723 partial specific volume, IV, 705-707 radius of gyration, IV, 707-708 refractive index increment, IV, 707 sedimentation behavior, IV, 709 ultraviolet absorption spectra, IV, 714-717

viscosity, IV, 710 reaction catalyzed, IV, 746-747 small molecule effectors, IV, 759-772 specificity, IV, 750-751 base, IV, 754-758 phosphate, IV, 758 sugar, IV, 752-754 steady state kinetic data ionic strength and, IV, 777-778 Michaelis constants and turnover number, IV, 772-777 organic solvents and, IV, 779-780 structure amino acid sequence, IV, 653-654 physical probes, 11, 396-401 three-dimensional, IV, 654-669 Ribonuclease N,, IV, 230-231 applications, IV, 232-234 preparation, IV, 231 properties, IV, 231-232 R.ibonuclease T, applications, IV, 222-223

preparation, IV, 212-213 properties, IV, 213-214 specificity and mode of action, IV, 215-218

structure and function, IV, 218-222 Ribonuclease Tz applications, IV, 229-230 preparation, IV, 223-224 properties, IV, 224-225 specificity and mode of action, IV, 225-229

Ribonuclease Uz applications, IV, 237-239 preparation, IV, 234-235 properties, IV, 235 specificity, IV, 235-237 Ribonucleic acid synthesis, VIII, 20-21 polynucleotide adenylyltransferases, VIII, 24-26 polynucleotide phosphorylase, VIII, 23-24

ribonucleic acid polymerase, VIII, 21-23

Ribonucleic acid polymerase(s) animal general properties, X, 280-283 historical, X, 262-264 inhibitors, X, 295-299 intracellular localization, X, 279-280 nomenclature, X, 264-266 physiological role, X, 299-300 purification, X, 266-269 regulation in vivo, X, 300 stirnulatory factors, X, 293-294 structure, X, 269-279 template specificity, X, 284-293 bacterial assay, X, 338-339 chain elongation, X, 359-366 chain initiation, X, 353-359 chain termination, X, 366-370 inhibitors, X, 370-374 outline of reaction, X, 346-348 purification, X, 335-338 template binding, X, 348-353 variety of reactions catalyzed, X, 344-346

chain elongation kinetics, X, 364-366

TOPICAL SUBJECT INDEX

nondissociable ternary complex, X, 359-361 specificity, X, 361-363 chloroplast, X, 329-330 covalent modification diphosphopyridine nucleotidedependent, VIII, 48-49 possible artifact, VIII, 49 inhibitors, X, 370 agents affecting template, X, 373374 agents interacting with enzyme, X, 371-373 fungal, X, 310-311 higher plant enzyme properties, X, 314-316 factors, X, 317-318 function, X, 318 molecular properties, X, 314 solubilization and purification, X, 311-314 mi tochondrial enzyme properties, X, 325-328 molecular properties, X, 324-325 solubilization and purification, X, 318-324 synthesis, X, 328 nuclear animal, X, 262300 higher plant, X, 311-318 yeast and fungi, X , 300-311 structure of bacterial enzyme dissociation and reconstitution, X, 342-343 missing subunit problem, X, 343-344 molecular weight, X, 341-342 subunits and, X, 340-341 yeast, X, 300-301 enzyme properties, X, 306-309 function, X, 310 molecular properties, X, 304-306 solubilization and purification, X, 301-304 stimulatory factors, 309310 Ribonucleoside 2’,3’-cyclic phosphate diesterase nervous ‘tissue, IV, 363-364 intracellular localization, IV, 364-356 physiological rale, IV, 365 properties and substrate specificity, IV, 364

x,

531 Ribonuceloside 2’,3’-cyclic phosphate diesterase with 3’-nucleotidase activity cellular localization, IV, 361-362 a metalloenzyme, IV, 362-363 physiological function, IV, 363 properties kinetics and mechanism of action, IV, 358-361 physical and chemical, IV, 358 substrate specificity, IV, 357458 Ribonucleotides reduction, regulation of, I, 442-443 Ribose-5-phosphate isomerase general, VI, 318-320 catalytic properties assay methods, VI, 321-322 mechanism, VI, 323-324 Michaelis constants, VI, 322-323 molecular properties, VI, 320-321 Ribosomal ribonucleic acid methyltransferase biological significance, IX, 189-190 isoIation and properties, IX, 187-189 occurrence, IX, 187 Ribosome(s) elongation and, X, 67, 77-78 sites involved, X, 09-77 structure and, X, 68-69 elongation factors and, X, 62-03, 66-67 release factor interaction bacterial, X , 101-103 mammalian, X, 103-106 sites involved in elongation guanosine triphosphatase and factor binding sites, X, 73-76 peptidyltransferase center, X, 69-72 role of 30 S proteins, X, 76-77 Ribulose-1,d-diphosphate carboxylase general considerations, VI, 169-173 kinetics and specificity, VI, 181-183 “carbon dioxide,” VI, 183-184 divalent metal ion, VI, 185-186 inhibitors, VI, 186-187 ribulose diphosphate, VI, 184-185 light activating factors, VI, 191-192 mechanistic considerations, VI, 187-191 molecular properties composite quaternary structure, VI, 179-180

532

TOPICAL SUBJECT INDEX

native enzyme, VI, 173-178 subunits, VI, 178-179 reaction, VI, 180-181 o-Ribulose-5-phosphate 3’-epimerase, properties, VI, 374375 ~Ribulose-5-phosphate 4’-epimerase, properties, VI, 372-373 Rubredoxin, chemical properties, XII, 12-15 historical background, XII, 4-6 physical properties, XII, 6-12 Rye grass, 3’-nucleotidase of, IV, 353

s Saccharomyces, see also Yeast acid phosphatase, IV, 497 Saccharomyces cerevisiae aspartate transcarbamylase of, IX, 302-306

aspartokinase of, VIII, 553 fructose-l$-diphosphatase, regulation, IV, 640 Salicylate hydroxylase, properties, XII, 206211

Salmonella phage lytic enzyme, V, 398 Sarcoplasmic membrane(s) calcium binding by, X, 450451 calciumdependent adenosine triphosphatase ion requirements, X, 446-447 lipid depletion and, X, 449 membrane permeability and, X, 445446

temperature and pH, X, 447 thiol reagents and, X, 449-450 calcium-independent adenosine triphosphate, X, 444445 calcium transport by absence of precipitating anions, X, 451 adenosine triphosphate extra splitting, X, 453-455 presence of precipitating anions, X, 451453

release from preloaded vesicles, X, 455456

composition lipids, X, 442 proteins, X, 440-442 in situ, X, 434-435

iso1ated contaminants, X, 437 procedures, X, 435-437 shape and size of vesicles, X, 437-439 lipid functions enzyme properties, X, 465467 membrane permeability, X, 465 lipoprotein structure, physical properties, X, 442-444 Saturation curves, fitting of, I, 361365 Scatchard plots, enzyme regulation and, I, 359-361 Schiff base(s) formation general characteristics, 11, 336-339 nonenzymic catalytic effects, 11, 339-346

other than pyridoxal, TI, 345-346 Schiff base enzyme(s) noncarbonyl, 11, 358-359 aldolases and transaldolases, 11, 359-360

8-aminolevulinate dehydratase, 11, 361-362

comparative properties, 11, 369 p-keto-acid decarboxylases, 11, 359 Seminal plasma bull, 5’-nucleotidase of, IV, 342-343 Serine hydroxymethyltransferase, properties, IX, 215-221 Serine peptides, microbial proteinases and, 111, 751-752 Serine residues, chemical modification, I, 173 chymotrypsin amide hydrolysis and, 111, 224231 ester hydrolysis and, 111, 218-224 ribonuclease, IV, 696 subtilisin, 111, 575-576 conversion to cysteine, 111,577-580 sequence in other proteinases, 111, 576-577

Serratia marcescens, adenosine diphosphorylglucose pyrophosphorylase of, VIII, 107-108 Serum, acid phosphatase in, IV, 495-496 Slime molds, fructose-l,6-diphosphatases of, IV, 640 Solvation, physical organic models, 11, 226-238

TOPICAL SUBJECT INDEX

Sorangium, proteinases of, 111, 747, 752-754, 789-790

Spinach adenosine diphosphoryl glucose pyrophosphorylase of, VIII, 86-89 chloroplast adenosine triphosphatase assay, X , 389 catalytic properties, X, 391393 cold inactivation, X, 390-391 molecular properties, X, 390, 391 nucleotide binding, X, 39-94 purification, X, 389-390 Spleen acid deoxyribonuclease catalytic properties, IV, 276-285 distribution, intracellular localization and biological role, IV, 285-287

physical and chemical properties, IV, 272276

acid exonuclease, IV, 330-336 acid phosphatase, IV, 493-495 deoxyribonuclease of, IV, 272-287 glycogen synthetase of, IX, 354 Squalene, epoxidase and, VII, 211 Staphylococcal nuclease active site, stereochemical probes,

IV, 195-196 behavior in solution, IV, 183-184 covalent structure, IV, 180-183 crystallographic studies, introduction,

IV, 156-159 fluorescence spectroscopy, 11, 429 fragments, complementation of, IV, 196-199

general, IV, 153-156 historical background, IV, 177-178 isolation, IV, 178-179 mechanism, IV, 174-175 peptide chain conformation, IV, 159-163

proposed future studies, IV, 175 studies in solution, IV, 17%174 substrate specificity and catalytic mechanisms polynucleotide substrates, IV, 185-187 size and specificity of active site,

IV, 191-195 synthetic substrates and inhibitors,

IV, 187-191

533 synthetic analogs and, IV, 199-204 synthetic substrates and inhibitors kinetic measurements, IV, 190-195 specificity, IV, 187-189 thymidine-3',5'-diphosphate and calcium ion binding, IV, 163-171 ultraviolet difference spectroscopy, 11, 413-414

a warning, IV, 174 Staphylococci, lytic phage enzymes, V, 398-399

Staphylococcus, acid phosphatase of, IV, 498

Staphylococcus aurews phage lytic enzyme, V, 400-401 virolysin, V, 399-400 Starch structure determination, debranching enzymes and, V, 228-234 Stearate, conversion to oleate, 11, 179-184 Stereochemistry a,j3 elimination reactions, 11, 309-312 proton transfer, 11, 313-315 Stereospecificity, nicotinamide adenine dinucleotide-dependent oxidoreductases, 11, 134-157 Steroid sulfatase(s), V, 4-6 androstenolone sulfatase, V, 7-9 cortisone sulfatase, V, 10 estrone sulfatase, V, 6-7 etiocholanolone sulfatase, V, 9-10 Strain, physical organic models, 11, 226-238

Streptococcal proteinase activation, 111, 627-628 amino acids active site, 111, 626-627 composition, 111, 624-625 N- and C-terminal, 111, 625-626 assay methods, 111, 627 immunological properties, 111, 615-619 kinetics of hydrolysis dielectric constant and, 111, 636-637 esterase vs. peptidase activity, 111, 638-639

p H and, 111, 633-636 structural inhibitors, 111, 637-638 mechanism of action, 111, 639-647 physical properties electrophoresis, 111, 614

534 gel filtration and chromatography, 111, 613-614 molecular weight, 111, 614 stability, 111, 614-615 preparation and crystallization proteinase, 111, 611413 zymogen, 111, 610-611 specificity esterase activity, 111, 631-632 peptide and amide bonds, 111, 628-631 transferase activity, 111, 632633 sulfhydryl group nature, 111, 622-623 reactivity, 111, 623-624 zymogen-to-enzyme transformation autocatalytic, 111, 621 bacterial cell walls, 111,621-622 preformed streptococcal proteinase, 111, 621 subtilisin, 111, 620-621 trypsin, 111, 619-620 Streptococci endonucleases, IV, 260-261 phage lytic enzyme, V, 402-403 enzyme assay, V,403-404 group C phages and, V,404-406 other phages, V, 406-408 Streptococcus faecalis, membrane adenosine triphosphatase, X,400-416 Streptomyces, proteinases of, 111, 746-747, 752-754, 768-769 Structural information other methods of obtaining Fourier difference maps and salt difference maps, I, 4145 noncrystallographic symmetry, I, 45-46 single isomorphous replacement and variations, I, 39-11 Submandibular gland, hyaluronidase of, V, 312-313 Substrate bridge complexes electronic structure of, ATP, 11, 478-481 formation mechanism, 11,48143 reaction mechanism, 11, 483-485 Subtilisin (s) active site studies histidine and, 111, 580-584

TOPICAL SUBJECT INDEX

serine and, 111, 575-580 chemical modification lysine residues, 111, 596-598 methionine residues, 111, 598-599 tyrosine residues, 111, 599-602 historical background and development, 111, 562-563 inhibitors, dye binding and, 111, 602-605 physical, chemical and stability properties subtilisin Amylosacchariticus, 111, 566-567 subtilisin BPN’, 111, 565-566 subtilisin Carlsberg, 111, 564-565 subtilisin Novo, 111, 566 practical uses, 111, 606-607 detergents and, 111, 608 protein sequencing, 111, 807 primary structure comparison of sequences, 111, 571-575 general comparison, 111, 567 subtilisin Amylosacchariticus, 111, 569-571 subtilisin BPN’, 111,567-569 subtilisin Carlsberg, 111, 569 subtilisin Novo, 111, 509 ribonuclease and, IV, 669-672 substrate specificity and enzymic properties mechanism of action, 111, 593-596 protein and peptide substrates, 111, 584-586 synthetic substrates, 111, 586-593 transesterification and transpeptidation, 111, 593 X-ray structure background, 111, 547452 catalytic site, 111, 553-560 comparison with subtilisin Carlsberg, 111, 560 general description, 111, 552-553 Subtilisin BPN’, chemical modification, I, 203-205 Subunits acetyl coenzyme A carboxylase, VI, 60-79 adenylosuccinase, VII, 192-193 aldolases, VII, 221-224

!K)PICAL SUBJECT INDEX

amino acid decarboxylases, VI, 248-253 argininosuccinase, VII, 176-177 aspartate transcarbamylase, IX, 230-243 aspartokinases, VIII, 518-519, 541 carbamate kinase, IX, 103-104 creatine kinase, VIII, 394-395 fumarase, V, 545-549 glucose-6-phosphate isomerase, VI, 279-281 glycerol kinase, VIII, 494-495 guanidino kinases, VIII, 4-88 hexokinase, IX, 8-10, 39-40 2-keto-3-deoxy-6-phosphogluconic aldolase, VII, 285-287 phosphofructokinase, VIII, 254-257 polynucleotide phosphorylase, VII, 543-546 purine nucleoside phosphorylase, VII, 514 thiolase, VII, 396-397 transcarboxylase, 95-101 triosephosphate isomerase, VI, 327-329 uridine diphosphate-n-glucose 4’epimerase and, VI, 366-368 Succinate dehydrogenase mammaliam, XIII, 222-223 enzymic properties, XIII, 236-245 inhibitors and modifiers, XIII, 245-247 mechanism, XIII, 251-254 molecular properties, XIII, 223-236 regulatory properties, XIII, 247-251 microorganisms and, XIII, 254-256 stereospecificity, 11, 176-179 Succinyl coenzyme A mutase, stereochemistry, 11, 206-210 Succinyl coenzyme A synthetase anomalous reactions, X, 605-606 catalytic intermediates others, X, 602603 phosphoensyme, X, 596-600 succinyl phosphate, X, 600-602 Escherichia coli active site, X, 590 characterization of phosphoenzyme, x, 583-584 purification, X, 582483 quaternary structure, X, 584-589 reactivity and stability, X, 689

535 size, X, 584 sulfhydryl groups, X, 590-591 other sources, X, 594 pig heart phosphoenzyme formation, X, 592-593 polymorphism, X, 593-594 quaternary structure, X, 591-592 size, X, 591 possible regulatory properties, X, 606 reaction catalyzed, X, 581-582 steady state kinetics, X, 603-605 substrate specificity coenzyme A, X, 595 nucleoside di- and triphosphates, X, 595-596 succinate, X, 594-595 Sucrose phosphorylase covalent glucose-enzyme configuration of bond, VII, 524 isolation and properties, VII, 521423 kinetics, VII, 510-521 mechanism of catalysis, VII, 524-526 purification and properties, VII, 518-519 water and alcohols as acceptors, VII, 528632 Sulfarnatases, V, 18-19 Sulfatase A other sources, V, 36-37 ox liver, V, 27-36 Sulfatase B other sources, V, 38-39 ox liver, V, 37-38 Sulfatophosphate sulfohydrolases, V, 17-18 Sulfate, activation of, VIII, 35-37 Sulfate-activating enzymes, regulation and control, X, 665-669 Sulfhydryl groups, see also Cysteine, Thiol groups amylases, V, 244-245 argininosuccinase and, VII, 177-178 aspartate transcarbamylase, IX, 263-265 streptococcal proteinase, 111,622-624 Sulfhydryl reagents, pyruvate carboxylase and, VI, 21-22 Sulfites organic, pepsin and, 151-152

TOPICAL SUBJECT INDEX

Sulfite oxidases liver catalytic properties, XII, 417-419 molecular properties, XII, 414-417 Sullite reductase(s), XIII, 286-287 NADPHdependent, properties, XIII, 287-295

reduced methyl viologendependent, XIII, 295 Sulfonamide, carbonic anhydrase and, V, 652-658, 661 Superoxide dismutase away methods, XII, 539-540 catalytic mechanism enzymic dismutation of 02-,XII, 552-556

inhibition, XII, 556 model complexes, XII, 657 crystallization, XII, 542 determination of purity and concentration, XII, 540 historical, XII, 533 molecular properties apoenzyme, XII, 548-549 native enzyme, XII, 542-548 reconstitution, XII, 549-551 redox properties, XII, 551-552 physiological role, XII, 533 prokaryotic and mitochondrial, XII, 537-538

purification methods, XII, 538-539 sources, XII, 538 suppression, mutations and, I, 24925 1

Synovial fluid, collagenases, of, 111, 693-696

T

Tabanin, thrombin and, 111, 304-305 Tadpole liver, glycogen synthetase of, IX, 357-358 Tartrate epoxidase, VII, 201 catalytic properties factors influencing, VII, 205 kinetics, VII, 204-205 measurement of reaction, VII, 203-204

specificity and products, VII, 204

molecular properties cysteine and amino acid composition, VII, 202 molecular weight, VII, 202-203 purification, VII, 202 Temperature creatine kinase and, VIII, 420-422 p-galactosidase and, VII, 634-635 phosphoglucomutase and, VI, 455 pyruvate carboxylase and, VI, 19-20 Terminal deoxynucleotidyl transferase biological directions, X, 169-171 historical background, X, 145-147 mechanism, X, 160-161 metal ligand inhibitors, X, 161-163 other inhibitors, X, 163-165 unititiated synthesis, X, 165-168 nature of the reaction enzyme, X, 154-157 initiators, X, 149-152 metal ions, X, 152-153 nucleoside triphosphates, X, 147-149 pyrophosphate, X, 153-154 practical applications oligomeric additions, X, 167-188 polymeric additions, X, 166-167 random copolymers, X, 168-169 N-Terminal exopeptidases, background, 111, 81-82 Terminator codons biochemical identification, X, 93-95 genetic aspects, X, 88-93 Testicle, hyaluronidase of, V, 311-312 Tetranitromethane, p-hydroxydecanoyl thioester dehydrase and, V, 455 Thiamine pyrophosphokinase, X, 624427 Thiolase active site, amino acid sequence, VII, 404-405

biological function control of enzymic activity, VII, 400 metabolic significance, VII, 399 regulation of synthesis, VII, 401 catalytic properties equilibrium, VII, 397 8-ketoacyl groups and, VII, 398-399 thiol groups and, VII, 397 inhibitors, VII, 401-402 historical background, VII, 391-392 isolation and stability, VII, 393-394

TOPICAL SUBJECT INDEX

mechanism, VII, 402-404 molecular properties, VII, 394-395 occurrence, VII, 392-393 subunit structure and reversible dissociation, VII, 395-397 Thiol groups, see also Cysteine, Sulfhydryl groups fumarase, V, 549-552 papain location of, 111, 514 modification of, 111, 515-516 phosphofructokinase, VIII, 269-272 Thiolsubtilisin, displacement reactions, I, 116117 Thioredoxin reductase general properties, XIII, 144-145 light-activated reduction-neutral scmiquinone, XIII, 147-148 metabolic functions, XIII, 142-144 reduced states, mechanism, XIII, 145-147 specificity of, XIII, 144 Threonine residues chemical modification, I, 173 ribonuclease, IV, 696 Threonine, serine dehydratase(s) inhibition by serine, VII, 47 mammalian liver, VII, 39-42 microbial, VII, 4 2 4 5 other sources, VII, 4647 Threonine synthetase, properties, VII, 59-60 Thrombin amino acid and carbohydrate composition, 111, 285-286 assay of, 111, 282-283 catalytic properties general, 111, 292-293 inhibitors, 111, 302306 mechanism, 111, 306-307 protein and polypeptide substrates, 111, 295-300 purported substrates, 111, 300-302 synthetic substrates, 111, 293-295 A and B chain amino acid sequences, 111, 287-290 disulfide bridges, 111, 290-291 general, 111, 278 historical background, 111, 278-280

537 homology and tertiary structure, 111, 291-292 isolation and purity, 111, 283-284 occurrence, 111, 280-282 physical properties and molecular weight, 111, 284-285 inhibitors polypeptide, 111, 304-305 protein, 111, 305-306 synthetic, 111, 302-304 sequence homology, 111, 290 N- and C-terminal 'amino acids, 111, 287 Thymidine 3',5'-diphosphate binding, staphylococcal nuclease and, IV, 163-171 Thymidine diphosphate-L-rhamnose synthetase, properties, VI, 375-376 Thymidylate synthetase, properties, IX, 210-215 Thymine 7-hydroxylase catalytic properties, XII, 174-176 Tosyl elastase tertiary structure comparison with hypothetical model, 111, 364-365 electron density map, 111, 357-358 model of, 111, 358-363 Tosyllysine chloromethyl ketone, results obtained, I, 112-113 Tosylphenylalanine chloromethyl ketone chymotrypsin and, I, 94-96 results obtained, I, 112-113 Transaldolase distribution and purification, VII, 262-265 enzymic properties, VII, 265-268 function of, VII, 259-260 historical background, VII, 261-262 mechanism of action, VII, 271-277 metabolic role, VII, 277-280 molecular properties, VII, 269-271 Schiff bases and, 11, 359-360 Transamination, see Amino group transfer Transcarboxylase catalytic properties assay and general properties, VI, ioaiog

TOPICAL SUBJECT INDEX

equilibria and free energy, VI, 109-111

kinetics and reaction mechanism, VI, 111-1 15

role of cobalt and zinc, VI, 111 electron microscopy, VI, 101-108 historical background, VI, 84-89 molecular weight, metal content, and biotin content and linkage, VI, 92-95

purification, VI, 91-92 role in propionic acid fermentation, VI, 89-91 subunits dissociation, VI, 95-99 reconstitution, VI, 99-101 Transfer ribonucleic acid aminoacylation assays, X, 509 reaction product; X, 508-509 enzyme complexes, formation and detection, X, 521-522 “recogniiion” problem, X, 522-523 chemical modification and, X, 524-525

conclusions, X, 528 heterologous aminoacylation, X, 525-526

isoacceptor ribonucleic acid sequencing, X, 523-524 method of “dissected molecules,” X, 525

mutant ribonucleic acid analysis, X, 524

topology of synthetase complexes, X, 526-528 structure, X, 518-521 Transfer ribonucleic acid methyltransferase biological significance, IX, 184-186 occurrence, IX, 168-169 properties ionic stimulation, IX, 174-175 substrate specificity, IX, 172-174 purification, IX, 169-172 regulation bacteriophage infection, IX, 182 hormonal, IX, 179-182 inhibition, IX, 176-179

tumor tissue, IX, 183-184 virus infection, IX. 182-183 2,4,6Trinitrobensene sulfonate, inorganic pyrophosphatase and, IV, 515-516

Triosephosphate isomerase catalytic properties function in vivo, VI, 337-338 kinetic parameters, VI, 333335 mechanism, VI, 338-340 phosphoglycolate and, VI, 335-336 substrate active states, VI, 336-337 history and general, VI, 326 modification of, I, 138 molecular properties, VI, 326-327 active site structure, VI, 330333 isoenzymes, VI, 329-330 molecular weight, VI, 327 subunit structure, VI, 327-329 Trypsin activation, 111, 244-245 active-site-directed reagents and, I, 103-112

chemical modification activation and, 111, 271-272 amino groups and imidazole rings, 111, 269-270 disulfide bridges, 111, 271 tyrosine and tryptophan residues, 111, 270-271 water-insoluble derivatives, 111, 272-273

chemical structure, 111, 255-260 crystalline, heterogeneity of, 111, 254 fragment, inhibitor complex with, 111, 454

inhibitors low molecular weight, 111, 273-274 naturally occurring polypeptides, 111, 274-275

inorganic pyrophosphatase and, IV, 514

mechanism and active site, 111, 260-261 catalytic site, 111, 261-262 specificity and binding sites, llI, 262-263

physicochemical properties and stability, 111, 254-255 ribonuclease and, IV, 673-674 substrates and specificity

TOPICAL SUBJECT INDEX

assays, 111, 267-269 modified substrates, 111, 267 role of side chain, 111, 263-265 role of substrate structure, 111, 266 substitution of amino and carboxyl groups, 111, 265-266 Trypsin inhibitor pancreatic, chemical modification of, 111, 443-444 Trypsinogen activation of, 111, 261-254 preparation of, 111, 251 Tryptophanase, properties, VII, 4849 L-Tryptophan 2,3-dioxygenase catalytic properties, XII, 129-130 historical, XII, 127-128 molecular properties, XII, 128-129 Tryptophan hydroxylase, properties, XII, 240-241 Tryptophan 5-monooxygenase, properties, XII, 291-292 Tryptophan residues, chemical modification, I, 173 trypsin, 111, 270-271 Tryptophan synthetase catalytic properties individual reactions, VII, 27-30 reaction mechanism, VII, 24-26 reactions catalyzed, VII, 22-24 historical background, VII, 1-4 molecular properties a-p2 affinity and equilibrium, VII, 21 a-chain purification and properties, VII, 8-12 p-component purification and properties, VII, 16-20 mutant a chain, VII, 13-16 mutant p subunits, VII, 20-21 mutations and, I, 256-257 occurrence, VII, 4-5 other organisms and, VII, 21-22, 30 tryptophanase and, VII, 6-8, 81 use in studies on protein synthesis and regulation, VII, 5-6 Tumor viruses, deoxyribonucleic acid polymerases of, X , 211-235, see ako Reverse transcriptase p-Tyrosinase, properties, VII, 49-51 Tyrosine hydroxylase, properties, XII, 238-240

Tyrosine 3-monooxygenase, properties, XII, 290-291 Tyrosine residues, chemical modification, I, 174 creatine kinase, VIII, 434-436 fructose-l,6diphosphatase, IV, 619-620 ribonuclease, IV, 684-685 subtilisin, modification of, 111, 599-602 trypsin, 111, 270-271

U Ultraviolet absorption peptide groups, 11, 379-380 ribonuclease, IV, 714-717 Ultraviolet difference spectroscopy protein structure, 11, 408409 solvent perturbation and, 11, 410-413 typical cases, 11, 413417 Urea a-galactosidase and, VII, 635 pyruvate carboxylase and, VI, 20-21 Urease(s) catalytic properties active site studies, IV, 20-21 kinetics, IV, 18-20 mechanism, IV, 15-16 substrate specificity, IV, 16-18 jack bean enzymic activity measurement, IV, 4-5 isolation and purification, IV, 2-4 molecular properties, IV, 5-8 chemical composition and behavior, IV, 11-12 derivatives, IV, 12-13 immunological behavior, IV, 13 molecular weight, IV, 8-10 other, IV, 10-11 other sources, IV, 13-15 Uridine-cytidine kinase assay, IX, 57-58 distribution and purification, IX, 5657 kinetic and molecular properties, IX, 59-60 reaction mechanism, IX, 60-61 regulatory properties, IX, 61-62 substrate specificity, IX, 58-59

540

TOPICAL SUBJECT INDEX

Uridine diphosphate-N-acetyl-n-glucosamine 2'-epimerase, properties, VI, 371-372

Uridine diphosphate-&glucose 4'-epimerase, VI, 357-358 activators, VI, 359362 bound pyridine nucleotide protein conformation and, VI, 368369

subunit association and, VI, 365368 kinetics and specificity, VI, 358-359 mechanism of catalysis, VI, 362-366 Uridine diphosphoryl glucose pyrophosphorylase analytical and synthetic applications, VIII, 54-55 measurement of activity, VIII, 52-53 metabolic function cytology, VIII, 55-56 metabolism, VIII, 5559 regulation, VIII, 59-62 properties kinetics, VIII, 65-68 mechanism, VIII, 69-71 optima, VIII, 62 specificity, VIII, 68-69 structure, VIII, 6 2 6 5 purification, VIII, 53-54 Uridine monophosphate kinase, properties, IX, 90-91 V Velocity curves, enzyme regulation and, I, 368-369 Venom enzymes hydrolyzing phosphate esters, IV, 328 5'-nucleotidase of, IV, 342 Venom exonuclease chemical nature, IV, 317-319 general, IV, 313-317 structural determination identification of (I and u terminals, IV, 326-328 ribooligonucleotide sequences, IV, 324-326

substrate structural characteristics conformation, IV, 319-320 monophosphoryl group and, IV, 322324

nature of bases, IV, 320-321 nature of sugar, IV, 320 Vibrio cholerae, neuraminidase of, V, 328-329

Viruses neuraminidases of, V, 324 virion deoxyribonucleic acid polymerase others, X, 214 tumor viruses, X, 213-214 Visual pigments(s) light interaction early intermediates, VI, 584-585 later intermediates, VI, 585-586 overall reaction, VI, 583-584 molecular properties criteria of purity, VI, 576 linkage between retinal and protein, VI, 580-582 lipids, VI, 579-580 preparation, VI, 575-576 protein, VI, 576-577 retinal chromophore, VI, 577-579 structure and color, VI, 582-583 nature of, VI, 573-574 regeneration following illumination, VI, 587489 sites in photoreceptor, VI, 575 Vitamin B,, mechanisms, metal complexes and, 11, 528-529

Vitamin BIZcoenzyme amino group migrations and, VI, 539540

mutases and, VI, 509-511 Vitamin BIZcoenzyme-requiring dehydrases apoenzyme properties, V, 496-497 coenzyme analogs and, V, 493-496 enzyme-coenzyme interaction, V, 492493

general considerations, V, 481-482 nature of hydrogen transfer, V, 485492

substrate to product interconversion, V, 482-485 Vitamin BIZmethyltransferase alkylation studies and light stability, IX, 137-143 assay, Ix, 122-123

541

TOPICAL SUBJECT INDEX

catalytic properties methyl transfers catalyzed, IX, 129135

propyl iodide inhibition, IX, 127-129 radioactive folate binding, IX, 1 3 6 137

mechanism, IX, 151-154 occurrence, IX, 162-164 physical properties absorption spectrum, IX, 124-125 resolution-reconstitution and molecular weight, IX, 125-127 purification, IX, 123-124 role of S-adenosyl methionine, IX, 143-151

Vitamin B,. adenosyltransferase catalytic properties activators and inhibitors, VIII, 151152

assay, VIII, 148-149 kinetics and substrate specificity, VIII, 150-151 reversibility, partial reactions and mechanistic considerations, VIII, 149-150

net reaction, VIII, 145-147 purification and physical properties, VIII, 147-148 significance and distribution, VIII, 144-145

W

activation refolding, 111: 182-183 arginine 145,111, 176179 catalytic site, 111, 179-182 isoleucine 16, 111, 175-176 methionine 192, 111, 179 clastase, 111, 353-356 glossary of symbols, I, 89 molecular symmetry determination and, I, 15-18 molecular weight determination and, I, 13-15 power and limitations, I, 3-5 sub tilisin background, 111, 547-552 catalytic site, 111, 553-560 comparison with subtilisin Carlsberg, 111, 560 general description, 111, 552-553 X-ray diffraction globular macromolecules heavy atom derivatives, I, 69-86 molecules studied, I, 52-69 lysozyme analysis of structure, VII, 682-692 conformation of egg-white model, VII, 692-707 crystallography of inhibitor complexes, VII, 707-717 origin of, I, 23-25 o-Xylonate dehydrase, properties, V, 582 D-Xylose isomerase properties, VI, 349-354 role of metals, 11, 511

Wheat acetyl coenzyme A carboxylase of, VI,

Y

78-79

3’-nucleotidase of, IV, 353-354 X

Yeast adenylosuccinase of, VII, 185-191 alcohol dehydrogenase of, XI, 2223, 171-186

Xanthine oxidase, see Milk xanthine oxidase metal complexes and, 11, 533-534 properties of, XII, 56 Xanthinuria, human, molybdenum hydroxylase genetics and, XII, 400402 X-ray crystallography carboxypeptidase A, 111, 1 7 4 6 chemical modification and, I, 201-202 chymotrypsinogen, 111, 169-175

aldolase of, 11, 515-516, VII, 258 enolase carboxymethylation, V, 533 photooxidation, V, 533-534 glycogen synthetase of, IX, 359-361 hex0kinases chemical studies, IX, 10-13 mechanism, IX, 13-28 modification by added proteases, IX, 6-7

542 molecular weight and subunit structure, IX, 7-10 regulation, IX, 29-31 two isozymes and endogenous proteases, IX, 2-6 p-hydrory-p-methylglutaryl coenzyme A synthase, VII, 429-431 inorganic! pyrophosphatase catalytic properties, IV, 534-539 molecular properties, IV, 530-539 invertase, V, 292-293 biosynthesis, V, 294-295 catalytic properties, V, 300303 localization and multiple forms, V, 293-294 properties, V, 298-300 purification, V, 295-298 isoamylase of, V, 206-208 mannose-6-phosphate isomerase of, VI, 304-305

TOPICAL SUBJECT INDEX

methyltransferase of, IX, 161 mitochondria1 adenosine triphosphatase properties, X, 386 purification, X, 386-387 nicotinamide adenine dinucleotide dehydrogenase of, XIII, 216221 nuclear ribonucleic acid polymerase, X, 301-310 5'-nucleotidase of, IV, 341442 proteases acid, 111, 723-744 diisopropylfluorophosphate-sensitive, 111, 744-765 metal-chelator sensitive, 111, 765-786 other, 111, 786-795 2

Zinc, carbonic anhydrase and, V, 641-642

A 8 C D E

6 7 0 9 O

F G H 1

1 2 3 4

1 5