The Enzymes, Vol XII: Oxidation-Reduction, Part B (Electron Transfer (II), Oxygenases, Oxidases (I)), 3rd Edition

The Enzymes VOLUME XI1 OXIDA TION-REDUCTION Part B ELECTRON TRANSFER (II) OXYGENASES OXIDASES (I) Third Edition CONT...

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The Enzymes VOLUME XI1

OXIDA TION-REDUCTION Part B ELECTRON TRANSFER (II) OXYGENASES OXIDASES (I) Third Edition

CONTRIBUTORS MITCHEL T. ABBOTT L.-E. ANDR~ASSON R. C. BRAY HAROLD J. BRIGHT W. DUPPEL OSAMU HAYAISHI PETER HEMMERICH MARTHA L. LUDWIG

B. G. MALMSTROM VINCENT MASSEY STEPHEN G. MAYHEW MITSUHIRO NOZAKI GRAHAM PALMER DAVID J. T. PORTER B. REINHAMMAR V. ULLRICH

ADVISORY BOARD BRITTON CHANCE BO MALMSTROM LARS ERNSTER VINCENT MASSEY

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

Volume XI1

OXIDA TIOWREDUCTION Part B ELECTRON TRANSFER (1I) OXYGENASES OXIDASES (I)

THIRD EDITION

ACADEMIC PRESS New York San Francisco London 1975 A Subsidiary of Harcourt Brace Jovanovich, Publishers

COPYRIGHT 0 1975, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN ANY FORM OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR ANY INFORMATION STORAGE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING F R O M THE PUBLISHER.

ACADEMIC PRESS, INC. 111 Fifth Avenue, New

York. New York 10003

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

Library of Congress Cataloging in Publication Data Main entry under title: The Enzymes. Includes bibliographical references. CONTENTS: v. 1. Structure and control.-v. 2. Hydrolysis: peptide kinetics and mechanism.-v. 3. Hydrolysis: other C-N bonds, phosbonds.-v. 4. phate esters. [etc.] 1. Enzymes. 1. Boyer, Paul D., ed. [DNLM: 1. Enzymes. QU135 B791eJ 5 74.1'925 75-1 17 107 QP601 .E523 ISBN 0-12-122712-X

PRINTED IN THE UNITED STATES OF AMERICA

Contents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

List of Contributors

vii

Preface

ix

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

xi

I Introduction . . . . . . . . . . . . . . . 11. Proteins with One Iron per Center: The Rubredoxins I11. Proteins with Two Irons per Center: Two-Iron Ferredoxins . . . . IV . Proteins with Four Irons per Center . . . . . . . . . V. Proteins with Four Irons per Center and Two Centers: 8 Fe Iron-Sulfur Proteins ; Bacterial Ferredoxins . . . . . . . . . . . VI . Model Compounds . . . . . . . . . . . . . VII . Iron-Sulfur Enzymes . . . . . . . . . . . . .

2 4 15 31

Contents of Other Volumes

.

1

Iron-Sulfur

Proteins

GRAHAM PALMER

.

2

.

. . . . .

37 46

47

Flavodoxins and Electron-Transferring Flavoproteins

STEPHENG. MAYHEW AND MARTHA L. LUDWIG I. Introduction . . . . . I1. Flavodoxins . . . . . I11. Electron-Transferring Flavoprotein

3

.

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

57

58 109

Oxygenases: Dioxygenases

OSAMUHAYAISHI, MITSUHIRO NOZAXI,AND MITCHELT. ABBOTT I . Introduction . . . . . . . I1. Heme-Con taining Dioxygenases . . I11. Nonheme Iron-Containing Dioxygenases IV . n-Ketoglutarate Dioxygenases . . .

4

.

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

120 127 132 151

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

191 193

Flavin and Pteridine Monooxygenases

VINCENTMASSEYAND PETER HEMMERICH I . Introduction . . . . . . I1. Internal Flavoprotein Monooxygenases V

vi

CONTENTS

I11. External Flavoprotein Monooxygenases IV . Pterin-Linked Monooxygenases . . V. Model Studies and Possible Mechanisms

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

204 231 241

5 . Iron- and Copper-Containing Monooxygenases

V . ULLRICH AND W . DUPPEL I. Introduction

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

I1. Occurrence and Biological Importance

I11. Iron-Containing Monooxygenases . IV . Copper-Containing Monooxygenases .

. . . . . . . . .

. . . . . . . . .

253 250 258 294

6 . Molybdenum Iron-Sulfur Flavin Hydroxylases and Related Enzymes

R . C. BRAY I . General Introduction

I1. Milk Xanthine Oxidase

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

I11. Other Molybdenum Hydroxylases . . . . IV . Genetic Studies and the Molybdenum Hydroxylases . . . . . . V. Sulfite Oxidase of Liver

.

7

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

300 303 388 400 414

Flavoprotein Oxidases

HAROLD J . BRIGHT AND DAVID J . T. PORTER I. Introduction . . . . . . . . . . . . . . . 421 I1. The Flavin Coenzyme . . . . . . . . . . . . . 423 I11. Kinetic Methods Applied to Flavoprotein Oxidases . . . . . . 425 IV . The Flavoprotein Oxidases : Molecular Properties and Kinetic Mechanism 445 V . The Chemical Mechanism of Flavoprotein Oxidases . . . . . . 474

8

.

Copper-Containing Oxidases and Superoxide Dismutase

B . G. MALMSTROM. L.-E. ANDREASSON. AND B . REINHAMMAR I. Introduction

. . . . . . . . . . . . . . . . . . . . . Dismutase . . . . . . . . . . . . .

I1. Enzymes Reducing Dioxygen to Hydrogen Peroxide

I11. Superoxide IV. The Blue Copper-Containing Oxidases

. . . . . . . . .

507 511 533 557

Author Index

. . . . . . . . . . . . . . . . 581

Subject Index

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

613

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

MITCHEL T. ABBOTT (119), Department of Chemistry, San Diego State University, San Diego, California L.-E. ANDREASSON (507), Department of Biochemistry, Chalmers Institute of Technology, and University of Goteborg, Goteborg, Sweden R. C. BRAY (299), The School of Molecular Sciences, University of Sussex, Brighton, England HAROLD J. BRIGHT (421), Department of Biochemistry, School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania

W. DUPPEL (253), Fachbereich Theoretische Mediain der Universitat des Saarlandes, Homburg/Saar, Germany OSAMU HAYAISHI (119), Department of Medical Chemistry, Kyoto University Faculty of Medicine, Kyoto, Japan PETER HEMMERICH (191) , Fachbereich Biologie, Universitiit Konstana, Konstana, Germany MARTHA L. LUDWIG (57), Biophysics Research Division and Department of Biological Chemistry, The University of Michigan, Ann Arbor, Michigan

B. P. MALMSTROM (507), Department of Biochemistry, Chalmers Institute of Technology, and University of Goteborg, Goteborg, Sweden VINCENT MASSEY (191) , Department of Biological Chemistry, The University of Michigan, Ann Arbor, Michigan STEPHEN G. MAYHEW (57), Department of Biochemistry, Landbouwhogeschool, Wageningen, The Netherlands MITSUHIRO NOZAKI (119) , Department of Medical Chemistry, Kyoto University Faculty of Medicine, Kyoto, Japan vii

...

v111

LIST OF CONTRIBUTORS

GRAHAM PALMER ( I ) ,Department of Biochemistry, Rice University, Houston, Texas DAVID J. T. PORTER (421), Department of Biochemistry, School of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania B. REINHAMMAR (507), Department of Biochemistry, Chalmers Institute of Technology, and University of Goteborg, Goteborg, Sweden V. ULLRICH (253), Fachbereich Theoretische Medizin der Universitat des Saarlands, Homburg/Saar, Germany

Preface As noted irl the Preface to Volume XI of this treatise tho important area of oxidation-reduction will be covered in Volumes XI-XIII. One Advisory Board, composed of Professors Britton Chance, Lars Ernster, €30 Malmstr6m1 and Vincent Massey, has served for these three volumes. Volume XI11 is in press. This volume continues coverage of electron transfer enzymes, opening with chapters presenting the rapidly expanded information on the ironsulfur proteins and the flavin electron transfer proteins. In this group remarkable molecular architecture has been revealed. The book continues with complete coverage of the dioxygenases and monooxygenases, a fascinating group of enzymes that have been recognized for only a little over twenty years. A closely related group of enzymes, the hydroxylases, where the oxygen introduced into substrates is derived from water instead of dioxygen, is also covered. The volume closes with the first part of the coverage on oxidases ; this includes chapters on flavoprotein oxidases and copper-containing oxidases together with superoxide dismutase. The wealth of information accumulated on superoxide dismutase in the tenyear period since its discovery provides a good example of the continued vitality of our field. T o all concerned with preparation of the volume, and particularly to the authors, a warm thanks is due.

PAULD. BOYER

ix

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Contents of Other Volumes Volume I: Structure and Control

X-Ray Crystallography and Enzyme Structure David Eisenberg Chemical Modification by Active-Site-Directed Reagents Elliott Shaw Chemical Modification as a Probe of Structure and Function Louis A . Cohen Multienzyme Complexes Lester J . Reed and David J . Cox Genetic Probes of Enzyme Structure Milton J . Schlesinger Evolution of Enzymes Emil L . Smith The Molecular Basis for Enzyme Regulation D . E. Koshland, 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 . Popjiilc Proximity Effects and Enzyme Catalysis Thomas C. Bruice Enzymology of Proton Abstraction and Transfer Reactions Irwin A . Rose Kinetic Isotope Effects in Enaymic Reactions J . H . Richards Schiff Base Intermediates in Enzyme Catalysis Esmond E. SneZl and Samuel J . DiMari Some Physical Probes of Enzyme Structure in Solution Serge N . Timasheff Metals in Enzyme Catalysis Albert S. Mildvan Author Index-Subject

Index

Volume 111: Hydrolysis: Peptide Bonds

Carboxypeptidase A Jean A. Hartsuclc 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 D . M . Blow Chymotrypsin-Chemical George P. Hess

Properties and Catalysis

Trypsin B. Keil Thrombin and Prothrombin Staffan Magnwson Pancreatic Elastase B. S. Hartley and D . M . Shotton Protein Proteinase Inhibitors-Molecular Aspects Michael Laskowski, Jr., and Robert W . Sealoclc Cathepsins and Kinin-Forming and -Destroying Enzymes Lowell M . Greenbaum Papain, X-Ray Structure J . Drenth, J . N . Jamonius, R . Koekoek, and B. G. Wolthers Papain and Other Plant Sulfhydryl Proteolytic Enzymes A . N . Glazer and E m 2 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

...

Xlll

xiv

CONTENTS OF OTHER VOLUMES

Other Bacterial, Mold, and Yeast Proteases Hiroshi Matsubara and Joseph Feder Author Index-Subj ect 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 Orlowslci 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, Tz,N,, and U, Tsuneko Uchidu and Fuji0 Egami Bacterial Deoxyribonucleases I. R . Lehman Spleen Acid Deoxyribonuclease Giorgio Bernardi Deoxyribonuclease I M . Laslcowski, Sr.

CONTENTS O F OTHER VOLUMES

Venom Exonuclease M . Laskowslci, Sr. Spleen Acid Exonuclease Albert0 Bernardi and Giorgk Bernardi Nucleotide Phosphomonoesterases George I . Drummond and Masanobu Yumamoto Nucleoside Cyclic Phosphate Diesterases George I. Drummond and Masunobu Yumamoto

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-l,6-Diphosphatases 8.Pontremoli and B. L. Horecker Bovine Pancreatic Ribonuclease Frederic M . Richards and Harold W . Wycko# Author Index-Subject Index

Volume V: Hydrolysis 1Sulfate Esters, Carboxyl Esters, Glycorider) , Hydraiion

The Hydrolysis of Sulfate Esters A . B. Roy

xv

xvi

CONTENTS OF OTHER VOLUMES

Arylsulf atases R. G. Nicholls and A . B. R o y Carboxylic Ester Hydrolases Klaus Krisch Phospholipases Dona1d J. H a mhan Acetylcholinesterase Harry C. Froede and Ivwin 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 To&, and Toshizo Isemura Cellulases D. R. Whitaker Yeast and Neurospora Invertases J . Oliver Lampen Hyaluronidases Karl Meyer Neuraminidases Alfred Gottschalk and A . S. Bhargava Phage Lysozyme and Other Lytic Enzymes Alcira Tsugita Aconitase Jenny Pickworth Glusker p-Hydroxydecanoyl Thioester Dehydrase Konrad Bloch Dehydration in Nucleotide-Linked Deoxysugar Synthesis L. Glaser and H . Zarkowsky

CONTENTS OF OTHER VOLUMES

xvii

Dehydrations Requiring Vitamin B,, Coennyme 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. Lindskog, L. E. Henderson, K . K. Kannan, A. Liljas, P . 0. Nyman, and B. Strandberg Author Index-Subject

Index

Volume VI: Carboxylation and Decarboxylation (Nonoxidative), lsomerization

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 Wishnick, and M . Daniel Lane Ferredoxin-Linked Carboxylation Reactions Bob B. Buchamn 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 Elijah Adams Coenzyme B,,-Dependent Mutases Causing Carbon Chain Rearrangements H . A. Barker

B,, Coenzyme-Dependent Amino Group Migrations Thressa C. Stadtman IsopentenylpyrophosphateIsomerase P. W . Holloway

Isomerieation in the Visual Cycle Joram Heller A5-3-Ketosteroid Isomerase Paul Talalay and Ann M . Benson Author Index-Subj ect Index

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

Tryptophan Synthetase Charles Yanofslcy and Irving P . Crawford 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 OTHER VOLUMES

xix

Argininosuccinases and Adenylosuccinases Sarah Ratner Epoxidases William B. Jakoby and Thorsten A . Fjellstedt Aldolases 3.L. Horecker, Orestes Tsolas, and C. Y . Lai Transaldolase Orestes Tsolas and B. L. Horecker 2-Keto-3-deoxy-6-phosphogluconicand Related Aldolases W . A. Wood Other Deoxy Sugar Aldolases David Sidney Feingold and Patricia Ann Hoffee 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. Higgins, Jack A . Kornblatt, and Harry Rudney a-Glucan Phosphorylases-Chemical and Physical Basis of Catalysis and Regulation Donald J . Graves and Jerry H . Wang Purine Nucleoside Phosphorylase R. E . Parks, Jr., and R. P. Agarwal Disaccharide Phosphorylases John J. Mieyal and Robert H. Abeles Polynucleotide Phosphorylase T. Godefroy-Colburn and M . Grunberg-Manago

xx

CONTENTS OF OTHER VOLUMES

The Lipases P. Desnuelle p-Galactosidase Kurt Wallenjels and Rudolf Weil Vertebrate Lysoeymes Taiji Imoto, L. N . Johnson, A. C . T,North, D. C . Phillips, and J . A . Rupley Author Index-Subj ect Index

Volume Vllh Group Transfer, Part A: Nucleotidyl Transfer, Nucleosidyl Transfer, Acyl Transfer, Phosphoryl Transfer

Adenylyl Transfer Reactions E . R. Stadtman Uridine Diphosphoryl Glucose Pyrophosphorylase Richard L. Tzrrnqu&t and R. Gaurth Hansen Adenosine Diphosphoryl Glucose Pyrophosphorylase Jack Pre&s The Adenosyltransferases S. Harvey Mudd Acyl Group Transfer (Acyl Carrier Protein) P. R o y Vagelos Chemical Basis of Biological Phosphoryl Transfer S. J. Benkovic and K . J . Schray Phosphofructokinase David P. Bloxham and Henry A . Lardy Adenylate Kinase L. Noda Nucleoside Diphosphokinases R. E . Parks, Jr., and R. P. Agarwal

CONTENTS OF OTHER VOLUMES

xxi

3-Phosphoglycerate Kinase R. K . Scopes Pyruvate Kinase F. J. K a yne 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 Truj'a-Bachi Protein Kinases Dona1 A . Walsh and Edwin G. Krebs Author Index-Subject

Index

Volume IX: Group Transfer, Part B: Phosphoryl Transfer, One-Curbon Group Transfer, Glycosyl Tronsfer, Amino Group Transfer, Other Transferases

The Hexokinases Sidney P. Colowick Nucleoside and Nucleotide Kinases Elizabeth P. Anderson Carbamate Kinase L.Raijrnan and M . E . Jones

N5-Methyltetrahydr~folate-HomocysteineMethyltransferases Robert T . Taylor and Herbert Weissbach Enzymic Methylation of Natural Polynucleotides Sylvia J . Kerr and Ernest Borek Folate Coenzyme-Mediated Transfer of One-Carbon Groups Jeanne I . Ruder and F . M . Huennekens

xxii

CONTENTS OF OTHER VOLUMES

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

N-Acetylglutamate-5-Phosphotransferase Ghza Dines Author Index-Subject

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 Lucas-Lenard and Laszlo Beres Polypeptide Chain Termination W . P. Tate and C . T. Caskey Bacterial DNA Polymerases Thomas Kornberg and Arthur Kornberg Terminal DeoxynucleotidyI Transferase F. J. Bollum Eucaryotic DNA Polymerases Lawrence A . Loeb

RNA Tumor Virus DNA Polymerases Howard M . Ternin 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 3. 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

CTP Synthetase and Related Enzymes D. E . Koshland, Jr., and A . Levitzki Asparagine Synthesis A1ton Meister Succinyl-CoA Synthetase William A . Bridger Phosphoribosylpyrophosphate Synthetase and Related Pyrophosphokinases Robert L. Switzer Phosphoenolpyruvate Synthetase and Pyruvate, Phosphate Dikinase R. A . Cooper and H . L. Kornberg Sulfation Linked to ATP Cleavage Harry D. Peck, Jr. Glutathione Synthesis Alton Meister 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 (I)

Part A: Dehydrogenases (II , Electron

Kinetics and Mechanism of Nicotinamide-Nucleotide-Linked Dehy drogenases Keith Dalziel Evolutionary and Structural Relationships among Dehydrogenases Michael G. Rossmann, Anders Liljas, Carl-Ivar Brcnde'n, and Leonard J. Banaszak Alcohol Dehydrogenases Carl-Ivar Brande'n, 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 . Austen, Kenneth M . Blumenthal, and Joseph F. Nyc Malate Dehydrogenases Leonard J. Banaszak and Ralph A . Bradshaw Cytochrornes c Richard E . Dickerson and Russell Timkovich Type b Cytochromes Bunji Hagihara, Nobuhiro Sato, and Tateo Yamanaka Author Index-Subject Index Volume XIII: Oxidationdeduction, Part A: Dehydrogenases (I1I , Oxidares ill) ,Hydrogen Peroxide Cleavage

Glyceraldehyde-3-phosphateDehydrogenase J. Ieuan Harris and Michael Waters

CONTENTS OF OTHER VOLUMES

xxv

Nicotinamide Nucleotide Transhydrogenases J . Rydstroim, J . B. Hoelc, and L. Ernster Flavin-Containing Dehydrogenases Charles H . Williams, Jr. Metal-Containing Flavoprotein Dehydrogenases Youssef Hate$ and Diana L. Stiggall Cytochrome c Oxidase Winslow S. Caughey, William J. Wallace, John A . Volpe, and Shinya Yoshikawa Cytochrome c Peroxidase Takashi Yonetani Catalase Britton Chance and Gregory R. Schonbaum Author Index-Subject

Index

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Iron-Sulfur Proteins GRAHAM PALMER I . Introduction . . . . . . . . . . . . . . I1. Proteins with One Iron per Center: The Rubredoxins . . A Background . . . . . . . . . . . . . . B . Physical Properties . . . . . . . . . . . C Chemical Properties . . . . . . . . . . . I11. Proteins with Two Irons per Center: Two-Iron Ferredoxins . A . Background . . . . . . . . . . . . . . B Physical Properties . . . . . . . . . . . C Effect of Perturbants on the EPR Spectra . . . . . D . Chemical Properties . . . . . . . . . . . I V . Proteins with Four Irons per Center . . . . . . . A Background . . . . . . . . . . . . . . B Low Potential Proteins . . . . . . . . . . C . High Potential Proteins . . . . . . . . . . V. Proteins with Four Irons per Center and Two Centers: 8 Fe Iron-Sulfur Proteins; Bacterial Ferredoxins . . . A . Background . . . . . . . . . . . . . . B . Physical Properties . . . . . . . . . . . C Chemical Properties . . . . . . . . . . . VI . Model Compounds . . . . . . . . . . . . . VII . Iron-Sulfur Enzymes . . . . . . . . . . . . A The Nitrogenase System . . . . . . . . . . B . Xanthine Oxidase . . . . . . . . . . . . C Mitochondria1 IronSulfur Proteins . . . . . . .

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2

GRAHAM PALMER

1. Introduction

Following the recommendation of the IUPAC-IUB Commission on Biochemical Nomenclature ( I ) , iron proteins are divided into three groups. If the iron atom is coordinated to a porphyrin the protein is called a hemoprotein, irrespective of the chemistry of the axial ligands. If, on the other hand, the iron is coordinated to sulfur, from either cysteine or from inorganic sulfur, then the protein is to be called an iron-sulfur protein; it is these proteins which are the subject of this review. Finally, any iron protein which does not belong in either of these two categories is disposed of as “other.” This third group includes proteins such as ferritin, transferrin, hemerythin, and possibly the dioxygenases [but see 12) I.

The IUPAC-IUB recommendation then proceeds to classify the various iron-sulfur proteins into four categories, essentially as follows : 1. Ferredozin. This group comprises those iron-sulfur proteins with an equal number of iron and labile sulfur atoms, and a negative midpoint redox potential a t pH 7. They are characterized by an E P R (electron paramagnetic resonance) signal with 8 < 2 for the reduced protein. Ferredoxins are present in plants, animals, and bacteria. Ferredoxin may be abbreviated Fd. Examples : chloroplast ferredoxin adrenal ferredoxin (formerly called adrenodoxin) Pseudomonas putidu ferredoxin (formerly called putidaredoxin) Clostridium acidi-urici ferredoxin 2. High Potential Iron-Sulfur Proteins. Certain microorganisms contain a unique class of iron-sulfur proteins containing acid-labile sulfur but differing from the ferredoxins in their physical properties. No E P R signal has been detected with the reduced form of this type of protein. The oxidized form is paramagnetic with an EPR signal with a g value of about 2. At pH 7 the midpoint potential is positive. Until further characterized, the descriptive but cumbersome name, “high potential iron-sulfur protein” should be retained with the source indicated as a prefix, e.g., Chromatiurn high potential iron-sulfur protein (HiPiP) . 3. Rubredoxins. This group comprises those iron-sulfur proteins with1. IUPAC-IUB Commission on Biochemical Nomenclature, BBA 310, 295 (1973) ; ABB 160, 355 (1974). 2. W. E. Blumberg and J. Peisach, Ann. N . Y . Acad. Sci. 222, 539 (1973).

1.

IRON-SULFUR

3

PROTEINS

out acid-labile sulfur characterized by having iron in a typical mercaptide coordination, i.e., one center surrounded by four cysteine or equivalent sulfur ligands. Oxidized rubredoxin has a distinctive EPR spectrum with a line at g = 4.3, whereas the reduced pigment gives no discernible E P R signal. The redox potential for those rubredoxins now characterized are negative at pH 7.0. The full name should be listed as (source) rubredoxin (function), e.g., Pseudomows oteovorans rubredoxin, alkane w-hydroxylation. 4. Conjugated Iron-Sulfur Proteins. This group comprises those proteins containing iron and labile sulfur or iron in a typical mercaptide coordination, but also containing additional prosthetic groups. Many of the iron-containing flavoproteins, molybdenum-iron proteins, or molybdenum-iron flavoproteins are included. Frequently these proteins may contain, as a component part of the enzyme complex, characteristics (EPR, optical spectra, or redox properties) similar to a protein classified in categories 1-3. However, since they are now considered in other nomenclature systems, no specific system of naming is now recommended. If desired, a cross-reference to this category of proteins may be included in addition to the present name in order to avoid ambiguity. These recommendations were drafted in 1968 ( 1 ) and, regrettably, not published until 1973. In the interim a vast amount of chemical and physical data has been accumulated which reveals striking structural differences between proposed members of a group, e.g., chloroplast and C. acidi-urici ferredoxin, and equal striking similarities in the detailed structures of compound which nominally belong to different categories, e.g., C. acidi-urici ferredoxin and Chromatium high potential iron-sulfur protein. Because of these developments this chapter is organized somewhat differently as indicated in Scheme I. This scheme uses the number of Iron-sulfur proteins

I

I

1 Fe/active center ( e . g . , rubredoxins)

I

I

Proteins af unknown structure cf table

2 Fe/actlve center ( e . g . , chbroplast ferredoxin)

I

Hlgh potential center (e.g., ckromatium hlgh potential protein)

I

Low potential

"f"

1 center proteln (e.g., B. polymyxa ferredoxln)

Scheme I.

2 center's/protein, (e.g., C. midi-un'n'

ferredoxin)

4

GRAHAM PALMER

iron atoms per active center as the primary discriminant among the various iron-sulfur proteins ; these primary categories are further divided as indicated. Recent reviews on the iron-sulfur proteins are references 3-7. I n particular an encyclopedic treatise entitled, “Iron-Sulfur Proteins,” was published in 1973 (8). It is obviously not practical to “compete” with that book; thus, in this chapter, the author will try to present an up-todate review of this field with some emphasis on the developments of the last 2 years. The properties of one member of each category is reviewed (Scheme I) in detail while available information on the other members of each category is summarized in the tables.

II. Proteins with One Iron per Center: The Rubredoxins

A. BACKGROUND The simplest iron-sulfur protein is typified by the rubredoxin from Clostridium pasteurianum. First observed as a contamination in crude preparations of ferredoxin from that organism the protein was isolated and crystallized by Lovenberg and Sobel in 1965 (9). It is a small protein, molecular weight 6127, composed of 54 amino acids and containing one atom of iron per mole. Unlike the vast majority of iron-sulfur proteins the rubredoxins do not contain inorganic (i.e., acid-labile) sulfur. The amino acid composition is distinguished by an abundance of aspartate and glutamate residues, the absence of both arginine and histidine, and by the presence of N-formyl methionine as the N-terminus (10). The amino acid sequence is presented in Fig. 1. Crystals of this protein have produced exceptionally good X-ray scattering, and an unusually high quality X-ray structure has been obtained by Jensen 3. W. H. OrmeJohnson, Annu. R e v . Biochem. 42, 159 (1973). 4. J. C. M. Tsibris and R . W. Woody, Coord. Chem. R e v . 5, 417 (1970). 5. G. Palmer and H. Brintzinger, in “Electron and Coupled Energy Transfer in Biological Systems” (T. King and M. Klingenberg, eds.), Vol. lB, pp. 379-476. Dekker, New York, 1972. 6. L.Jensen, Annu. R e v . Biochem. 43, 461 (1974). 7. M. Llinas, Strucl. Bonding (Berlin) 17, 125 (1973). 8. W. Lovenberg, ed., “Iron-Sulfur Proteins,” Vols. 1 and 2. Academic Press, New York, 1973. 9. W. Lovenberg and B. E. Sobel, Proc. Nut. Acad. Sci. U. S. 54, 193 (1965). 10. K. McCarthy and W. Lovenberg, BBRC 40, 1053 (1970) ; K. McCarthy, Ph.D., Thesis, George Washington University, Washington, D. C., 1972.

1.

IRON-SULFUR

5

PROTEINS 20

GLU PRO

ASP

GLV

ASP

I PRO I ASP

1

F-N

3a 64

FIG. 1. The amino acid sequence of rubredoxin from C . pasteurianum. From Eaton and Lovenberg (16).

FIG. 2. Chain model of the three-dimensional structure of rubredoxin. From Jensen (6).

and his colleagues (11J2) (Fig. 2 ) . The most striking structural feature of this protein is tetrahedral coordination of the metal ion by four mer11. K. D. Watenpaugh, L. C. Sieker, J.R. Herriott, and L. H. Jensen, Acta Crystallogr., Sect. B 29,943 (1973). 12. J. R. Herriott, L. C. Sieker, L. H. Jensen, and W. Lovenberg, JMB 80, 423 (1974).

6

GRAHAM PALMER

captide sulfur atoms present in the protein; no other amino acid functions are coordinated to the metal. Although the geometry is not rigorously tetrahedral, S-Fe-S angles varying from 101O to 115O, the deviations are not so large as to deny tetrahedral as the best “first description” of the coordination geometry. However, it is clear from the details present in Raman spectra of this protein (IS) that a significant distortion of the iron-sulfur tetrahedron persists in solution. Possibly more important is the observation that the Fe-S bond length to Cys-42 is unusually short, being some 0.2 A smaller that the average Fe-S bond length in this complex; the sulfur atom of Cys-42 lies a t the surface of the protein and may well be the point a t which electrons are transferred to and from the iron. From a low resolution difference Fourier map i t has been established that no major changes in the positions of the iron and sulfur atoms occur on reduction, and it can thus be inferred that the coordination geometry in ferrous rubredoxin is also tetrahedral; this conclusion is supported by optical data taken in the near infrared (14) (see below). Although i t is well established that this rubredoxin undergoes oxidation-reduction with n = 1 and E,’ = -57 mV (9) the physiological role of this protein is not known. Indeed, with the exception of a rather unusual rubredoxin from Pseudomonas oleovorans, which functions in the o-hydroxylation of alkanes ( 1 5 ) ,the only biochemical function which has been established for these proteins is as a substitute electron carrier in certain ferredoxin-requiring reactions ; this may well be a biochemical artifact.

PROPERTIES B. PHYSICAL The coordination chemistry which is exhibited by rubredoxin is extremely rare and thus this protein has been very thoroughly characterized physicochemically for it was anticipated that tetrahedral coordination by sulfur might be central to the structure (8) of the ferredoxins ( 1 6 ) . The effective magnetic moment of oxidized rubredoxin is 5.85 & 0.2 Bohr magnetons; this establishes that the iron is high-spin (S = g) Fe3+ for which a value of 5.92 Bohr magnetons is predicted (17). I n agree13. T. Yamamoto, G. Palmer, L. Rimai, D. Gill, and I. Salmeen, Fed. Proc., Fed. Amer. SOC. E x p . Biol. 33, 1372 (1974). 14. W. Eaton and W. Lovenberg, JACS 92, 7195 (1970). 15. E. T. Lode and M. J. Coon, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 1, p. 173. Academic Press, New York, 1973. 16. G. Palmer, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 2, p. 285. Academic Press, New York, 1973. 17. W. D. Phillips, M. Poe, J. F. Weiher, C. C. McDonald, and W. Lovenberg, Nature (London) 227, 574 (1970).

1. IRON-SULFUR

7

PROTEINS

ment with this conclusion the oxidized protein exhibits strong EPR with a g value of 4.3;this resonance is typical of Fea+ subjected t o relatively small distortions [D g * B H , E/D ‘v +; (IS)]. For the reduced protein the effective magnetic moment is 5.05 i 0.2 Bohr magnetons, close to the value of 4.90 predicted for the high-spin (S = +) ferrous ion (17). No E P R has been observed for reduced rubredoxin; this is a typical property of a paramagnet containing a n even number of unpaired electrons. The magnetic and valence characteristics of the iron atom in rubredoxin in both its redox states dictate the other physical properties of this protein; i.e., all the physical properties of this active center are determined by the number of unpaired electrons and their distribution in the metal orbitals. The most commonly measured physical properties is unquestionably the visible spectrum. For high-spin Fe3+ in any symmetry the optical spectra intrinsic to the metal ion are extremely weak for these transitions are both parity and spin forbidden; molar extinction coefficients =1 might be typical (19). Thus, the intense visible color observed with oxidized rubredoxin (Fig. 3) with a molar extinction coefficient of 8800 M-’ em-1 a t 480 nm is clearly resulting from S + Fe3+charge-transfer transitions, i.e., to electronic transitions in which the center of gravity of the optical electron moves from the sulfur atom(s) in the ground state to

WAVELENGTH (nml

(a)

WAVELENGTH (nm)

fb)

FIQ.3. The optical spectra of (-) oxidized and (---) reduced spinach (a) ferredoxin and (b) rubredoxin. From Palmer (16). 18. R. Aasa, J. Chem. Phys. 52, 3919 (1972). 19. A. B. P. Lever, “Inorganic Electronic Spectroscopy,” p. 133. Elsevier, Amsterdam, 1968.

8

GRAHAM PALMER

the iron atom in the excited state. By decreasing the positive charge on the metal ion (e.g., by reduction) these transitions involving electron transfer become energetically more difficult and thus move to shorter wavelengths. I n oxidized rubredoxin the lowest energy (longest wavelength) transition is at 750 nm. On reduction the protein decolorizes and the lowest energy transition is observed a t 320 nm ( 2 0 ) .These observations are quantitatively consistent with Jorgensen’s theory of optical electronegativities for charge-transfer transitions (16). Oxidized rubredoxin exhibits intense and detailed optical activity in the visible (21); the spectrum changes dramatically on application of a magnetic field to the sample ( 2 2 ) , but no theoretical analysis of either the circular dichroism or magnetic circular dichroism spectra of this protein has been published. However, this property may well be of empirical value. Even though reduced rubredoxin is colorless it exhibits a unique and valuable optical transition a t ca. 1.6 pm (6000 cm-l) (14). This transition is the 6E + 5T d-d promotion of an electron on the metal ion; although weak ( A , N 50 M-I cm-I) it is extremely useful for structural investigations because it is diagnostic of tetrahedral coordination by mercaptide anions. In addition to its energy and intensity, this transition is also optically active; the demonstration that he/€ (the ratio of the circular dichroism to linear absorbance) is as large as 0.05 is strong proof for the d-d character of this band [this point is discussed in detail in Eaton and Lovenberg (14)1. The use of this band for structural assignments should rely on all three spectroscopic parameters, viz., energy, intensity, and optical anisotropy (23). The Mossbauer spectra of rubredoxin in both redox states are, qualitatively, those expected for high-spin iron (Table I) (24,27a,b).At low temB. E. Sobel and W. Lovenberg, Biochemistry 5 , 6 (1967). W. Lovenberg, Protides Biol. Fluids, Int. Proc. Conf. 14, 165 (1966). D. D. Ulmer, B. Holmquist, and B. L. Vallee, BBRC 51, 1054 (1973). W. A. Eaton, G . Palmer, J. A. Fee, T. Kimura, and W. Lovenberg, Proc. Nut. Acud. Sci. U.S. 68, 3015 (1971). 24. K. K. Rao, M. C. W. Evans, R. Cammack, D. 0. Hall, C. L. Thompson, P. J . Jackson, and C. E. Johnson, BJ 129, 1063 (1972). 25. W. A. Eaton and W. Lovenberg, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 2, p. 131. Academic Press, New York, 1973. 26. H. Bachmayer, L. H. Piette, K. T. Yasunobu, and H. R. Whiteley, Proc. Nut. Acad. Sci. U . S. 57, 122 (1967). 27. E. T. Lode and M. J. Coon, JBC 246, 791 (1972). 27a. C. L. Thompson, C. E. Johnson, D. P. E. Dickson, D. 0. Hall, U. Weser, and K. X. Rao, BJ 139,97 (1974). 27b. R. H. Holm, B. A. Averill, T. Herskovitz, R. B. Frankel, H. B. Gray, 0. Siiman, F. J. Grunthan, JAGS 96, 2644 (1974). 20. 21. 22. 23.

1. IRON-SULFUR

PROTEINS

9

peratures (-4.2OK) ferric rubredoxin exhibits a six-line spectrum with relative intensities 3 :2: 1:1:2 :3. From the separation of these lines one can calculate the size of the magnetic field at the 57Fe nucleus produced by the five unpaired electrons of the iron; a value of 370 -t 3 kG has . value should be compared (Table I) with been reported ( 1 7 9 7 ~ )This -550 kG for coordination by six oxygen atoms (Fe,03), -475 k G for coordination by six sulfur atoms (ferric Tris-pyrrolidyl dithiocarbamate) , and is probably typical of tetrahedral coordination by four sulfur ligands. At higher temperatures (77OK) the electron relaxation rate increases and the hyperfine splitting collapses ; no prominent features are apparent in the spectrum ( 2 4 ) .This is presumably because the electron T, is comparable to the nuclear precession frequency in the hyperfine field. [The report of a prominent doublet in the 77OK spectrum of ferric rubredoxin probably results from a contamination by extraneous Fe from the reconstitution procedure ( 1 7 ) . ] For the ferrous rubredoxin the electron relaxation rate is much more rapid and the nucleus sees the average hyperfine field (which in the H,, is zero because the number of spin-up and spin-down electrons are the same). A precise measurement of both the isomer shift and quadrupole splitting is possible (Table I ) . The increase in isomer shift is consistent with reduction, and the large quadrupole splitting is typical of high-spin ferrous iron ( 2 4 ) . Application of a magnetic field to the sample produces a change in the populations of spin-up and spin-down electrons (via the electron Zeeman effect) and the nucleus now experiences a net hyperfine field from the electrons. As a result, the spectrum changes its shape dramaticaIly and from these new spectra it is possible t o calculate that the hyperfine field a t the nucleus is ca. 200 kG. This is much smaller than 6 X -370 = -300 kG, the value one might anticipate from the value of ferric rubredoxin, because the sixth electron generates small amounts of both a n orbital field and a dipolar field a t the nucleus which total about +90 kG [see Thompson et al. (27a) for an elaboration of this point]. An important qualitative observation was the behavior of the two quadrupole lines as the magnetic field was “turned on.” The low energy line (negative velocities) splits into a doublet and the high energy line into a triplet. This establishes that the electric field gradient is negative and that the sixth (reducing) eIectron occupies dz*. From the observation that the quadrupole splitting is almost independent of temperature it follows that the next available orbital, ( d S ~ 4is) some 800 cm-l to higher energy. Thus the iron-sulfur cluster suffers a marked distortion consistent with the X-ray and Raman result. The Mossbauer spectra of rubredoxin do not offer any great surprises, and the quantitative values of the MOSS-

TABLE I MOSSBAUER PARAMETERS FOR SOMEREPRESENTATIVE IRONSULFUR PROTEINS AND SOMESELECTED INORGANIC COMPOUNDS Compound (source)

Oxidation state

Isomer shift/T (OK)

Quadrupole splitting/T ("K)

Rubredoxin (C. pasteuriunum)

Oxidized

-

-

2 Fe protein (spinach)

2 Fe protein (P. putidu)

4 Fe protein

Reduced 0.65(77)

3.1(198), 3.16(77), 3.16(1.4)

Oxidized 0.28(4.2)

0.65(4.2)

Reduced FeIrI; 0.26(4.2) Fe"; 0.55(4.2)

Fe"'; 0.68(4.2) Fe"; 3.0(4.2)

Comments

Ref.

Electron relaxation precludes any data at 195" 94 or 77°K. Large magnetic hyperfine (370 kG/Fe) observed at 4.2"K in absence of applied field. Hyperfine field = -200 kG/Fe Diamagnetic. Only nuclear Zeeman seen in 66 applied field. Paramagnetic. Strong magnetic hyperfine seen at 4.2"K, which collapses with increasing temperature, aa the electron relaxation rate increases. At ca. 77°K a simple spectrum for two inequivalent irons k observed.

Oxidized 0.18(150), 0.27(4.2) 0.60(150), 0.60(4.2) As for the spinach 2 Fe protein except the 97b magnetic hyperfine spectrum is observed 0.27(4.2) FeIII; 0.60(4.2) at all temperatures for the reduced protein. Reduced Fe"1 FeII 0.58(4.2) Fe"; -2.7(4.2)

--

Oxidized 0.28(145), 0.31(77)

0.77(195), 0.80(77)

Complex magnetic hyperfine observed at d7u 4.2"K. Irons possibly inequivalent in pairs.

Reduced 0.38(195), 0.42(77) 0.44(4.2)

1.01(195), 1.13(77)

All spectra, both with and without applied

(Chromatium) HiPiP

CL

0

field, strikingly similar to oxidized 8 Fe protein. No evidence for inequivalent irons.

GI *I

6

k B

8 Fe protein (C. pasteurianum)

Oxidized 0.39(195), 0.43(77) 0.44(4.2) Reduced 0.52(195), 0.57(77) 0.58(1.54)

[FeB4(CHzPhdl2-

0.36(300), 0.36(4)

0.75(195), 0.91(77) 1.08(4.2) 1.07(195), 1.25(77) 1.54(4.2)

All spectra similar to reduced HiPiP (see 98 above). No evidence for inequivalent irons. Complex magnetic hyperfine observed a t 4.2"K. No evidence for inequivalent irons.

1.1(300), 1.26(4)

Formerly equivalent to reduced HiPiP, and d7b oxidized 8 Fe protein.

Y

3

3 w

z

W

Fe'+ Fea+

1.4(77) 0.55(77)

Six oxygen ligands

FenOa FeBaSLOlo

Fe*+

0.87(77)

Four oxygen liganda

FeClr FeCL

Fea+ Fez+

0.53 1.20

Six chloride ligands

98a

(NMe,) FeC14 (NMe,) 2FeC14

Feg+ Fez+

0.2(77) 1.05(77)

Four chloride ligands

98a

Fe-Tris-dtca

Fea+

0.50(77)

Six sulfur ligands

98a

FeSiFb-

0

Fe-Tris-dtc stands for ferric Tris-pyrrolidyl dithiocarbamate.

98a

3

12

GRAHAM PALMER

bauer parameters should be extremely valuable in deciphering the spectra of the polyiron proteins. The high resolution NMR spectra of rubredoxin are rather disappointing because no contact-shifted resonances of the type so abundant in the polyiron iron-sulfur proteins have been observed (I?'). This is presumably because of large magnetic moments and long electron spin relaxation times which shift the resonances of interest out of instrumental range. None of these physical data presents any great surprises. Each property studied was predictable from the assumed electronic configuration of the metal and, a t least, qualitatively, the internal consistency is very satisfying. [However, attempts to provide a more rigorous analysis of the spectral properties lead to apparent contradictions, see Eaton and Lovenberg (25).1

C. CHEMICAL PROPERTIES As might be expected from the structure of these proteins, the integrity of the visible chromophore is destroyed by mercurials. Likewise, carboxymethylation of the cysteine residues by iodoacetate only occurs after the iron has been completely removed. On the other hand, neither the methionine nor any of the lysines appears to play a role in maintaining the integrity of the active center. However, reaction of the protein with N bromosuccinimide, 2-hydroxy-5-nitrobenzylbromide, or acylatiop of the tyrosines prevents reconstitution of the active center (26). The technique of using denaturation to prepare the apoprotein of ironsulfur proteins is well established. With rubredoxin, however, unusually vigorous conditions have been used to effect complete removal of the iron. Thus, Lovenberg and Williams (28) precipitated the protein with 8% trichloracetic acid and subsequently extracted the residue with warm 70% ethanol containing o-phenanthroline. On the other hand, Rao et al. (24) reported that two precipitations with 8% trichloracetic acid are adequate. The more vigorous method of denaturation may be responsible for the extraneous absorption observed in Mossbauer spectra of samples prepared with this method (17). A particularly elegant set of chemical experiments has been reported with the rubredoxin isolated from Pseudomonas oleovorans. This protein is unusual in several regards. First, its physiological function is established-to mediate electrons between a flavoprotein, NADH rubredoxin oxidoreductase, and a hydroxylase, o-hydroxylase, capable of oxidizing alkanes to the corresponding n-alcohols. 28. W. Lovenberg and W. M. Williams, Biochemistry 8, 141 (1969).

1. IRON-SULFUR

13

PROTEINS

Second, the molecular weight of this protein is 19,500 and it is a single polypeptide chain but contains two ferric ions; i.e., there appear to be two active centers. One of these appears to be labile. By all spectroscopic criteria that have been applied these centers appear very similar to each other and to the protein from C. pasteurianum. The amino acid sequence and metal sites are shown schematically in Fig. 4. Noticing the symmetry in the sequence Lode and Coon (27) surmised that each cluster of four cysteines comprised a metal-binding site. Using cyanogen bromide they cleaved the polypeptide chains a t the single methionine strategically located between the two clusters and obtained the N- and C-terminal peptides. Each of these could be reconstituted to give products with optical spectra very similar to the native protein. The reconstituted C-rubredoxin was quite stable and exhibited an activity about one-fourth of the native protein (on an iron basis). The N-rubredoxin was quite unstable ( t l I zN 30 min, 2 5 O ) and only had onetenth the native activity. Yasunobu and Lovenberg (29) have prepared the antibody to rubredoxin from C. pasteurianum. The antibody precipitated the parent rubredoxin and apo-rubredoxin but not those from M . aerogenes, P. elsdenii, and P. oleovorans. However, all four proteins were retarded on a Sepharose-antirubredoxin column suggesting that the parent rubredoxin is polyvalent whereas the foreign rubredoxin was monovalent. However, the activity of the rubredoxin-stimulated NADPH-ferredoxin reductase(Met) 'NH3-77

ss

HH

s

H

ss

=

HH

t? 7 v c o , ss

HH

s

H

ss

HH

(Met) +NH3j--yPf--)

__

-

c 0;

(Met) (A)

(6)

FIG. 4. Schematic diagram of P . oleoiiorans rubredoxin and possible models for iron-binding sites (A and €3). From Lode and Coon (27). 29. K. T. Yasunobu and W. Lovenberg, ABB 158, 84 (1973).

14

GRAHAM PALMER

cytochrome c system was readily inhibited irrespective of the rubredoxin employed. The reduction potential, Eo', of rubredoxin is about -0.057 V, n = 1 (C. pasteuriunum) (9) though there is a small variation with species; for the protein from P. oleovorans the value is -0.037 (SO). Because of its well-understood physical properties, rubredoxin is a particularly favorable protein with which to study electron transfer reactions. Furthermore, the retention of ligands and coordination geometry during electron transfer make it extremely likely that oxidation and reduction occur by an outer sphere mechanism, i.e., the oxidant and reductant do not enter the primary coordination sphere of the iron as shown diagrammatically (Fig. 5 ) . This should be contrasted with an inner sphere mechanism in which Fea+ and 0, are linked b y a bridging ligand (either S or L) in the binary complex. Jacks et al. (32) have investigated the reduction of clostridial rubredoxin b y hexammineruthenium(I1), vanadous and chromous ions. These are three well-characterized inorganic reductants. The reaction with R u ( N H 3 ) P appeared to be second order with lc = 9 X lo4 M-' sec-' in the pH range 6.3-8. No evidence for rate saturation was observed t.hough the fastest observed rate was only 77 sec-I. The reaction proceeded t o an equilibrium position which agreed well with the reduction potentials of the reactants. The reactions with V(H20)e2+and Cr(H20)e2+were significantly slower with Ic = 1.1 X lo4 M-' sec-I and 1.2 X lo3 M-' sec-', respectively. From the rate of reduction of rubredoxin with Ru" together with the equilibrium constant, one can calculate an effective rate constant for the reverse reaction; the calculated value = 3 X lo7 M-I sec-l. Thus, the inS

s-F:%

I

A.

+ L-0,-L

4

S

FIQ.6. Schematic representation of outer sphere electron transfer reactions; S and L are sulfur ligands and "generalized" ligands, respectively; 0. and 0,-are the two valences of the oxidizing metal ion. 30. J. A. Peterson and M. J. Coon, JBC 243, 329 (1968). 31. C. A. Jacks, L. E. Bennett, W. N. Raymond, and W. Lovenberg, PTOC.Nut. Acad. Sci. U.S. 71, 1118 (1974).

1.

IRON-SULFUR

15

PROTEINS

trinsic reactivity of the metalloprotein is high. This is made more striking by calculating the electron exchange rate between oxidized and reduced rubredoxin, using the relative Marcus theory (32): AEo'

1% k l l = 2 log kll - log k z z - __ 0.059

where k11, k 2 2 , and k12 are the rate constants for the processes, oxidized rubredoxin with reduced rubredoxin, Ru" with RuIII, and oxidized rubredoxin with RuII, respectively; k Z 2= 3 X lo3 14-' sec-l (31).A value of 1 X log M-' sec-l was calculated for k l l , which must be close to the diffusion limit for the reaction of two protein molecules of this size. A smaller value for k l l , niz., 2 X lo8M-' sec-I, was calculated from the rate of reaction of rubredoxin with the vanadous ion. This is still an extremely fast rate. From these results it would seem that the intrinsic reactivity of sulfurcoordinated tetrahedral iron is extremely high, a desirable property for a redox active protein. 111. Proteins with Two Irons per Center: Two-Iron Ferredoxins

A. BACKGROUND Table I1 presents a compilation of the available data on the chemical, physical, and biological properties of a number of two-iron iron-sulfur proteins. The prototype of the class is spinach ferredoxin; this is 8 small protein of molecular weight 10,660 which contains two atoms of iron and two of sulfide. The amino acid sequence (Fig. 6 ) (SS) reveals a large contribution from acidic residues while very few basic amino acids are present. The result is an extremely low isoelectric point (pH = 4.0). Five cysteine residues are present and it is probable that four of these residues are part of the active center (see below). Indeed, in the two-iron ferredoxins from Equheturn (%la) and Aphanothece sacrum (3%) the cysteine at residue 18 is replaced by a valine indicating that i t is the residues at ca. 39, 44, 47, and 77 that are present a t the active center. Methionine, 32. L.E.Bennett, Progr. Znorg. Chem. 18, 1 (1973). 33. K. T. Yasunobu and M. Tanaka, in "Iron-Sulfur Proteins" (W. Lovenberg, ed.), Vol. 2, p. 27. Academic Press, New York, 1973. 33a. H. Kagamiyama, K. K. Rao, D. 0. Hall, R. Cammack, and H. Matsubara BJ 145, 121 (1975). 33b. K. Wada, H. Kagamiyama, M. Shin, and H. Matsubara, J . Biochem. 76, 1217 ( 1974).

TABLE I1 PROPERTIES OF 2 Fe IRONSULFUR PROTEINS Source Spinach Parsley Adrenal mitochondria (adrenodoxin) Pseudomonas putida

Fe :S :g-atom protein

EPR("K)"

2:2:10,660 2:2:10,700 2:2: 13,100

1.89, 1.95, 2.05(40) 1.90, 1.96, 2.06(40) 1.93, 1.93, 2.02(100)

2:2:12,500

1.93, 1.93, 2.02(100) ~~

a

g values and temperature for convenient observation.

Wavelength (nm) and absorbance of millimolar solution.

EO' -0.420 -0.413 -0.274 -0.360 -0.235

V V V(?) V

No. of electrom

Optical properties of oxidized protein & ( A d ) *

1 1 1

325(11.9), 420(9.4), 465(8.5) 330( 12.0), 422(9.2), 463(8.3) 330(12.0), 415(9.8), 453(8.5)

1

325(150), 415(10.0), 455(9.6)

1. IRON-SULFUR

GLY Lys -

17

PROTEINS

LEU

LYS

THR

-

SER

LEU

ASN

GLN

-

ASP

ASP -

50

ALA 97

FIG.6. Comparison of the primary structure of spinach ferredoxin with those of taro, alfalfa, Leuceana glauca, and Scenedesmus. Invariant residues are underlined and conservative substitutions overlined. After Yasunobu and Tanaka (33).

histidine, phenylalanine, tryptophan, and tyrosine are scarce. Much speculation centers around the possibility that plant ferredoxins may be a product of duplication of the gene for the bacterial (8 Fe) iron-sulfur protein; the reader is referred to the literature for those arguments (54,35).Interestingly, while adrenodoxin and putidaredoxin have much 34. D. 0. Hall, R. Cammack, and K. K. Rao, Nature (London) 233, 136 (1971).

35. H. Matsubara, T. H. Jukes, C. R. Cantor, Brookhaven Symp. Biol. 21, 201 (1969).

18

GRAHAM PALMER ,.. - - .

'.' SER . .

ILE

GLN

THR

-

HIS

PHE

10

ASN

ARG

ASP -

GLY -

GLU

THR -

LEU

ARG

THR

LYS

GLY

LYS

20

ILE

GLY -

ASP

SER -

LEU -

LEU

ASP

GLN

30

ASN

LEU

ASP

ILE

ASP 34

GLY -

THR

LEU

ALA

CYS -

SER

GLY PHE GLY -

ALA

Cys

GLU

LEU

ILE

PHE

GLU

Lys

LEU

GLU

-

40

THR CYS HIS -

50

ARG

HIS

GLU

GLN

HIS

ILE

60

LEU

ILE

THR

ASN GLU -

ALA

TYR

Em

LEU -

80

GLY

CYS GLN -

ILE -

CYS

LEU

ARG

ASP VAL PRO -

ALA

VAL

PHE -

-

GLU -

ASN

ASN

MET

THR

ASP

ARG

SER

- LEU -

-

ASP

LEU c

-

FIG.7. Homology in sequence between adrenodoxin (shown) and putidaredoxin. Invariant residues are underlined while conservative substitutions are overlined. The sequences are aligned to maximize homology. Residues 1-6, 17, 39, 71, 110, and 111 are absent in putidaredoxin. After Tanaka et al. (36).

in common with the spinach protein (Table 11) their amino acid sequences (Fig. 7) , (36), while similar to each other, are entirely unrelated 36. M. Tanaka, M. Haniu, K. T. Yasunobu, K. DUB,and I. C. Gunsalus, JBC 249, 3689 (1974).

1.

IRON-SULFUR

PROTEINS

19

to the chloroplast protein. A protein physically similar to adrenodoxin has been isolated from E . coli (36‘~). In general, proteins in this class exhibit a rich red-brown color though the optical spectrum is rather undistinguished with poorly defined maximum a t ca. 330, 420, and 460 nm superimposed on a long absorption tail (Fig. 3 ) . The most accurate extinction coefficients appear to be those described by Moss et al. (37) using in situ addition of solid mercurial, Fe3+reagent and finally a small aliquot of a standard iron solution to a solution of ferredoxin in a spectrophotometer cell. Thus, spectra could be taken at all stages of the analysis with no changes in volume. A value of 9400 M-’ cm-* a t 420 nm was obtained for the spinach protein (assuming 2 Fe/mole protein). The protein can be reversibly reduced by the addition of one electron (38) which in the laboratory is usually provided by solid sodium dithionite (it is prudent to add the very minimum of this reagent (3-5 crystals) and to have the solution well buffered at ca. pH 8 ) . The reduction potential for this process has been measured as -0.42 V by several laboratories (e.g., 39). Interestingly) adrenodoxin and putidaredoxin are not as negative. The most reliable value is that of Wilson et at. (40) who reported En’= -0.235 V for putidaredoxin. Above p H 7 the potential became more negative with increasing p H with a slope of about 30 mV/pH unit. The data could be described adequately by assuming three protonic dissociations) two in the oxidized protein (pK = 8 and pK = 10) and one in the reduced protein (pK = 9). Substitution of the sulfide by selenide produced only a trivial change in the observed potentials. The data for adrenodoxin are extremely unreliable) values ranging from -0.274 V (41) to -0.36 V (4.2) have been reported. On reduction these proteins lose 50% of their visible color and the measured spectrum is even less distinguishable (Fig. 3). However) whereas the oxidized protein is diamagnetic and exhibits no EPR at low temperatures (77’K) the reduced protein contains a net unpaired electron (8= +) and exhibits a strong EPR resonance of the g = 1.94 type. I n this case the 36a. H. E. Knoell and J. Knappe, Eur. J. Biochem. 50,245 (1974). 37. T. H. Moss, D. Petering, and G. Palmer, JBC 244, 2275 (1969). 38. 5. G. Mayhew, D. Petering, G. Palmer, and G. P. Foust, JBC 244,2830 (1969). 39. K. Tagawa and D. I. Arnon, BBA 153,602 (1968). 40. G. S. Wilson, J. C. M. Tsibris, and I. C. Gunsalus, JBC 248, 6059 (1973). 41. K. Mukai, J. J. Huang, and T. Kimura, BBA 338,427 (1974). 42. K. Suzuki and R. Estabrook, quoted in R . W. Estabrook, K. Suzuki, J. I. Mason,

J. Baron, W. E. Taylor, E. R. Simpson, J. Purvis, and J. McCarthy, in “IronSulfur Proteins” (W. Lovenberg, ed.), Vol. 1, p. 193. Academic PreEs, New York, 1973.

20

GRAHAM PALMER

spectrum is rhombic with three principal g values g. = 1.89, gv = 1.96, and gE = 2.05. Putidaredoxin and adrenodoxin exhibit similar spectroscopic properties except that the E P R spectrum is axial with two principal features 811 = 2.02 and g1 = 1.94.

B. PHYSICAL PROPERTIES By virtue of a very extensive series of physicochemical measurements, principally spectroscopic, there now seems to be very IittIe doubt that the salient features of the structure of the two-iron ferredoxins are as depicted in structure I. Thus we see that the active center contains two

iron atoms linked by bridging sulfide ions and each externally coordinated by two mercaptides. The evidence supporting this structure has been extensively discussed in a recent review (16) which also presents a qualitative discussion of the phenomenon of antiferromagnetism as it is relevant to the properties of these proteins. In summary the data (43) supporting the structure are discussed below. The effect of 57Feand 7iSe isotopic substitution experiments on E P R hyperfine interactions has demonstrated that both iron atoms and both labile sulfur atoms are involved in bonding a t the active center of putidaredoxin and adrenodoxin. Similar data on parsley and spinach ferredoxins establish the role of both sulfur atoms in these proteins also. However, from the EPR data no decision on the number of participating iron atoms can be made. Experiments on putidaredoxin grown on 33Senriched media indicate that a t least one cysteine or methionine sulfur is also involved in the active site. The magnitudes of the principal components of the 67Fe magnetic hyperfine tensors have been measured by ENDOR experiments on proteins which were chemically substituted with 67Fenuclei. These experiments give effective A values for two nonequivalent iron atoms in the 43. W. R. Dunham, G . Palmer,

R. H. Sands, and A. Bearden, BBA 253,373 (1971).

21

1. IRON-SULFUR PROTEINS

reduced proteins: one iron has an almost isotropic effective A tensor of magnitude about 46 MHz (17 electron gauss), the other iron has a highly anisotropic effective A tensor with principal values of about 17, 24, and 35 MHz in adrenodoxin and putidaredoxin. The Mossbauer spectra of the oxidized proteins show a slightly broadened, single quadrupole pair which is temperature independent from 4.2" t o 77°K. The Mossbauer spectra of the reduced proteins are strongly temperature dependent with the spectrum obtained a t 4.2"K in a n applied magnetic field exhibiting well-resolved magnetic hyperfine splittings given by hyperfine tensors which are in agreement with the ENDOR results. Two nonequivalent, spin-coupled iron sites are observed for the proteins: one with the same isomer shift and quadrupole splitting exhibited in the oxidized protein spectra and with a slightly anisotropic effective A tensor for the ground ( I = 8) state of 67Fea t around -46 MHz; the other iron is a high-spin ferrous ion (large isomer shift and quadrupole splitting), and has a highly anisotropic A tensor. I n the case of the high-spin ferrous atom, the identity of the orbital ground state is contained in the electric field gradient tensor and the hyperfine tensor a t this site. This ground state has d,, symmetry in the case of parsley and spinach ferredoxin, with the symmetry in the other proteins as yet undetermined. Magnetic susceptibility measurements demontrate antiferromagnetic coupling of the iron atoms in both the oxidized and reduced forms of the proteins. These couplings result in molecular diamagnetism a t temperatures below 50°K for the oxidized proteins, and a molecular paramagnetism corresponding to that of a single unpaired electron for the reduced protein. The most precise studies (44; also A. Ehrenberg, personal communication) of the magnetic susceptibility show the existence of higher magnetic states which become populated as the temperature is increased. The factor J in the spin Hamiltonian term, +2J5,.S2, where S , and S , are the spins of the individual iron atoms, is measured to be -182 cm-l in oxidized spinach ferredoxin and -80 to -100 cm-l in reduced spinach ferredoxin. The infrared spectra of reduced parsley and spinach ferredoxin and adrenodoxin show absorption bands at about 1.6 and 2.5 pm which coincide almost exactly with those found in rubredoxin. These bands are broad, of low intensity (4, 50) and are very optically active (&/c 0.03),which characterizes them as being the electric dipole forbidden, magnetic dipole allowed d-d transitions of the ferrous iron. I n addition, the following chemical data are important. The mercurial

-

H

44. G. Palmer, W. R . Dunham, J. A. Fee, R. H. Sands, T. Iiauka, and T. Yonetani,

BBA 245, 201 (1971).

22

GRAHAM PALMER

titer of the two-iron ferredoxins can be interpreted in terms of the number of reacting cysteine and sulfide anions present in the denatured protein. In spinach ferredoxin, nine mercurial equivalents are necessary to titrate the protein. The five cysteine residues and two labile sulfur atoms are accounted for exactly by assigning the following valences to the sulfur atoms: RS- for the cysteine and S2- for the labile sulfur. To propose a persulfide structure at the iron site, one must postulate that mercurials can promote a reductive scission of the persulfide bond to satisfy the stoichiometry of the above data. Because of the unlikelihood of this reaction the mercurial titer data for the two-iron ferredoxins is an argument against a structure a t the iron centers which involves persulfides. Likewise, the titration of these proteins by oxidizing agents (potassium ferricyanide) gives a stoichiometry consistent with the oxidation of sulfur to the zero-valent state (&), again supporting the formal assignment of -1 to the mercaptide and -2 to the sulfide in the intact protein structure. These data strongly support a model for the active center of this protein which is the synthesis of two early speculations. The first of these, by Brintzinger et al. (46) proposed Structure I as that of the active center. The second, by Gibson et al. (47), proposed that in the oxidized protein the two iron atoms were high-spin Fea+ (8 = g) while in the reduced protein one iron atom was high-spin Fe3+ and the other was high-spin Fea+ (S = +). I n both redox states the iron atoms were postulated to be coupled antiferromagnetically, to yield a diamagnetic center in the oxidized protein and a paramagnetic (S = +) center in the reduced protein. Both of these suggestions appear to be correct, or more accurately, all of the available data can be adequately accounted for in terms of these two proposals. Blumberg and Peisach (48) have described a systematic characterization of the EPR properties of the 2 Fe proteins in which they relate the variations in g values to differences in charge density a t the metal ions in the different proteins. The use of selenium as a spin label for the labile sulfide was introduced by Tsibris et al. (49,60) in their penetrating work on putidaredoxin and 45. D. H. Petering, J. A. Fee, and G. Palmer, JBC 246, 643 (1971). 46. H. Brintzinger, G. Palmer, and R. H. Sands, Proc. Nut. Acad. Sci. 47. 48. 49. 50.

U.8. 55, 397 (1966). J. F. Gibson, D. 0. Hall, J. F. Thornley, and F. Whatley, Proc. Nut. Acud. Sci. U.S. 56, 987 (1986). W. E. Blumberg and J. Peisach, ABB 162, (1974). J. C. M. Tsibris, M. J. Namtvedt, and I. C. Gunsalus, BBRC 30,323 (1968). J. C.M. Tsibris, R. L. Tsai, I. C. Gunsalus, W. H. OrmeJohnson, R. E. Hansen, and H. Beinert, Proc. Nat. Acud. Sci. U.8.59,959 (1968).

1. IRON-SULFUR

PROTEINS

23

adrenodoxin ; a thorough characterization of the selenium-substituted parsley ferredoxin has been reported ( 5 1 ) . More recently, Kimura has extensively characterized adrenodoxin containing almost all possible combinations of isotopes, 6iFe, sOSe,and 7iSe. In agreement with the earlier work (49-61), he found that replacing S by Se (41,52,53) (1) has little effect on biological activity (V,,, was unchanged while K , increased approximately twofold; (2) modifies the EPR spectrum; (3) shifts the visible absorption spectrum to longer wavelengths; (4)has little, if any, effect on the near-infrared bands of the reduced protein; (5) the redox potential decreased from -274 to -288 mV; (6) Raman lines a t 397, 350, and 297 cm-I are replaced by lines a t 350, 355, and 263 cm-l on substitution. The line at 350 cm-l is interpreted as an FeS (Cys) stretch while the other lines were attributed to vibrations involving the chalcogenide. The electron nuclear double resonance spectra of the two derivatives have been compared (64). A mixed S-Se derivative was also identified from its E P R spectrum (5,931. C. EFFECT OF PERTURBANTS ON THE EPR SPECTRA

A variety of compounds produce changes in the spectral properties of these proteins. Particularly dramatic is the demonstration that in the presence of 5 M urea the intensity of the circular dichroism spectrum of oxidized spinach ferredoxin is reduced ca. 10-fold by reducing the ionic strength to 0.02 and can be restored by increasing the ionic strength to 1.0 ( 5 5 ) . Changes in the optical spectrum of adrenodoxin were also observed using dimethyl formamide, dimethyl sulfoxide, and ethylene glycol (66). The C D and optical spectrum of oxidized spinach ferredoxin does not change on incubation for one hour a t room temperature in the following solvents: 40% methanol, 20% acetone, 20% ethanol, 50% dimethyl sulfoxide, 60% ethylene glycol, 20% dimethyl formamide, 80% glycerol, and 20% 2-methylpropane diol (pH 7.4,1M NaCI) ( 5 7 ) .Higher concentration of these solvents produce discernible changes. Smaller but nonetheless real changes can be produced in the E P R 51. J. A. Fee and G. Palmer, BBA 245, 175 (1971). 52. K. Mukai, J. J. Huang, and T. Kimura, BBRC 50, 105 (1973). 53. S.-P.W. Tang, T. G. Spiro, K. Mukai, and T. Kimura, BBRC 53,869 (1973). 54. M. Bowman, L. Kevan, K. Mukai, and T. Kimura, BBA 328, 244 (1973). 55. D. H. Petering and G. Palmer, ABB 141, 456 (1970). 56. T. Kimura, BBRC 43, 1145 (1971). 57. G. Palmer, unpublished results (1970).

24

GRAHAM PALMER

spectrum of the reduced spinach ferredoxin by methanol and n-propanol (68) and by several chaotropic agents (69). Small changes in the EPR and ENDOR spectra of reduced adrenodoxin induced by dehydration have been described (60). The solvents methanol, dimethyl sulfoxide, dimethyl formamide, and glycerol were harmless at 30% V/V. Strong et al. (61) have observed that the EPR line widths of spinach ferredoxin varies almost linearly with magnetic field. The line width a t gz extrapolates to a value of ca. 5 G at zero field. Similar effects have been seen with a variety of ferredoxins including adrenodoxin and some eight-iron proteins. The implication is that the active centers of these proteins exhibit a distribution of conformation each characterized by its own set of physical parameters, e.g., g values. Whether this distribution is established by the freezing process or preexists in solution has not been resolved; however, the effect is not peculiar to the ferredoxin but has also been observed, for instance, with the cytochromes. With cytochrome c the magnitude of the effect (62) is comparable to that observed with spinach ferredoxin. Apart from the work of Eaton et al. (23)on the use of optical transitions near 1.6 to diagnose tetrahedral Fe(II)S, the only attempt to provide a detailed interpretation of the optical spectra of spinach ferredoxin focuses on relatively weak transitions between 600 and 1000 nm. Rawlings et al. (6‘3) found that the oxidized protein exhibits three obvious transitions in this region, a t 720 nm ( A , = 800), 820 nm ( A , = 260), and a t 920 nm ( A , = 80). (cf. 23,64). On reduction the bands a t 820 and 920 nm were essentially unchanged while that a t 720 nm was replaced by one a t 652 nm ( A , = 600). Rawlings et al. proposed, reasonably enough, that the longer wavelength bands are associated with the nonreducible FerI1 and suggested that they arise either from a distorted tetrahedral cluster or an Fe (111)S, unit bearing additional ligands. The bands at 720 nm in the oxidized protein and 650 nm in the reduced protein are assigned as the 6A + 4T and 5E+ ST d-d transitions of the Fe (111)Sa and Fe (11)S4 cores. However, rubredoxin, which in the oxidized form has a band a t 720 58. R. E. Coffman and B. W. Stavens, BBRC 41, 163 (1970). 59. R. Cammack, K. K. Rao, and D. 0. Hall, BBRC 44,s (1971).

60. K. Mukai, T. Kimura, J. Helbert, and L. Kevan, BRA 295, 49 (1973). OrmeJohnson I. 61. L. H. Strong, D. H. Palaith, and R. H. Sands, Fig. 3 in W. € and R. H. Sands, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 2, p. 202. Academic Press, New York, 1973. 62. C. Mailer and C. P. S. Taylor, Can. J . Biochem. 49, 695 (1971). 63. J. Rawlings, 0. Siiman, and H. Gray, Proc. N u t . Acad. Sci. U.S. 71, 126 (1973). 64. D. F . Wilson, ABB 122, 254 (1967).

1. IRON-SULFUR

25

PROTEINS

nm very similar to that observed in spinach ferredoxin, does not exhibit any band in the visible on reduction, the first transition being detected a t 320 nm (Fig. 3) ; in this system the 750 and 320 nm bands are nicely interpreted in terms of the optical electronegativities of the metal ion and its ligands (see above) (16). Thus, the assignment of the band a t 650 nm to a 6E + ST does not seem reasonable and Rawlings' alternative suggestion that the band at 650 nm is a Fe" -+ FeX1Iintervalence transition seems more plausible. Recently, Mayerle et al. (66) have synthesized the compound bis[uxylyldithiolato-p2-sulfidoferrate(111)] (Fig. 8) which has a striking resemblance to the proposed structure for spinach ferredoxin. This compound exhibits Mossbauer and optical spectra that are qualitatively similar t o that of the protein while the magnetic susceptibility and contactshifted PMR spectra are essentially identical with the oxidized protein. The complex exhibits two, well-separated, one-electron reduction potentials both of which are considerably more nEgative than the protein.

D. CHEMICAL PROPERTIES 1. Apoproteim and Reconstitution Apart from their ability to undergo one-electron oxidation reduction reactions of very low potential, the most striking property of these proteins is the ease with which one can reconstitute a totally competent molecule from a mixture of iron, sodium sulfide, a mercaptan, and any one of several different apoproteins. Sill

FIQ. 8. The structure of [FeS(SCH2)2C6H41?-anion showing the 50% probability ellipsoids of thermal vibration and omitting hydrogen atoms. After Mayerle et al. (66).

65. J. J. Mayerle, R. B. Frankel, R. H. Holm, J. A. Ibers, W. D. Phillips, and J. F. Weiher, Proc. Nut. Aead. Sci. U . S. 70,2429(1973).

26

GRAHAM PALMER

The simplest way to prepare a reconstitutable apoprotein is by precipitation of the native ferredoxin with 5% trichloroacetic acid. This procedure appears to work best if the protein is incubated briefly a t p H 12 in the presence of excess Tiron (catechol disulfonate) ( 6 6 ) . The TCAapoprotein does not contain iron, sulfide, or free sulfhydryl group, but does contain 2.5 disulfide bonds/mole ( 4 5 ) ; it must therefore be a polymer. Mercurial apoprotein is prepared by the addition of stoichiometric amounts of mercurial, usually mersalyl, to the protein; for spinach ferredoxin this means the addition of 9 moles of Hg/mole protein (5 RS-, 2 s*-) . Finally, urea-oxidant apoprotein is obtained by reacting the protein with either oxygen or ferricyanide in 5-9 M urea ( 4 5 ) .This reaction is particularly intriguing because the product so obtained contains no iron, no sulfide, and no free cysteine yet reconstitution can be effected by addition of Fe and thiol only, i.e., no sulfide need be added. Petering et al. (45) produced a variety of chemical data to suggest that the sulfide is in fact trapped in the protein as sulfur zero (SO), most probably in the form of cysteine trisulfide. It seems likely that this process is the principal mechanism for the oxygen instability of many iron proteins ( 4 5 ) . Hosein (67) has partially characterized these apoproteins and reported the following extinction coefficients: TCA-apoprotein, A , = 13,100 M-l cm-l (276 nm) ; urea apoprotein, A , = 14,300 (276 nm) ; and mersalyl apoprotein, A , = 21,500 (21,500).She further found that all three apoproteins react with the antibody to native spinach ferredoxin, whereas putidaredoxin, adrenodoxin, parsley and Synechococcus lividus ferredoxin, and ferredoxin-TPN' reductase did not react with this antibody. [Similar results have been obtained by Tel-Or et aZ. (68) while Hiedemann-Van Wyk and Kannagara (69) have used antibody for the localization of ferredoxin in the thylakoid membrane.] From far-UV C D measurements, all three apoproteins are judged to have some structure but were predominantly random. In particular, since almost one-fourth of the residues of the polypeptide chain bear COOH groups, a comparison of the mercurial apoprotein with polyglutamic acid seems apt. This apoprotein exhibits unusually large negative dichroism at 189 nm, which may be characteristic of the extended helical structure 66. W. R. Dunham, A. J. Bearden, I. T. Salmeen, G . Palmer, R. H. Sands, W. H. OrmeJohnson, and H. Beinert, BBA 253, 134 (1971). 67. €3. Hosein, Ph.D. Thesis, University of Michigan, Ann Arbor, 1973. 68. E. Tel-Or, S. Fuchs, and M. Avron, FEBS (Fed. Eur. Biochem. Soc.) Lett. 29, 156 (1973). 69. D. Hiedemann-Van Wyk and C. C. Kannagara, 2.Naturforsch. B 26,40 (1971).

1.

IRON-SULFUR PROTEINS

27

proposed for polyglutamic acid ( 7 0 ) , suggesting that the high density of negative charges in the apoprotein confers some polyglutamate-like structure on the polypeptide chain. Hosein has made a very detailed study of the optimum parameters needed for rapid and quantitative reconstitution. I n this work she exploited the circumstance that the appearance of the native protein could be followed directly in the highly colored reconstitution mixture by monitoring the relatively intense and characteristic circular dichroism of the active center. Her conclusions (67) can be summarized thus: 1. Apoprotein should be prepared at 4 O and stored a t liquid nitrogen temperatures when not in use. As the apoprotein ages at 4 O the rate of production (but not the yield) of reconstituted protein decreases. 2. The kinetics of reconstitution is complex. The maximum initial velocity-is of mixed order with a p H optimum a t 9.2, but at higher pH values reconstitution becomes autocatalytic. Thus, reconstitution is complete in 30 min a t pH 10.5 but takes 3 hr at p H 8.8. 3. The optimum temperature is 28O. The rate of reconstitution is very slow a t 4 O and 3 6 O . An Arrhenius plot was linear up to 30° with an apparent activation energy of 15 kcal mole-l. Above 30° another process seems to become important. This is probably irreversible denaturation for cooling a reconstitution mixture from 3 6 O to 2 5 O did not produce any additional reconstitution. 4. Increasing ionic strength decreased the rate of reconstitution but increased the stability of the product. The ionic composition did not appear to be critical, Tris-chloride, phosphate, and borate buffers being of comparable effectiveness. 5. The optimum reconstitution is obtained if the order of addition of reagents to the apoprotein is iron, thiol (routinely dithiothreitol) , and finally sulfide. Some evidence was obtained for the formation of an iron-protein precursor. Using careful anaerobic techniques (67) no evidence could be found for a requirement for oxygen in the reconstitution process. This is a surprising result for the major product is the oxidized protein. Some evidence was obtained for optically active intermediates but the reduced protein could not be detected, and yet in the reconstitution mixture the great excess of diothiothreitol guarantees that the iron is ferrous. Thus the question arises as to the origin of the oxidizing equivalents. One possible source is a trace disulfide contamination in the dithiol reductant: None could be detected analytically. A second alternative is that the disulfide produced by reduction of the apoprotein disulfide bonds is subsequently 70. M. L. Tiffany and S. Krimm, Biopolymers 6,1359 (1968).

28

GRAHAM PALMER

rereduced by the reduced ferredoxin formed initially. This possibility might be eliminated by investigating the reconstitution of mersalyl protein under anaerobic conditions. 2. Reaction with Exopeptidases

When spinach ferredoxin is incubated with carboxypeptidase a t 40° and PH 8.0 there is little change in the visible spectrum for 2 hr. Thereafter the absorbance decreases steadily to a new level which is about 60% of the original. Amino acid analyses (57) performed on samples withdrawn during the course of the reaction show that during the initial 2 hr about 1 mole of alanine and minor amounts of threonine and leucine had been released. By the end of the incubation period these latter two amino acids had also been released in substantial yield. Only trace amounts (<< 0.1 mole) of other amino acids could be detected on the analyzer record. Samples of ferredoxin removed from the incubation mixture during the first 2 hr and separated from the carboxypeptidase by absorption on DEAE-cellulose did not show any further decrease in absorbancy a t 420 nm. When a similar experiment was performed with leucine aminopeptidase (67), the absorbance at 420 nm decreased in an apparent first-order reaction. Identical behavior was observed if the protease was omitted, but its activator Mn2+was included in the reaction mixture. Controls lacking either the Mn2+or the Mn?+ plus the peptidase showed no measurable change in absorbance a t 420 nm even after 20 hr a t 40°. Amino acid analyses showed that no amino acids <0.01 mole) were liberated under any condition. This effect of Mnz+is probably a consequence of hydrolysis to MnOz+which is a powerful oxidant and can therefore oxidize the sulfur-rich active center. Clearly, however, the amino terminus of this protein does not react with leucine aI’nino peptidase while alanine can be split from the carboxyl terminus with no gross results: Subsequent removal of the two penultimate residues leads to denaturation. 3. Chemical Modification

Reaction of native spinach ferredoxin with tetranitroformate a t 25O and either pH 8.0 or 6.0 leads to a rapid reaction as judged by the production of the nitroformate anion a t 350 nm ( 6 7 ) .Amino acid analyses indicate that no tyrosines are modified under these conditions and it seems probable that, once again, oxidation of the active center has occurred. Reaction of the TCA apoprotein with tetranitromethane leads to the for-

1, IRON-SULFUR

29

PROTEINS

mation of 1-2 moles of mononitrotyrosine ( 6 7 ) . This product has not been reconstituted. Acetylation of the apoprotein with N-acetylimidazole leads t o the modification of 4 moles of tyrosine ( 6 7 ) ; this derivative has also not been reconstituted. 4. Electron Transfer Reactions

I n the laboratory the favorite reductant for these proteins is dithionite. The reactions of this compound are potentially complicated reactions for both S2042- and its monomeric product SO2- can function a s reductants. I n the case of spinach ferredoxin, however, the observed rate is a n accurately linear function of [S2042-]1/2over a 400 X rate of concentration (71). Thus SO2- is the sole reductant with this protein wit,h a rate constant of 8.6 M-1/2 sec-’. Since the equilibrium constant for dithionite monomerizat.ion is 1.4 X lo+ R I the true rate constant for the reaction is 2.3 X 105 M-l sec-I. One “traditionally” popular view of the biological role of the plant ferredoxins can be represented schematically, thus P-700 2 Fd-

--

+ Fd \photosyetern I Fd- + P-7OOt hv

+ T P N + redurtsae

Fd

+ T P N H + H+

Of these two reactions only the first has ever really been questioned. In particular there have been intermittent reports of a number of factors which were suspected as being the primary acceptor of the photoejected electron: I n these alternative proposals ferredoxin would then function as a secondary mediator in the electron transfer path. See, for instance, Evans and Cammack ( 7 l a ) who find that photooxidation of P-700 is only irreversible if Fd is initially oxidized . . . implying an earlier acceptor. The status of these compounds has been reviewed by Siedow et al. ( 7 2 ) . Recently, Bearden and Malkin (73) have discovered that whole and fragmented chloroplasts contain an iron-sulfur protein with an EPR spectrum significantly different from soluble spinach ferredoxin in that gz = 1.86 compared with gz = 1.89 for the soluble protein, i.e., the high field peak is shifted some 50 G upfield. This species can be produced by irradiation with 715 nm (photosystem I ) light a t 20°K. This irradiation also produces a strong radical signal which is presumably the P-700 71. D.0.Lambeth and G. Palmer, JBC 248, GO95 (1973). 71a. M. C. W. Evans and R. Cammack, BBRC 63, 187 (1975). 72. J. Siedow, C.F. Yokum, and A. San Pietro, Curr. Top. Bioenerg.

73. A.Bearden and R. Malkin, BBA 283, 456 (1973).

5, 107 (1973).

30

GRAHAM PALMER

cation. Warming the sample to 150°K for 2 min and reexamining a t 25OK showed that about one-half of the intensity of the P-700 and g = 1.86 resonances had disappeared suggesting a dark reversal of electron transfer (74).From very careful quantitative EPR measurements Bearden and Malkin (73) calculated that there was one ferredoxin for every 350 chlorophyll molecules and 1.23 iron-sulfur proteins per P-700in chloroplasts. Both the stoichiometry and the ease of electron transfer in both directions a t cyrogenic temperatures are persuasive data in support of the formulation that an iron-sulfur protein is the primary electron acceptor for photosystem I. Related experiments using optical spectroscopy (76) suggest that this new iron-sulfur protein is P-430-,a pigment with an optical spectrum similar to soluble spinach ferredoxin which gets bleached a t rates (-100 nsec) comparable to the production of P-700'. From potentiometric titrations of digitonin-treated photosystem I subchloroplasts, Ke et al. (76) identified three iron-sulfur centers: one with g values of 2.05 and 1.92;one with g values a t 2.05,1.92,and 1.89;and, finally, a resonance at g = 1.86. They were unable to locate the other components of this last resonance. Because of the many overlapping resonances it is not possible to get totally reliable estimates of intensity. However, one interpretation which satisfies the data rather simply is that there is only one principal iron-sulfur center present. As the potential of the solution is lowered from -450 to -550 mV this species appears with g values of 1.86,1.94,and 2.05.The titration curves for these species are very similar in this region. Below -550 mV the system changes and the iron-sulfur center exhibits a modified E P R spectrum in which the Q = 1.86 line is replaced by one at g = 1.89.The line a t g = 1.94 is unchanged and the line at g = 2.05 gets narrower. The titration curves for the disappearance of the g = 1.86 line, the appearance of the g = 1.89 line, and augmentation of the g = 2.05 line are also very similar. I n this interpretation the potential a t -500 mV would correspond t o an electron transfer to the iron-sulfur protein whereas that a t -575 mV would result from electron transfer to an unspecified species with a concomitant change in configuration a t the active center of the iron-sulfur protein. A similar explanation has been preferred by Evans et al. ( 7 6 ~ )However, . the report (76) that n = 2 for these titration curves raises more difficulties. At the other end of the electron transfer step involving the plant fer74. A. J. Bearden and R. Malkin, Fed. Proc., Fed. Amer. SOC. Ezp. Biol. 33, 1289 (1974). 75. B. Ke, BBA 301, 1 (1973). 76. B. Ke, R. E. Hansen, and H. Beinert, Proc. Nut. Acud. Sci. U. S. 70, 2941 (1973). 76a. M. C. W. Evens, S. G. Reeves, and R. Cammack, FEBS Lett. 4!3, 111 (1974).

1.

IRON-SULFUR

31

PROTEINS

redoxin is the flavoprotein, ferredoxin TPN-reductase. The interaction between the iron protein and the flavoprotein has been examined by workers in several laboratories who are in general agreement (see, for example, 77). I n particular, it has been shown that the two proteins form a relatively tight 1 :1 complex, the strength of which decreases markedly with ionic strength; a value of cu. 5 X M (pH 7.0) was obtained for Kd by extrapolating to zero ionic strength, whereas a t an ionic strength of 0.1 Kd was ca. M . The rate of combination of the two proteins is too fast to measure by both stopped flow and T-jump experiments (78). It is to be anticipated that the overall reduction of TPN’ by reduced spinach ferredoxin proceeds in three steps, viz.,

FTR*-

+ + +

+ +

FdFTR e Fd FTRFdFTR- e Fd FTR*T P N + H+ e F T R TPNH

+

+

where FTR- and FTR2- are the one- and two-electron reduced flavoprotein, with the reducing equivalents presumably on the flavin. However, Forti et al. (79) reported that during photosynthetic electron transport in vivo the flavoprotein is only reduced to the semiquinone form (FTR-) . This being the case, it would imply that the subsequent reduction of the pyridine nucleotide produces the neutral pyridinyl radical. This species has previously been produced in pulse radiolysis experiments and its spectrum is characterized (80). However, there is no direct evidence for its participation in a biochemical system and a more reasonable deduction from Forti’s result is that turnover proceeds via a small amount of fully reduced flavoprotein. Alternatively, there may be a second nonflavin redox active group present in this enzyme. Since no evidence for such a component is available the former interpretation is to be preferred (see also 81) .

IV. Proteins with Four Irons per Center

A. BACKGROUND

A number of examples now exist of 4 Fe iron-sulfur proteins and it seems that these can be divided into two classes: those typified by the 77. G . P. Foust, S. G. Mayhew, and V. Massey, JBC 244,964 (1969). 78. J. Siedow, D. Ballou, D. Rorabacher, and G. Palmer, unpublished. 79. G. Forti, B. A. Melandri, A. San Pietro, and B. Ke, ABB 140, 107 (1970). 80. E. J. Land and A. J. Swallow, BBA 234,36 (1971). 81. V. Massey, R. G. Mathews, 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, 1969.

32

GRAHAM PALMER

high potential iron protein (HiPiP) which has been isolated from Chrornatium (82), Rhodospirillum gelatinosa (82), Thiocapsa pfennigii (83) and possibly mitochondria (84); and a second group which includes two proteins from Bacillus polymyxn (86), one from Desulfovibrio desulfuricans (86) and possibly proteins from Desulfovibrio gigas (87,88) and Spirochaeta aurantia (89). From a comparison of the X-ray structure of HiPiP which contains a single iron cluster composed of 4 Fe3+,4 S", and 4 cysteine residues (90-92) with that of the 8 Fe ferredoxin from M . lactylyticus which contains two of these cubanelike iron-sulfur clusters (SS), it is clear that structure I1 is a common structural element in 4 n iron-sulfur proteins

T

cys 43

(n = 1, 2 ) . However, certain physical properties of these several proteins are strikingly different; in particular, HiPiP is diamagnetic when reduced and paramagnetic (8 = 4) when oxidized (37) while for the 8 Fe bacte82. K. Dus, H.DeKlerk, K. Sletten, and R. G. Bartsch, Biochim. Biophys. Actu 140, 291 (1967). 83. T . E. Meyer, S. J. Kennel, S. M. Tedro, and M. D. Kamen, BBA 292, 634 ( 1973). 84. F. J. Ruzicka and H. Beinert, Fed. Proc, Fed. Amer. SOC.Exp. RioE. 33, 1255 (1974). 85. N. A. Stombaugh, R. H. Burris, and W. H. OrmeJohnson, JBC 248, 7951 (1973). 86. J. A. Zubieta, R. Mason, and J. R. Postgate, BJ 133,851 (1973). 87. E. J. Laishley, J. Travis, and H. D. Peck, J. Bucterwl. 98, 302 (1969). 88. J. Travis, D.J. Newman, J. LeGall, and H. D. Peck, BBRC 45, 452 (1971). 89. P. W.Johnson and E. Canale-Parola, Arch. Mikrobiol. 89, 349 and 3363 (1973). 90. C. W. Carter, J. Kraut, S. T. Freer, Nguyen-huu Xuong, R. A. Alden, and R. G. Bartsch, JBC 249, 4212 (1974). 91. S. T.Freer, R. A. Alden, C. W. Carter, and J. Kraut, JBC 249, 6339 (1974). 92. C. W. Carter, J. Kraut, S. T. Freer, and R. A. Alden, JBC 250 (1975) (in press). 93. E. T. Adman, L. C. Sieker, and L. H. Jensen, JBC 248, 3987 (1973).

1. IRON-SULFUR

33

PROTEINS

rial ferredoxin and the proteins from B. polymyxa the reverse is true (94). Furthermore, the optical spectra of reduced HiPiP and the oxidized bacterial ferredoxin are identical (16). These facts can be rationalized in terms of the following three-state : scheme (16,95) Oxidized HiPiP

+e

reduced HiPiP

E

oxidized ferredoxin

+e

reduced ferredoxin

or using formal valences (16,95a) [Z Fe(III), 1 Fe(II)]

+ e = [2 Fe(III), 2 Fe(I1) f e

=

[Fe(III), 3 Fe(II)]

The question then arises as to the condition which restricts HiPiP to the left-hand redox couple and the ferredoxins to the right-hand couple. It seems probable that this results either from subtle differences in the properties of the cluster arising from differences in polypeptide conformation or, alternatively, from different microenvironments. I n HiPiP the redox cluster is buried in a hydrophobic pocket (90-92) and the protein itself is basic: I n the bacterial ferredoxins the cluster is close to the surface also in a nonpolar site and the overall charge on the protein is negative. The first direct evidence for the validity of this concept was recently provided by Cammack (96) who showed th at addition of dithionite to a solution of HiPiP in 80% dimethyl sulfoxide led to the formation of a n E P R signal with gll = 2.04 and g, z 1.93, similar to a ferredoxin. This phenomenon could be reversed by addittion of ferricyanide regenerating the original E P R. The protein could be recycled through this procedure several times without loss of optical or EPR absorption. Finally, dilution and rechromatography on DEAE-cellulose led to the recovery of 77% of the starting material. So far, this demonstration of both HiPiP-like and ferredoxin-like E P R spectra in the same protein has only becn achieved using HiPiP as the starting material. Clearly one would like to see the same phenomena demonstrated b y overoxidation of a ferredoxin, though at this time there seems to be no reason to doubt that this should also be possible. 94. M. Poe, W. D. Phillips, C. C. McDonald, and W. Lovenberg, Proc. Nut. A c u ~ . Sci. U . S . 65, 797 (1970). 95. C. W. Carter, Jr., J. Kraut, S. T. Freer, R. A. Alden, L. C. Sieker, E. Adman, and L. H. Jensen, Proc. Nut. Acad. Sci. U.S. 69, 352 (1972). 95a. The use of formal valence in this scheme is purely for pedagogic reasons. There is good reason to believe that the individual iron atoms in these clusters are essentially indistinguishable and that the mixed-valence representation which was useful for spinach ferredoxin should be abandoned. 96. R. Cammack, BBRC 54, 548 (1973).

GRAHAM PALMER

B. Low POTENTIAL PROTEINS The best characterized of these proteins was obtained from Bacillus polymyxa (86).This organism contains at least two of these 4 Fe ferredoxins, abbreviated as B. polymyxa Fd I and Fd 11. The proteins are very similar in properties (Table 111) with 4 Fe and 4 Sz-per 9000 molecular weight, very negative values for Eo (pH 7.0),n = 1, and optical spectra similar to those of 8 Fe ferredoxin from C. pasteurianum (Amax = 395 nm; A,/Fe N 4000 M-I cm-I). These three proteins show comparable activity in the phosphoroclastic reaction Pyruvate

+ n Fd + CoA

~

acetyl-CoA

+ n Fd

where n = 1 for the ferredoxin from C. pasteuriunum and 2 for those from B . polymyxa. However, the amino acid compositions of the two proteins from B. polymyxa show many differences though of a semiconservative nature. Both proteins lack histidine and tryptophan and Fd I1 also lacks arginine. The most striking difference in the physical properties of the 4 F e and 8 Fe ferredoxins is to be found in their EPR spectra. Both proteins from B. polymyza show simple rhombic EPR spectra with gz = 1.88, gv = 1.93, and ga = 2.06, somewhat similar to that observed with spinach ferredoxin. However, the EPR spectrum of the fully reduced 8 Fe protein from C. pasteurianurn shows a most complex spectrum, the nature of which will be discussed below. No Miissbauer spectra of the proteins from B . polymyxa have been published (see below). TABLE I11 COMPARISON OF 4 FE FERREDOXINS

MW Iron/mole S*-/mole Am.* (nm) A, Eo' (V) n 951

gY1 g.'

B . polymyxa In

B . polymyxa 110

D. desulfuricansb

8.aurantiaC

9 500 4.2 3.8 390 16,000 -0.38 I 1.88, 1 . 9 2 , 2 . 0 6

8800 4.0 3.3 390 16,800 -0.42 1 1.8,1.92,2.06

6300 4 4 400 31, OOOd -0.33

6300 4 4 390-400 12,000"

1

1.89,1.95,2.06

Stombaugh et al. (86). Zubieta et al. (86). c Johnson and Canale-Parola (89). d Probably erroneously calculated per 8 Fe [see Zubieta et al. (8611. Estimated from published spectra. a

-

-

1. IRON-SULFUR

35

PROTEINS

C. HIGHPOTENTIAL PROTEINS The X-ray structure of the HiPiP of Chromatiurn has been fully described in three recent articles (90-92), and the geometry of the Fe-S cube compared with the analogous structure in the bacterial Fd. These data are shown in Table IV together with the corresponding dimensions for a synthetic analog [ (E-,N) 2Fe4S1(SCH,Ph) 4 1 recently prepared and characterized by Holm and his colleagues ( 9 7 ) .The agreement between the values for bond lengths and angles is remarkable and provides a striking demonstration of the similarities in the FeS, cluster in each of the three compounds. The basic structure is that of a distorted cube with the vertices bearing Fe atoms being slightly “inside” those of the sulfide atoms so that each face of the cube is a nonplanar rhomb. It was these similarities that prompted the model relating the relationship of HiPiP to bacterial ferredoxin described above. The amino acid sequence of HiPiP has been determined (97a). The Mossbauer properties of HiPiP are summarized in Table I. The most striking data (98) are those obtained for the reduced protein shown in Fig. 9 where it is compared with data taken on the 8 Fe protein (27a) in the oxidized form. The similarities in these spectra at all temperatures and magnetic fields are obvious and provide most persuasive documentation for a similar electronic configuration for reduced HiPiP and oxidized bacterial ferredoxin and consequently for the three-state hypothesis described above. Similar Mossbauer spectra have been reported for the (27b). synthetic analog [Fe4S4(SCH,Ph,) The observations (Table I ) that (1) the isomer shifts and quadrupole splittings of oxidized and reduced bacterial ferredoxin are essentially constant and (2) all the iron atoms appear to be equivalent (certainly there is no evidence for the large quadrupole splitting associated with Fez+ found in spinach ferredoxin) shows that a “mixed-valence” model which adequately describes the two-iron proteins is not relevant for the fouriron clusters. The remaining alternatives are a fractional valence description in which each iron atom is identical and has a net charge of +2%, +2&, and +24 2-73-]

97. B. A. Averill, T. Herskovits, R. H. Holm? and J. A. Ibers, JACS 95, 3523 (1973). 97a. K. Dus, S.Tedro, and R. G. Bartsch, JBC 248,7318 (1973). 97b. E. Munck, P. G. Debrunner, J. C. M. Tsibris, and I. C. Gunsalus, Biochemistry 11, 855 (1972). 98. D. P. E. Dickson, C. E. Johnson, R. Cammack, M. C. S. Evans, D. 0. Hall, and K. K. Rao, BJ 139, 105 (1974). 98a. C. E. Johnson, J . Phys. (Paris) 35, (21-57 (1974).

TABLE IV MEANVALUESFOR EQUIVALENT A N D MODEL COMPOUND BONDLENGTHS AND ANGLESIN THE HIPIP FERREDOXIN FE&* CLUSTERS Parameter Bond lengths (A) Fe-Fe Mean Range Rms deva n Fe-S * Mean Range Rms dev n Fe-S Mean Range Rms dev n Bond angles (") Fe-S*-Fe Mean Range Rms dev n S*-Fe-S* Mean Range Rms dev n S*-Fe-S Mean Range Rms dev n 0

hot-mean-square deviation.

[Fe4S4(SCHzPh),I--]

2.72 2.68-2.78 0.04 6 2.26 2.10-2.39 0.08 12 2.20 2.17-2.22 0.02 4 74 72-76 1.3 12 104 101-109 2.4 12 115 107-120 4.9 12

2.81 2.74-2.87 0.045

6 2.32 2.18-2.45 0.09 12 .2,22 2.14-2.26 0.03 4

76 72-80 2.4 12 104 99-107 2.6 12 116 106-126 5.3 12

2.85 2.65-3.04 0.10

12 2.30 1.93-2.67 0.23 24 2.19 1.90-2.44 0.17 8

78 66-9 1 5.8 24 101 88-116 8.4 24 116 95-140 9.5 24

2.776 0.01 2 2.310

2.732

-

0.005

4 2.239 0.004 4

-

0.003 8 2.251 0.003 4 73.8

-

0.3 12 104.1 0.2 12

-

111.7-117.3

-

12

1.

37

IRON-SULFUR PROTEINS

(A)

( B)

FIG.9. A comparison of the Mossbauer spectra of (A) reduced HiPiP with the oxidized ferredoxin from (B) C. pasteuriunum. (a) At 195"K, (b) at 77"K, (c) a t 4.2"K, and (d) at 4.2"K in a magnetic field of 3 T applied perpendicular to the y-ray direction, (e) at 4.2"K in a magnetic field of 6 T applied perpendicular to the y-ray direction. (1 T = 10' G). After Thomson e t al. ( $ 7 ~ )and Dickson et al. (98).

in the three states, and a description in which the excess electrons are based on the sulfur atoms. This is still very much an open question (2'7%). The four iron clusters from both the oxidized high potential iron protein of Chromatiurn and the reduced ferredoxin from C . pasteurianurn have recently been studied by ENDOR (98b). The data suggest that the iron atoms are inequivalent in the former and equivalent in the latter.

V. Proteins with Four Irons per Center and Two Centers: 8 Fe Iron-Sulfur Proteins; Bacterial Ferredoxins

A.

BACKGROUND

The structure of the oxidized ferredoxin from Peptococcus aerogenes at 2.8 A resolution has been described by Adman et al. (93) (Fig. 10). The most striking feature of the structure is that the protein contains two active centers, each being a 4 Fe-4S2-four cysteine cluster very similar t o that described above for high potential iron protein. Indeed, a t 98b. R. E. Anderson, G. Anger, L. Petersson, A. Ehrenberg, R. Cammack, D. 0. Hall, R. Mullinger, and K. K. Rao, BBA 376,63 (1975).

38

GRAHAM PALMER

12

FIQ.10. X-Ray structure of the ferredoxin from P. aerogenes showing the carbon, iron, and sulfur positions. From Adman et ul. (93).

the present level of crystallographic resolution, the clusters in the two proteins can be said to be identical (Table I V ) . The two clusters in P. aerogenes are separated by ca. 12 A (center-tocenter) and are canted with respect to one another. For the most part it appears as if the polypeptide chain is wrapped over the surface of a prolate ellipsoid which has its principal axis defined by the line joining the two clusters. A curious feature of the structure becomes apparent when one contemplates the amino acid sequence of this protein: The eight cysteine residues occur in two clusters of four, one near the amino terminus comprising Cys-8, -11, -14, and -18, and the other, close to the carboxyl terminus, being composed of Cys-35, -38, -41,and -45 (Fig. 11). An immediate anticipation is that each of these groups of cysteines belong to one or other of the two clusters but, in fact, the polypeptide chain takes a strange turn. Cysteine residues 8, 11, and 16 all function as ligands to one cluster (I); thereupon the polypeptide chain proceeds onto the second cluster (111, which is coordinated to Cys-18, -35, -38, and -61. Finally, the polypeptide returns to cluster I to provide the fourth ligand, Cys-45. The biological motives for this surprising sequence are not a t all clear, but it may well be relevant that when C. acidi-urici ferredoxin is cleaved at Arg-29 (between the two groups of cysteine resi-

1.

IRON-SULFUR

ALA

TYR

39

PROTEINS

VAL

ILE -

ASP

SER

PRO

-

ILE

ALA

GLU

PRO

-

VAL

SER

ILE

-

5

1

- CYS

ASN

GLy

ALA

CYS

LYS

10

15

-

ASN

ALA

ILE

ILE

GLN

GLN

20

GLY 25

ASP -

ALA

ASP -

SER

30

GLY

ALA

SER

CYS

CyS ASP -

35

cys -

ALA

SER

- -s VAL

40

PRO

TYR

CY

VAL

GLY

45 ASN

PRO

50

GLY

ASP 54

FIG.11. Sequence of P. aerogenes ferredoxin. The underlined amino acids are invariant in seven anaerobic ferredoxins. Conservative substitutions occur a t the overlined amino acids. Spaces indicate assumed deletions to align sequence. After Adman et at. (93).

dues), the two fragments obtained can be reconstituted to give products with similar spectroscopic properties as the native protein but with reduced enzymic activity and significantly poorer stability (99) suggesting that the unusual sequence just described results in a gross environment for the clusters which improves their stability. Relatively few amino acid side chains occupy the volume between the clusters while the immediate environment of each center is heavily hydrophobic; cluster I (‘sees” 2 Val, 2 Ile, 2 A h , 1 Gly, 1 Tyr, and 1 (2) Pro, while I1 “sees” 3 Ile, 1 Val, 1 Gly, 1 Tyr, and 1 Pro (?). Although each cluster is associated with a tyrosine residue in M . aerogenes, the suggestion that these residues might be associated with the electron transfer process seems unlikely in view of some recent chemical experiments (see below). As yet no data are available on the X-ray structure of the reduced protein.

B. PHYSICAL PROPERTIES The Mossbauer spectra of the 8 Fe ferredoxins are discussed in Section IV,C. The EPR spectra of reduced 8 Fe iron-sulfur proteins are complicated and contain many more features (bumps) than is common (100). Orme99. W. H. Orme-Johnson, Biochem. SOC. Trans. 1,30 (1973). 100. G. Palmer, L. Mortensen, and R. H. Sands, BBRC 23,357 (1966).

40

GRAHAM PALMER

Johnson and Beinert found that the shape of the spectrum varied considerably with the degree of reduction (101). Protein which is minimally reduced exhibits a relatively simple spectrum similar to that observed with spinach ferredoxins and the 4 Fe proteins found in B. polymyxa. As the degree of reduction is increased the shape of the spectrum changes and the spectrum of the fully reduced protein is totally unlike the partially reduced compound. However, the integrated intensity of this resonance accounts for two electrons. The origin of this mysterious behavior became apparent when the spectra of partially and fully reduced protein were obtained a t two different frequencies 9 GHz (X band) and 35 GHz (K band) (102). D a ta for the protein from M . lactylyticus are shown in Fig. 12. At X band the spectrum of the partially reduced protein was principally that of a simple rhombic species with g, = 1.89, gv = 1.94, and gr = 2.07. At K band the “g values” did not change demonstrating that these features are in fact genuine g values (the minor contribution from fully reduced protein present a t X band is considerably broadened a t the higher frequency and is t,hus not apparent). This simple behavior does not hold for the fully reduced protein and the spectra a t X and K band are totally different. Thus, the features present in the spectrum of the fully reduced protein cannot be assigned as g values but, rather, it follows that the paramagnetic electrons are influenced by two magnetic fields, an internal field which is characteristic of the sample and is a constant (Hint)and the variable laboratory magnetic field (Ho). A value for the order of magnitude of Hint can be obtained from Fig. 12; the single peak present a t 11,859 G in the partially reduced protein is split into a doublet a t 11,762 and 12,012 G in the spectrum of the fully reduced case. Onehalf of this separation of 260 G is a good approximation to Hint. The simplest interpretation of these data is that the partially reduced sample contains no more than one electron per protein molecule, that is, only one of the two (assumed) identical sites is reduced and the protein exhibits single-site EPR similar to that observed with Fd I from B. poEyrnyxa (see above). In the fully reduced protein, however, both sites are paramagnetic and each exerts a magnetic field a t the other. The most obvious mechanism for this interaction is magnetic dipole-dipole (equivalent to the interaction between two bar magnets). Assuming the two sites are indeed identical

101. W.H.OrmeJohnson and H. Beinert, BBRC 36, 337 (1969). 102. R. A. Mathews, S. Charlton, R. H. Sands, and G. Palmer, JBC 249,4326 (1974).

1.

IRON-SULFUR

41

PROTEINS

I

'

I

ii2w

I

limo

I

iaw

I "

FulIY reduced

I

Imw

I

ism

I

ism

H (gouwl IEI

FIG. 12. (A) X-Band EPR spectra of the ferredoxin from M . lactylyticus 20% reduced (upper spectrum) and fully reduced (bottom spectrum). (B) Same samples observed at K band (102).

where r (A) is the distance between centers, Po is the magnetic moment, and 0 the angle subtended by the direction of Ho and the line connecting the two sites (this will change from molecule to molecule and all values of 0 should be present in frozen solution). For S = Q, Po2 is lo4 G and the angular term can be as large as 2. Then H i n t= 30,000/r3 predicting a value of 20 G if r is taken as 12 A, as indicated by the X-ray data on the oxidized protein. The value of r must be reduced to about 6 A to satisfy the experimental observation which implies a substantial conformational change on reduction such that the two centers come very much closer together. Evidence supporting a more compact structure for

42

GRAHAM PALMER

the reduced protein from C. acidi-urici has been obtained using tritium exchange methods (103). Alternatively, the value of Hintand r = 12 A and the above equation can be used to calculate a value for pLo.The value so obtained is about two and one-half larger than that for a single electron, which could only arise if the individual iron atoms were high spin and a large fraction of their magnetic moment contributed to the effective spin. Also, Hintcould arise through an electrostatic exchange interaction (J) which is anisotropic (D# 0). From Hint ( = 2 0 ) we can calculate J:

a value of 10 cm-1 is obtained. This value is large enough that it would produce a large variation in EPR intensity (and magnetic susceptibility) as the temperature is changed in the range 10°-200K. If J were negative (antiferromagnetism) then the intensity of the E P R should decrease substantially with decreasing temperature. In fact, a small increase was observed (102)which implies that J is positive (ferromagnetism) . However, as the EPR spectrum is broadening rapidly a t the higher temperatures the lower intensities observed may just be a consequence of the problems inherent in integrating spectra which contain a significant amount of intensity. Furthermore, the ferromagnetic case, leading to an S = 1 ground state, requires that the absolute EPR intensity be four-thirds of an unpaired spin per active center, while the experimental data indicate no more than one spin per active center (103a). This latter quantitation is consistent with the dipolar mechanism. Proof that the active centers are magnetically coupled comes from the observation of half-field (Am,= 2) transitions (Fig. 13). These lines are centered a t g = 2 X 1.96 indicating their origin is the g = 1.96 resonance; they are isotropic and weak, as anticipated, and can only arise from magnetically coupled species. However, this observation itself offers no obvious discrimination between the alternative mechanisms. It thus seems clear that the origin of the atypical E P R spectra observed with bacterial ferredoxins is the result of an intramolecular magnetic coupling between adjacent active centers which is almost certainly dipolar in origin. The strength of this coupling is small (ca. 1 cal), and thus the possibility that this has some bearing on the biological function of these proteins seems remote. 103. J. S. Hong and J. C. Rabinowitz, JBC 245,4995 (1970). 103s. R. A. Mathews, Ph.D. Thesis, University of Michigan, 1973; University Microfilms 73-24, 637.

1. IRON-SULFUR

PROTEINS

43

FIG.13. X-band EPR spectra of three fully reduced ferredoxins in the half-field (Am = 2) region. The overall sensitivity is about 5000 times greater than necessary t o get good quality spectra in the g = 1.91 region of the samples, and this results in a large number of background signals being observed. The line occurring a t 1700 G in all three samples appears as the degree of reduction increases above 20%; the remainder of the lines are from extraneous material. Arrows a t the top of the figure indicate the expected locations of Am = 2 signals resulting from two equivalent coupled spins both having g values of 2.00 or 1.94. The location of the Am = 2 signal a t 1700 G is consistent with both coupled spins having an average g value of 1.94 as is observed in the partially reduced samples (108).

The eight-iron ferredoxins exhibit extremely rich contact-shifted (104) NMR spectra in both redox states. These resonances were postu-

lated to arise from the protons of the cysteine residues chelated to the iron-sulfur cluster (106).This suggestion has received substantial support from the observation that the tetranuclear model compounds (106) ex104. Magnetic nuclei which bear some electron spin density will resonate a t N M R

frequencies displaced from the normal range of 0 to -10 ppm, thus rendering them particularly amenable to observation. The location of these resonances will be temperature dependent, thus facilitating their identification as contact-shifted. A detailed discussion of many aspects of this phenomenon and its exploitation will be found in Lamar et nl. (lo&). 104a. G. N. Lamar, W. D. Horrocks, and R. H. Holm, “NMR of Paramagnetic Molecules.” Academic Press, New York, 1973. 105. W. D. Phillips, and M. Poe, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 2, p. 255. 1973.

44

GRAHAM PALMER

hibit resonances of similar location and temperature dependence (cf. Table I1 ; 106). I n particular the resonances are displaced further downfield with increasing temperature consistent with the anticipated antiferromagnetism of the cluster. Furthermore, the eight (p-CH,) protons are inequivalent in both the model compounds and the protein though this effect is more prominent in the proteins, presumably because of conformational restrictions imposed by the polypeptide chain. The [13C]NMR of the ferredoxin isolated from C. acidi-urici grown on [l-W]glycine have been interpreted (107) as showing that the aromatic residues are adjacent to the metal clusters as anticipated from the X-ray structure on M . aerogenes. C. CHEMICAL PROPERTIES The 8 Fe iron-sulfur proteins are two-electron acceptors ; however, the Nernst n is one. Thus, the two centers function as independent one-electron acceptors ; note, however, the apparent midpoint potentials for the two sites will differ by RT In 4 (i.e., 36 mV) even if the sites are intrinsically identical (cf. 16). The most extensive results have been obtained by Stombaugh (108). Typically E,,' is close to -0.40 V with little dependence on pH. Note, however, that the protein from Chromatium has a value of -0.48 V for E"'. There is some evidence for electron transfer between the two sites. The contact-shifted NMR of the partially reduced protein from C . pasteurianum is not simply the sum of the spectra observed with the fully oxidized and fully reduced forms. This behavior is the opposite to that observed with the single site proteins HiPiP (109) and the ferredoxin from B. polymyxa (110), suggesting, though by no means proving, that intermolecular electron transfer is slow (< lo4 sec-') while intramolecular transfer is fast (> lo4 sec-I). A direct test of this hypothesis is provided by the concentration dependence of the NMR spectra-the intramolecular process should, of course, be concentration-independent. Evidence from E P R spectra for a magnetic coupling between the sites was described above. 106. R. H. Holm, W. D. Phillips, B. A. Averill, J. J. Mayerle, and T. Herskovitz, JAG'S 76, 2109 (1974). 107. E. L. Packer, H. Sternlicht, and J. C. Rabinowitz, Proc. Nut. Acad. Sci. U.S. 69, 3278 (1972). 108. N. A. Stombaugh, R. H. Burris, and W. H. Orme-Johnson, Fed. Proc., Fed. Amer. Sac. E x p . Biol. 33, 1254 (1974). 109. W. D. Phillips, M. Poe, C. C. McDonald, and R. G. Bartsch, Proc. Nut. Acad. Sci. U . S. 67, 682 (1970). 110. W. D. Phillips, C. C. McDonald, N. A. Stombaugh, and W. H. OrmeJohnson, Proc. Nat. Acad. Sci. U.S . 71, 160 (1974).

1.

IRON-SULFUR

PROTEINS

45

The chemical properties of these proteins have been thoroughly reviewed by Malkin (111). Much of the early work on these proteins was plagued by variable and inaccurate values for the molar absorbance of the chromophore. However, a recent thorough study by Hong and Rabinowita (112) provides a very reliable value, zliz., A , (390 nm) = 30,600 (k900) M-1 cm-l for the C . acidi-urici ferredoxin. This value was obtained by using carboxypeptidase A to determine protein via the C-terminal alanine, a technique which has also served for the protein from M . Zactylyticus (105a) and the apoprotein from spinach (67). One source of the variability in A , encountered during the early studies has been described by Gersonde et al. (115). With preparation under aerobic conditions or during lyophiliaation, the protein from C. pasteurianum undergoes partial denaturation with formation of a dimer. This dimeric protein, of molecular weight 12,000, contains 8 Fe, 8 5’-, and S cysteine and exhibits one-half activity in the phosphoroclastic assay, which suggests that during the denaturation each monomer, of molecular weight 6000, loses an iron-sulfur cluster and the associated cysteine residues from disulfide bonds with a decomposed center in a second molecule of monomer. Alternatively, the internal molecular crosslinks may arise via polysulfide bonds ( 4 5 ) as has been observed in the oxidative denaturation of spinach ferredoxins and other proteins. Subsequently Lode et al. (114) have described a most elegant piece of chemistry which provides strong evidence against the direct role of tyrosine-2 in electron transfer. Using the Edman degradation technique, the two amino terminal amino acids (Ala* and Tyr?) were removed from the 8 Fe ferredoxin of C . acidi-urici and either leucine and alanine or glycine and alanine “grafted” onto the des(A1al - Tyr’) apoferredoxin yielding the (Leu2) and (Gly‘) polypeptides which were subsequently reconstituted. The (Leu’) protein was as active as native protein in the phosphoroclastic assay and in the ferredoxin-dependent reduction of cytochrome c by ferredoxin TPN reductase and TPNH, under conditions in which the iron-sulfur protein was rate-limiting. EPR spectroscopy showed that both centers could be reduced by dithionite. There is thus a strong implication that TyrZ plays no role in the electron transfer function of center 11. A possible weakness in this argument arises from the suspicion (see above) that intracluster electron transfer 111. R. Malkin, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 2, p. 1 (1973). 112. J. S. Hong, and J. C. Rabinowitz, JBC 245, 4982 (1970). 113. K. Gersonde, E. Trittelvits, H.-E. Schlaak, and H. H. Stabel, Eur. J. Biochem. 22, 57, (1971). 114. E. T. Lode, C. L. Murray, W. V. Sweeney, and J. C. Rabinowitz, Proc. Nut. Acad. Sci. U . S . 71, 1361 (1974).

46

URAHAM PALMER

may be rapid (> lo4 sec-I). This alternative was discussed by Lode et al. (114), but they could not rule out the possibility that in this modified ferredoxin communication with the exterior was restricted to center I with rapid intramolecular electron transfer to cluster 11, whereas in the normal protein both clusters could communicate equally well with external redox agents. However, since Tyr2 is not invariant (the second residue is histidine in Clostridium tartarivorum, Peptostreptococcus elsdenii, and Clostridium thermosaccharolyticum) it is more reasonable to adopt the obvious interpretation ; Tyr-30, however, is highly conserved. Furthermore, it is perhaps worth pointing out that the energetics required in either adding or removing an electron from tyrosine is considerably larger than the overall free energy accompanying the reactions of these protein. A value of -2 V has been estimated for the reduction potential of tyrosine (116) which would make sodium in liquid ammonia a preferred but hardly physiological reductant. VI. Model Compounds

The characterization of the properties of the iron-sulfur proteins has received an enormous boost by the development (97) of synthetic methods for the preparation of realistic model compounds for the active centers of these proteins. In outline the method is 3 RSH [Fe(SR,)lm

+ 3 NaOMe + FeCL + NaSH + NaOMe -+

[Fe(SR)&, ??? crystalline product

The reaction is carried out under nitrogen with methanol as the solvent. The character of the intermediate depended critically on the nature of R, which can be either aliphatic or aromatic. The final product had the general formula [Fe4S4(SR),l2-and was usually obtained as the t-butyl ammonium salt. The structure of the compound obtained when R = benzyl mercaptan has been reported ( 9 7 ) ; it has the cubanelike structure described earlier. Some of the crystallographic parameters are shown in Table IV. If RSH is a dithiol, specifically a-xylyl-apt-dithiol, then the product has the formula [Fe,S,(SzR)2] (66) and the structure shown in Fig. 8. These compounds are being characterized by X-ray, Mossbauer, EPR, NMR, optical, and photoelectron spectroscopies (66,97,106,116), magnetic susceptibility, electrochemistry (117), and 115. R. X. Ewall and L. E. Bennett, JACS 96, 940 (1974). 116. R. B. Frankel, T. Herskovitz, B. A. Averill, R. H. Holm, P. J. Krusic, and W. D. Phillips, BBRC 58, 974 (1974). 117. B. V. DePamphilis, B. A. Averill, T. Hemkovitz, L. Que, Jr., and R. H. Holm, I A C s 96, 4159 (1974).

1.

IRON-SULFUR

47

PROTEINS

chemical reactivity and there is a gratifying correspondence between the properties of these model compounds and those observed with the protein. The reader is referred to the original papers for a full discussion of this very elegant work. One intriguing aspect of the chemistry of these compounds is the ability to undergo ligand substitution reactions of the type (108),

+

[Fe4SI(SR)4]2- nR'SH

e [FeS,(SR'),(SR),_,]*-

This reaction can be monitored by changes in optical spectrum as R'Sdisplays RS or, more spectacularly, by following the NMR of the bound and unbound alkyl groups. The efficacy of substitution approximately parallels aqueous acidities and aryl thiols which displace the t-butyl mercaptan stoichiometrically. There are several ways in which this phenomenon might be exploited biochemically (118) : (1) extrusion of intact Fe-S clusters from proteins by treatment with thiols in sufficient concentration [this could be of enormous value in characterizing clusters of unknown composition and structure which are suspected of existing in several iron-sulfur proteins (e.g., hydrogenase and nitrogenase proteins) ] ; (2) the possibility arises that electrons may be transferred to the iron-sulfur cluster via a cysteine residue in a second protein which has displaced a mercaptide group intrinsic to the cluster and formed a bridge between the two proteins; and (3) the preparation and characterization of model compounds not accessible by the direct synthetic method. The kinetics and mechanism of extrusion ) the technique have recently been studied by Dukes and Holm ( 1 1 8 ~and used with bacterial ferredoxin (118b) and other iron-sulfur proteins (118c).

Clearly, the possible utility of these chemical developments is enormous and there should be some fascinating consequences. VII. Iron-Sulfur Enzymes

The proteins that we have been considering so far are presumed to have a simple electron transfer function in much the same way as the cytochromes. In addition to these there are a number of iron-sulfur proteins which have an identifiable enzymic activity. The best known of these iron-sulfur enzymes are shown in Table V. The remainder of this 118. L. Que, Jr., M. A. Bobrik, J. A. Ibers, and R. H . Holm, JAG'S 96, 4168 (1974). 11Sa. G. R. Dukes and R. H. Holm, JACS 97,528 (1975). 118b. L. Que, Jr., R. H. Holm, and L. E. Mortensen, JACS 97, 463 (1975). 11Sc. J. R. Bale and W. H. OrmeJohnson, Proc. Nut. Acud. Sci. U . S. 72, (1975). (in press).

TABLE V PROPERTIES OF IRONSULFUR ENZYMES" Protein

Other cofactors

Fe/mole

Pyruvic dehydrogenase (C. acidi-urici)

2.4 X 10'

6

6

Thiamine

Nitrate reductaae ( M . denitrifians)

1.4 X 106

8

8

Mo

1.88.1.95,2.06

<30

Sulfite reductase (E. coli)

6 . 7 X 10s

16

16

4FAD.4FMN 4 siroheme

1.90,1.93,2.04

<30

8

8

4 heme

1.93,1.95.2.09

<30

Adenylyl sulfate reductme (D. nibas) Hydrogenase

(D.gigas)

2

x

10s

St-/mole

EPR parameters

MW

Reaction catalyzed

OKb

+

+

Pyruvate CoA 2 Fd e acetyl-CoA 2 Fd f Coz

n.a.

+

+ T P N H + H+ Not- -t + TPN+ 3 TPNH + sulfite + 3 HCe 3 TPN+ + sulfide + 3 HtO Adenylyl sulfate + 2e e sulfrte +

NOaHtO

AMP 6 X 10'

4

4

None

1.86,--,2.03

None Mo

1.87.1.94.2.06 2.01,3.65,4.28

<30

Nitrogenase (C. pasteurionum) A. Feprotein B. Be-Mo protein

5.5 2.2

104 10'

4 24

4 24

Xanthine oxidsse (milk)

3.6 X 106

8

8

2 FAD, 2 Mo

1.90,1.94,2.02 1.91,2.007,2.11

Aldehyde oxidase (liver)

2.8 X 106

8

8

2 FAD, 2 Mo

1.93,1.93.2.02

-

Dihydroorotate dehydrogenase (Zvmobacterium orolicum)

1.1

x

4

4

2FAD.2FMN

1.92,1.92,2.01

x

x

10s

2 H+

+ FdZ-SHx + Fd

+ + + + + Xanthine + On * uric acid f HzOz +

Nx 3 Fdr12 ATP 6 H+ 29 NHj 12 ADP 12 Pi 3 Fd

+ Ot f HtO e + HtOz Dihydroorotate + DPNf d orotate + DPNH -k H

Acetaldehyde acetic acid

zP

E

Mitochondria1 iron-sulfur centers

(A) DPNH-ubiquinone reductaaa

16/FMN

(B) Uhiquinone cytochrome c reductase

2/6

2/C1

Cytochromes b, CI

1.809.1.887.2.026

(C) Succinate-ubiquinone reductase

8/flavin

8/flavin

8.-substituted flavin

1.91,l.93,2.01

Flavin

(1) 1.92,1.94,2.02

DPNH-ubiquinone reductme (Candida utilis)

n.s.

16/FMN

n.a.

FMN

(1) 1.923,1 ,938.2.022
DPNH

+ COQ + H+ + DPN+CoQHz

5r + 2 cytochrome e + 2 cytochrome cz+ + 2 H+ Succinate + CoQ e fumarate +

COQHI CoQ

cg+

CoQHt

(2) ? ,1.92,2.05 (3) (4)1.85,1.98. and 2.10

+

7

* After Orme-Johnson (8)with modifications. Values given are for substrate reducible iron-sulfur centers. The other cofactors often exhibit EPR. Addition of dithionite modifies1 the E P R in some cases. b E P R conveniently observed below this temperature.

B v d

I2

2

50

GRAHAM PALMER

chapter discusses some of the recent developments with the iron-sulfur enzymes.

A. THENITROGENASE SYSTEM The component proteins of the nitrogenase system [Fe protein (azoferredoxin, Azo) and Fe-Mo protein (molybdoferredoxin, Moly) 3 have been highly purified and extensively characterized (119,120). The salient features are presented in Table VI. One unpleasant property of these proteins is their extreme sensitivity to oxygen: Less than a minute's exposure to air is enough to produce substantial denaturation in both cases (119).As a consequence the preparation and handling of these compounds is quite difficult. To minimize these complications the preparations are manipulated under inert gas and dithionite (1 mM) is included in all reagents. Furthermore, since all reactions are studied in water, it follows that H' is also always present. Now, in addition to reducing N2 to 2 NH, and acetylene to ethylene, the nitroThus, genase system also exhibits an ATP-driven reduction of H+ to Hz. under normal preparative conditions, the proteins of the nitrogenase complex contain the source of electrons (dithionite) and the oxidant (H', occasionally N,)and it is difficult if not impossible to execute experiments in which these ingredients are omitted. (The only barrier to turnover is thus the absence of MgATP.) As isolated (i.e., in the presence of dithionite) both proteins exhibit EPR (121). That exhibited by the Fe protein is very similar to those seen in single four-iron ferredoxin clusters with g values of 2.06, 1.94, and 1.87; however, the narrowness of the central feature is puzzling and leads to the suspicion that there is a second species (possibly an excited state) underlying the major rhombic species. The integrated intensity associated with this signal has been estimated to be as high as 0.8 electron per mole protein (122). Addition of RiIgATP changes the shape of the E P R spectrum markedly from rhombic to axial (121-123): The new g values are gL = 1.93 and 911 = 2.04. This transition is also produced by MgADP, MgGTP, and Mg &y-methylene ATP, but not other nucleotides; Mg2+ is necessary 119. R. R. Eady, B. E. Smith, K. A. Cook, and J. R. Postgate, BJ 128, 655 (1972) 120. J. S. Chen, J. S. Multani, and L. E. Mortensen, BBA 310, 51 (1973). 121. G. Palmer, J. S. Multani, W. C. Cretney, W. G. Zumft, and L. E. Mortensen, ABB 153, 325 (1972). 122. W. H. OrmeJohnson, W. D. Hamilton, T. Ljones, M.-Y. Tso, R. H. Burris, V. K. Shah, and W. J. Brill, Proc. N a t . Acad. Sn'. U.S. 69, 3142 (1972). 123. B. E. Smith, D. J. Lowe, and R. C. Bray, BJ 135,331 (1973).

1. IRON-SULFUR

51

PROTEINS

TABLE VI PHYSICOCHEMICAL PROPERTIES O F THE

Property Sedimentation coefficient Diffusion coefficient Frictional ratio Apparent partial specific volume Molecular weight From amino acid composition From diffusion and sedimentation coefficients From gel chromatography on Sephadex G-200 From disc electrophoresis From subunit composition Subunit composition

Acid-labile sulfide content (g-atom/mole) Metal content (g-atom/mole) Molybdenum Iron Copper Magnesium Calcium Zinc Cadmium Manganese Cobalt Specific activity Substrates Acetylene (nmole of substrate reduced min-1 mg-1 H+ hmole of substrate reduced min-1 mg-1 Nitrogen (nmole of substrate reduced min-1 mg-1 ATP (nmole of phosphate min-' mg-I Oxygen sensitivity No. of MgATP sites Ka No. of MgADP sites

Ka E P R parameters (as isolated) E P R parameters +Mg ATP Eo (PHI in presence of MgATP

NITROQENASE COMPONENTS"

MoFe protein (moIybdoferredoxin, Kpl)

Fe protein (arof erredoxin KPZ)

11 4.95 x 10-7 1.08 0.73

4.8 5.55 x 10-7 1.45 0.69

229,000 200,400

67,800 68,200

220,000

62,000

217,000 221,800 69,200 One type of subTwo types of subunit, M W unit, MW 34,600 2000 51,300 i 1700 and 59,600 f 1900 3.85 16.7 rt 10

*

1.04 f 0.1" 17.5 It 0.7 1.4 1.8 1.2 0.8 N.D.b N.D.

0.02 4 <0.1
N.D.

N.D. N.D.

1200

980

1500

1050

380

276

5400

4350

tt

= 10 min

0"

2.02, 3.78,4.27" No change -0.03 Vc No change

-

ti :0

45 17 p M c 1 (strong) 1 (weak)? 5rM 1.87, 1.94, 2.06" 1.93, 1.93, 2.04" -0.294 V" -0.400 Vc

a Modified from Eady et al. (119).Unless indicated otherwise the values used are for the proteins from K . pneumoniae. * N.D. indicates not detected. c Values for proteins from C. pasteurianum.

52

GRAHAM PALMER

for this phenomenon. With the azoferredoxin from C. pasteurianurn tho changes in shape of the E P R spectrum are complete with 2 moles of MgATP (124). This stoichiometry has been confirmed by equilibrium dialysis measurements which also yield a value of 17 p M for Kd (125), thus rationalizing the abrupt spectral transition seen with this protein. With azoferredoxin from Klebsiella pneumoniae the affinity must be considerably smaller for the titration behavior was hyperbolic rather than stoichiometric (la2). With isolated azoferredoxin it takes 2-3 sec for this conversion to be completed (126). In addition to this change in the EPR spectrum addition of MgATP produces several other profound effects. First, it causes the redox potential of the paramagnetic center in the protein to fall by over -0.10 V, from -0.29 V to -0.40 V [pH 7.5 (126)l.This decrease in potential is presumably of importance to the catalytic mechanism. Furthermore, it renders the iron readily available to chelation by dipyridyl ( C . pasteurianum) (197); it exposes an additional 8 SH groups over those normally available to 5,5’-dithiobis- (2-nitrobenzoate) ; and, in the absence of dithionite, causes a polymerization of the protein ( K . pneumoniae) (128) ; and, finally, potentiates the air sensitivity of a membrane-bound nitrogenase (Azotobacter).This last effect may be a consequence of metal chelation by the ATP for it is duplicated by Chelex (129). Changes in the EPR spectra similar to those described above have been obtained by addition of 5 M urea to the protein (124). All of these data are consistent in pointing to a major conformational change in the protein associated with the addition of MgATP. Interestingly, MgADP, which has one high affinity site on the protein (Ka= 5 p M ) , produces a biphasic change in the EPR spectrum. One mole of MgADP produces a smaller but characteristic loss in anisotropy; it requires a large excess of MgADP to convert this intermediate species into the characteristic “MgATP form.” MgADP also lowers the potential of azoferredoxin (126) but is much less effective than MgATP in exposing the iron (127‘). The E P R spectrum of the Fe-Mo protein is quite unusual with g values of 2.01, 3.78, and 4.27. This resonance is characteristic of a paramagnetic system containing three unpaired electrons (8 = $) (121). Experiments using 96Mo-enriched protein ruled out the possibility that the resonance resulted from MorI1 (4 d 3 ) (I%), and the fact that the protein is main124. W. G. Zumft, G. Palmer, and L. E. Mortensen, BBA 292, 413 (1974). 125. M.-Y. Tao, and R. H. Burris, BBA 309,263 (1973). 126. W.G.Zumft, L. E. Mortensen, and G. Palmer, Eur. J. Biochem. 46,525 (1974). 127. G.A. Walker and L. Mortensen, BBRC 53, 904 (1973). 128. R. N.F.Thorneley and R. R. Eady, BJ 133,405 (1973). 129. M.G.Yates, Eur. J. Biochem. 29, 386 (1972).

1.

IRON-SULFUR

PROTEINS

53

tained in dithionite throughout the preparation made it quite unlikely tha t the species responsible for the E P R in high-spin Fe3+in five coordination for which the S = $ state has been observed (130). It seems most probable that the resonance results from an iron-sulfur cluster containing three or more iron atoms which are spin-coupled to give a resultant S = Q. I n a very interesting set of Mossbauer measurements, Smith and Lang (132) have provided some additional insights into the proteins from K . pneumoniae. Azoferredoxin (Fe protein) exhibits Mossbauer parameters similar to the 4 Fe cluster of the bacterial ferredoxin, thus reinforcing the suppositions based on the E P R spectra (121) and optical spectra (132). In molybdoferredoxin (Fe-Mo protein) Smith and Lang deduced the presence of three species labeled M,, M,, and M,; M, is ascribed to a dimer containing two high-spin ferrous ions, while M, and M, are suggested to be clusters (octamers?) of low-spin ferrous iron and a ferric-ferrous mixture, respectively. (The analytical data indicate 18 Fe/220,000 in this protein.) It was speculated that M, is the species giving rise to the novel EPR. If either protein is treated with oxygen it is irreversibly denatured. However, cautious treatment with organic oxidants or ferricyanide yields an oxidized (EPR-free) protein which can be reduced back to the EPRpositive state by dithionite. The rate of this reaction is very rapid for azoferredoxin ( tM5 5 msec) but extremely slow for molybdoferredoxin (t% 3-5 min) (119). The reduction potential for molybdoferredoxin has also been determined by E P R and a value of -0.03 V (pH 7.5) obtained for the protein from C . pasteurianum (126). However, the protein for Chromatium apparently exhibits two potential determining steps with E,' = -0.06 and -0.26 V ( pH 7.5) (133). No evidence for this more negative step was found with the protein from C . pasteurianum. The midpoint potential for the conversion of acetylene to ethylene is -0.465 V for the nitrogenase from Chromatium (133a). When the two proteins are mixed together, the resultant E P R spectrum appears to be very close to the sum of the two component spectra. Addition of MgATP produces two effects: 1. The azoferredoxin changes to the MgATP form very rapidly (msec) (134). This should be contrasted with the behavior of free azoferredoxin

-

130. R. L. Martin and A. H. White, Znorg. Chem. 6, 712 (1967). 131. B. E . Smith and G. Lang, BJ 137, 169 (1974). 132. T. Ljones, BBA 321, 103 (1973). 133. S.L. Albrecht and M. C. W. Evans, BBRC 55, 1009 (1973). 133a. M. C. W. Evans and S. L. Albrecht, BBRC 61, 1187 (1974). 134. L. E . Mortensen, W. Zumft, and G. Palmer, BBA 292, 422 (1973).

54

GRAHAM PALMER

described above, suggesting that the reactivity of this protein is enhanced by ca. three orders of magnitude in the nitrogenase complex. The amplitude of the azoferredoxin resonance then decreases by 50% (134). 2. The amplitude of the molybdoferredoxin E P R decreases substantially a t all g values. This effect is most readily seen a t low field (g = 4.38 and 3.7). The magnitude of the decrease depends on experimental conditions. If MgATP is added in the absence of a regenerating system, then the EPR amplitudes fall to ca. 50% of their original value and begin to increase again as ADP accumulates (MgADP is a potent competitive inhibitor). Further addition of MgATP initiates another cycle of this behavior. However, in the presence of creatine and creatine kinase the EPR amplitudes fall to about 10% of their starting value and stay down. The half-time for this process is about 60 msec which is smaller than the turnover time calculated for the experimental conditions (134).It is perhaps worth reiterating that nitrogenase is functioning in the steady state-there is a small but constant ( M ) concentration of the oxidant, H’, present. If turnover experiments are performed under conditions in which dithionite is limiting the end point consists of a state in which the molybdoferredoxin E P R is regenerated while the azoferredoxin E P R has almost disappeared ( l M , l S 4 ) ; subsequent addition of dithionite restores the steady-state levels. The interpretation of these data requires making some assumptions concerning the meaning of the disappearance of an EPR resonance since both oxidation and further reduction can eliminate the EPR of a paramagnetic species. This point was stressed early in this work (134) and it was appreciated that both oxidative and reductive mechanisms could be invoked. For concreteness the ramifications of an oxidative pathway were explored (13.4) but no judgment was made in favor of either possibility. Subsequently (I%?), however, it was asserted that a specific mechanistic pathway had been promulgated, in fact a “strawman” was created (123). The best interpretation of the available data is for a reductive pathway, as follows (MgATP is abbreviated to ATP) :

+

Azo Fd e Azo FdAZOFd2 ATP [AZO(ATP)z][Azo(ATPz]- MoFd AZO MoFd2 ADP 2 Pi [Azo(ATP)~]- MoFdAZO M0Fd’2 ADP 2 Pi MoFd*- 2 H+ MoFd H2

+ +

+

~

+

+ + +

+ + +

+ +

(la) (1b) (2) (3)

(4)

Several comments are needed to complement these equations. (1) It seems most likely that all of these discrete reactions occur in the nitroge-

1.

IRON-SULFUR

PROTEINS

55

nase complex; (2) reactions ( l a ) and ( l b ) might be inverted with electron transfer subsequent to ATP binding; (3) the paramagnetic species discussed above are Azo Fd, (Azo (ATP),)- and MoFd-; (4) MoFd and AoFd are physiologically oxidized species and MoFd2- is a physiologically super-reduced species; (5) since the rate of reduction of all acceptors (N2, H+ acetylene, etc.) is the same on a per electron basis, the rate-limiting step is probably Eq. (4); and (6) since MoFd*- is substantially depressed in the steady state but returns when dithionite is consumed then reaction (2) is significantly faster than the reaction ( 3 ) . Thus, reaction (2) > reaction (3) 2 reaction (4), and in the steady-state MoFd is in the form MoFd2-. Mossbauer observations made on molybdoferredoxin in the steady state indicate that the species Mo (giving the g = 4.27 E P R ) is further reduced to M,; M, and M, have Mossbauer parameters analogous to oxidized and reduced HiPiP, respectively (131). It thus seems that the scheme depicted above is a good first approximation to the reaction pathway. Aspects of this scheme which have been glossed over include (a) mechanism of reduction of N, (as opposed to H+), (b) the relevance of complex formation, and (c) the structure and function of the several different iron-sulfur clusters present in this system. From titration experiments with a variety of oxidants, Walker and Mortensen (135) determined that the conversion of Azo- to Azo required two electrons per 55,000 while oxidation of Moly- to Moly required two electrons per 110,000 molecular weight. I n a second experiment (136) comparing the amount of oxidant consumed by molybdoferredoxin in the resting and steady states in the presence of identical concentrations of dithionite i t was deduced that an additional two electrons per 110,000 were involved in the reduction of Moly- to Moly2-. Thus a 1:1 complex of the two proteins could provide six electrons required for N2 reduction. At the high protein concentrations employed in most analytical methods molybdoferredoxin has a molecular weight of 220,000; that is, it dimerises. Eady, using the proteins from Klebsiella, has provided good physical data to slow the existence of a 1:l complex between this dimer and azoferredoxin (55,000) ( 1 3 7 ) . His activity measurement, however, can be used to support this 1: 1 ratio when molybdoferredoxin is varied a t fixed azoferredoxin and 1:2 when azoferredoxin is varied a t fixed molybdoferredoxin (137). Values of 1:l (138) and 1:2 (139) have also been M. Walker and L. Mortensen, BBRC 54, 669 (1973). M. Walker and L. Mortensen, JBC 249,6356 (1974). R. R. Eadg, BJ 135,531 (1973). L. E. Mortensen, W. G . Zumft, T. C. Huang, and G . Palmer, Biochem. Sac. Trans. 1, 33 (1973). 139. M.-Y. Tso, T. Ljones, and R. H. Burris, BBA 261,600 (1972). 135. 136. 137. 138.

56

GRAHAM PALMER

obtained for the ratio of proteins present in the nitrogenase complex from C . pasteurianum. Presuming that the electron stoichiometries described above are correct, then the most attractive alternatives are a unitary complex between azoferredoxin (55,000) and molybdoferredoxin (110,000) or a dimer of this complex (equivalent to the 1:2 complex of the preceding text). The 1 : 1 complex described above would contain 10 electrons which has no obvious value. This argument, however, places considkrable credence in the accuracy of the electron-balance experiments and should be taken cum granis salis.

B. XANTHTNE OXIDASE Xanthine oxidase is unquestionably the best characterized of the iron-sulfur enzymes, and a recent series of papers has made a major contribution to the understanding of the principles a t play in the functioning of redox enzymes containing several prosthetic groups (140-14s). This work is reviewed in the article by Bray in this volume.

C. MITOCHONDRIAL IRON-SULFUR PROTEINS Recent data have shown that mitochondria contain a large number of iron-sulfur centers. As yet these species are identified by their E P R characteristics and their precise function has not been established. The best sources of information are Orme-Johnson et al. (143,144) and Ohnishi (146).

140. D. Edmondson, D. Ballou, A. Van Heuvelen, G. Palmer, and V. Massey, JBC 248, 6135-6144 (1973). 141. J. S. Olsen, D. Ballou, G. Palmer, and V. Masaey, JBC 249, 4350 (1974). 142. J. S. Olsen, D. Ballou, G. Palmer, and V. Mawey, JBC 249, 4323 (1974). 143. N. R. Orme-Johnson, R. E. Hansen, and H. Beinert, JBC 249, 1922 (1974). 144. N. R. OrmeJohnson, R. E. Hansen, and H. Beinert, JBC 249, 1928 (1974). 145. T.Ohnishi, BBA 301, 105 (1973).

Flawodoxins and Electron-Transferring Flavoproteins STEPHEN G. MAYHEW

MARTHA L. LUDWIG

I. Introduction. . . . . . . . . . . . . . . . 11. Flavodoxins . . . . . . . . . . . . . . . . A. Background . . . . . . . . . . . . . . . B. Structures . . . . . . . . . . . . . . . C. Flavin-Protein Interactions: Chemical and Physical Studies Solution . . . . . . . . . . . . . . . 1). Spectroscopic Properties . . . . . . . . . . . E. Oxidation-Reduction Potentials . . . . . . . . . F. Reactivity . . . . . . . . . . . . . . . 111. Electron-Transferring Flavoprotein . . . . . . . . . A. Introduction. . . . . . . . . . . . . . . B. Molecular Properties . . . . . . . . . . . . C. Catalytic Properties . . . . . . . . . . . .

. .

. .

57 58 68 66

in

. .

. . . . . .

82 88 98 102 109 109 111 116

1. Introduction

This chapter discusses two classes of flavoproteins which function solely to mediate electron transfer between the prosthetic groups of other proteins. Beinert ( 1 ) and co-workers discovered the first flavoprotein of this type, a soluble FAD protein obtained from mitochondria which couples the oxidation of acyl-CoA dehydrogenases to the reduction of components of the terminal electron transfer chain. According to its function the protein was termed “electron-transferring flavoprotein” (ETF). 1. H. Beinert, “The Enzymes,” 2nd ed., Vol. 7, p. 467, 1963. 57

58

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

More recently, a different class of flavoprotein carriers has been isolated from microorganisms. Because of their functional interchangeability with the ferredoxins, these proteins have been called flavodoxins. The major portion of this chapter is devoted to the flavodoxins because they are the first flavoproteins for which three-dimensional structures have been determined.

II. Flavodoxinr

A. BACKGROUND 1. Discovery, Nomenclature, and Distribution In the decade that followed the discovery of ferredoxin (2,S),a further class of microbial proteins which transfer electrons at low potential was recognized. These proteins are also small and acidic and in many reactions they substitute efficiently for ferredoxin. However, in contrast to the ferredoxins which contain iron and acid-labile sulfide, proteins in the second group utilize a molecule of flavin mononucleotide as their redoxactive component. Smillie (4,6) purified the first flavoprotein of this kind from extracts of the blue-green alga Anacystis niduhns and termed the protein “phytoflavin.” Shortly afterward, an FMN protein with similar catalytic properties was isolated from a strictly anaerobic bacterium, Clostridiunz pasteurianum, and crystallized by Knight and co-workers (6-8). To indicate the functional similarity with ferredoxin, Knight et al. (6) proposed the term “flavodoxin” for their FMN protein. This term has been adopted for similar flavoproteins that were subsequently isolated from a variety of microorganisms (9-18),and it has been extended by some authors 2. L. E. Mortenson, R. C. Valentine, and J. E. Carnahan, BBRC 7,448 (1962). 3. K. Tagawa and D. I. Arnon, Nature (London) 195, 537 (1962). 4. R. M. Smillie, Plant Physiol. 38, 28 (1963). 5. R. M. Smillie, BBRC 20, 621 (1965). 6. E. Knight, Jr., A. J. D’Eustachio, and R. W. F. Hardy, BBA 113, 626 (1966). 7. E. Knight, Jr., and R. W. F. Hardy, JBC 241,2752 (1966). 8. E. Knight, Jr., and R. W. F. Hardy, JBC 242, 1370 (1967). 9. J. LeGall and E. C. Hatchikian, C. R. Acad. Sci., Ser. D 264, 2580 (1967). 10. M. Dubourdieu, J. LeCall, and F. Leterrier, C. R . Acad. Sci., Ser. D 267, 1653 (1968). 11. M. Dubourdieu and J. LeGall, BBRC 38, 965 (1970). 12. S. G. Mayhew and V. Massey, JBC 244, 794 (1969). 13. S. G. Mayhew, BBA 235,276 (1971). 14. M. A. Cusanovich and D. E. Edmondson, BBRC 45, 327 (1971)

2.

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59

to include not only Smillie’s phytoflavin but also a protein from Azotobacter vinelandii (19,200) , which in earlier publications had been called “free radical flavoprotein” (21,22), “Shethna flavoprotein” (23), or “azotoflavin” (24,25). All of these flavoproteins catalyze the transfer of electrons to and from other proteins, and as they are also broadly similar in their chemical and physical properties, it is appropriate that they should be described by a single term. In this article the general name flavodoxin has been retained because it has been used most often in the literature. As pointed out by Beinert (26) however, this nomenclature is imprecise by comparison with the corresponding term “ferredoxin” for the iron-sulfur proteins. Organisms from which flavoproteins of the flavodoxin type have been isolated include several strictly anerobic bacteria, representatives from the obligately aerobic [ A . vineEandii (19,222)1, facultatively anaerobic [Escherichia coli ( 16 ) ] , and photosynthetic [ Rhodospirillum rubrum (14) ] groups of bacteria, blue-green algae [e.g., Anacystic nidulans (46) and Synechococcus lividus (17)] and a eukaryotic green alga [Chlorella fusca ( 1 6 ) ] (Table IV). They have not yet been found in higher animals and plants. 2. Chemical Composition, Moleculay Weight, and Purification

Flavodoxins contain one equivalent of FMN. Flavin is their only known prosthetic group (5-18) , and they lack transition metals, in particular the iron-sulfur chromophore of the ferredoxins. A tabulation of the amino acid compositions of 12 flavodoxins (27) discloses some general similarities. Acidic amino acids always predominate over basic residues; nine 15. W. G. Zumft and H. Spiller, B B R C 45, 112 (1971). 16. H. Vetter, Jr., and J. Knappe, Hoppe-Seyler’s 2. Physiol. Chem. 352, 433 (1971). 17. H. L. Crespi, U. Smith, L. Gajda, T. Tisue, and R. M. Ameraal, B B A 256, 611 (1972). 18. H. L. Crespi, J. R. Norris, and J. J. Katz, Nature (London), New Biol. 236, 178 (1972). 19. B. van Lin and H. Bothe, Arch. Mikrobiol. 82, 155 (1972). 20. H. Bothe and B. Falkenberg, 2.Naturforsch. B 27, 1090 (1972). 21. Y. I. Shethna, P. W. Wilson, and H. Beinert, B B A 113, 225 (1966). 22. J. W. Hinkson and W. A. Bulen, JBC 242, 3345 (1967). 23. D. E. Edmondson and G. Tollin, Biochemistry 10, 113 (1971). 24. J. R. Benemann, D. C. Yoch, R. C. Valentine, and D. I. Arnon, Proc. Nut. Acud. Sci. U . S.64, 1079 (1969). 25. D. C. Yoch, J. R. Benemann, R. C. Valentine, and D. I. Arnon, Proc. Nut. Acud. Sci. U . S. 84, 1404 (1969). 26. H. Beinert, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 207. Univ. Park Press, Baltimore, Maryland, 1971. 27. J. L. Fox, S. S. Smith, and J. R . Brown, 2.Naturforsch. B 27, 1096 (1972).

60

STEPHEN G. MAYHEW AND MARTHA

L.

LUDWIG

flavodoxins either lack histidine altogether or contain only a single residue. Yet flavodoxins from differentstrains of Desulfovibrio or Clostridium have distinctive compositions (If ,IS). For the flavodoxins surveyed (27), the cysteine content varies from one to five residues and tryptophan from one to six residues. The polypeptide chain lengths, based on amino acid compositions, lie between approximately 120 and 220 residues, corresponding to apoprotein molecular weights between 14,500 and 23,000. According to size, the flavodoxins seem to fall into two groups. One category has molecular weights of 14,500to 17,000, and the other, 20,000 to 23,000. Unless otherwise noted, the molecular weights given in Table IV (Section I1,E) are calculated from compositions. Determinations by ultracentrifugation or gel filtration have occasionally produced somewhat different values. The several published procedures for the purification of flavodoxins exploit their low isoelectric points, which result in retention of these proteins by DEAE-cellulose under conditions where many other proteins are eluted (5,7,12,14,22). Cell-free extracts are often applied directly to DEAE columns. Development with salt gradients results in substantial purification. Ammonium sulfate fractionation and a second DEAE chromatography, followed by gel filtration, provides a highly purified preparation. Several flavodoxins crystallize readily from solutions of ammonium sulfate (7,12,IS,28,29). All are very stable and can be stored for long periods in frozen solution or as crystals a t 4O.

3. Function Flavodoxins do not react directly with small molecules such as the pyridine nucleotides, and their only known biochemical “substrates” are other redox proteins. Nevertheless, the number of reactions known to utilize low potential carriers is impressive; e.g., a recent review (SO) lists 18 ferredoxin-dependent enzymes of fermentative bacteria. Replacement of ferredoxin by flavodoxin has not been attempted in every one of the ferredoxin-requiring reactions, and there are a few systems in which flavodoxins seem unable to fill the role of ferredoxins (3I,S2). However, flavodoxins prove to be efficient carriers in numerous reactions. Smillie (5)was the first to demonstrate that a flavodoxin (phytoflavin from A . nidulam) could replace ferredoxin in the light-dependent reduc28. M. L. Ludwig, R. D. Andersen, S. G. Maphew, and V. Massey, JBC 244, 6047 (1969). 29. K.D. Watenpaugh, L. C. Sieker, L. H. Jensen, J. LeGall, and M. Dubourdieu. ?roc. Nat. Acad. Sci. U . S. 69, 3185 (1972). 30. D. C. Yoch and R. C. Valentine, Annu. Rev. Microbiol. 26, 139 (1972). 31. U. Gehring and D. I. Arnon, JBC 247,6963 (1972). 32. L.L. Barton and H. D. Peck, Bacterial. Proc. p. 134 (1970).

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tion of NADP' by plant chloroplasts. Knight and Hardy (7,8) subsequently showed that flavodoxin from C. pasteurianum is likewise a mediator in the photosynthetic system. I n this chain of reactions, ferredoxin or flavodoxin transfers electrons to the flavoenzyme ferredoxin-NADP+reductase (2,5,33-35). In addition, flavodoxins substitute for bacterial ferredoxins in the phosphoroclastic oxidation of pyruvate (7,8,IZ,gO). Hf

+ CH&OCOO- + HP04'-

+ CH3COOPOs'-

+ COz + Hz

(1) The clastic system in extracts of C. pasteurianum and other anaerobic bacteria includes the enzymes pyruvate dehydrogenase, phosphotransacetylase, and hydrogenase. It utilizes coenzyme A as cofactor; the low potential electron carrier is required to mediate the oxidation of pyruvate dehydrogenase and the reduction of hydrogenase. Reducing equivalents made available by the oxidation of pyruvate can be transferred by ferredoxin or flavodoxin not only to hydrogenase but also to other enzymes that reduce a variety of compounds, including molecular nitrogen (7,19,S6) and pyridine nucleotides (8). Reduced ferredoxin also participates in the reversal of the first step of the clastic reaction, namely, the formation of pyruvate from CO, and acetyl-CoA, catalyzed by pyruvate synthase in various anaerobes and photosynthetic bacteria (S7). Peptostreptococcus elsdenii flavodoxin is able to substitute for ferredoxin in the analogous fixation of CO, into butyrate (38). The dissimilatory pathway of sulfate reduction in Desulfovibrio species relies on ferredoxin or flavodoxin to mediate transfer of electrons between hydrogen and sulfite. Formation of H,S from sulfite proceeds in several steps and the precise role of flavodoxin (or ferredoxin) has not been conclusively established ; the overall reaction is also dependent on cytochrome CRI

(99,401.

I n many cases the requirement for an electron carrier has been established using crude extracts or partially purified enzymes, and for this reason the direct interaction of flavodoxin or ferredoxin with particular A. Son Pietro and H. M. Lang, JBC 231, 211 (1958). M. Shin, K . Tagawa, and D. I. Arnon, Biochem. Z. 338, 84 (1963). M . Shin and D. I. Arnon, JBC 240, 1405 (1965). M. G. Yates, FEBS (Fed. Eur. Biochem. Soc.) Lett. 27, 63 (1972). R . Bachofen, B. B. Buchanan, and D. I. Amon, Proc. Nat. Acad. Sci. U . S . 51, 690 (1964). 38. M. J. Allison and J. L. Peel, BJ 121,431 (1971). 39. E. C. Hatchikian, J. LeGalI, M. Bruechi, and M. Dubourdieu, BBA 258, 701 (1972). 40. K. hie, K . Kobayashi, M . Kobayashi, and M. Ishimoto, J. Biochem. ( T o k y o ) 73, 353 (1973). 33. 34. 35. 36. 37.

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STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

redox enzymes has not always been rigorously demonstrated. An exception is the reaction of carriers with ferredoxin-NADP+-reductase, which has been investigated in considerable detail (Section II,F,4). Purified preparations of nitrogenase have been shown to accept electrons directly from Azotobacter chroococcum flavodoxin (367, but it is not clear whether the site of transfer is the Mo or the nonheme iron moiety of nitrogenase. It has been proposed that a nonheme iron center of the bacterial pyruvate dehydrogenase is oxidized by electron carriers (41) . The relative activities of flavodoxins and ferredoxins as electron carriers have been determined in both plant and microbial systems, with results which indicate that transfer rates vary somewhat according to the source of the carrier; for example, C. pasteurianum and A . niduluns flavodoxins are reported to be twice as active, on a molar basis, as bacterial and A . nidulans ferredoxins, respectively, in stimulating production of NADPPH by washed chloroplasts (5,8). On the other hand, the activity of Chlorella fusca flavodoxin is less than that of Chlorella ferredoxin in the same assay (16). Flavodoxins are generally less efficient than ferredoxins in the phosphoroclastic reaction, although at saturating levels of the carriers the rates become approximately equal (9-17). The interchangeability of carriers with quite different structures and chromophores suggests a lack of recognition in the electron transfer reactions, yet the existence of tight complexes between carriers and ferredoxin-NADP+-reductase has been demonstrated (42-46). Flavodoxin from A . vinelandii differs from other flavodoxins in showing abnormally low activity in several ferredoxin-dependent reactions. As a result some time elapsed before a catalytic function could be ascribed to this protein (21,22). I n 1969, Benemann and co-workers (24) discovered that it is weakly active as an electron carrier between spinach chloroplasts and nitrogenase of A . vinelandii. The activity in nitrogen fixation was confirmed by van Lin and Bothe (19) who showed further that, contrary to earlier indications (14,22,24),this flavodoxin also substitutes for ferredoxin in the photosynthetic reduction of NADP' by plant chloroplasts. The critical difference between the experiments of van Lin and Bothe (19) and previous negative results was the use of an anaerobic gas phase. However, even under these conditions, the catalytic efficiency of this flavodoxin is low, and the maximum rate with saturating concentrations is only about half of the maximum rate observed with fer41. K.Uyeda and J. C. Rahinowitz, JBC 246,3111 (1971). 42. G. P. Foust, S. G. Mayhew, and V. Massey, JBC 244, 964 (1969). 43. N. Nelson and J. Neumann, BBRC 30,142 (1968). 44. M.Shin and A. San Pietro, BBRC 33, 38 (1968). 45. M.Shin, BBA 292, 13 (1973).

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redoxin and flavodoxin from A . nidulans. Bothe and Falkenberg (20) subsequently showed that A . vinelandii flavodoxin can function in the phosphoroclastic oxidation of pyruvate catalyzed by extracts of C . pasteurianum. The principal metabolic role of A . vinelandii flavodoxin still remains to be established. Enzyme assays based on the activity of flavodoxin in the photosynthetic reduction of NADP’ (46), the production of hydrogen from dithionite in the presence of hydrogenase ( 7 ), and the phosphoroclastic oxidation of pyruvate (12) have been described. These are unsatisfactory in several respects: first, they are insensitive; second, they depend on crude and unstable preparations of other fractions; third, they do not distinguish between flavodoxin and ferredoxin, and consequently flavodoxin can be positively identified only after it has been obtained in pure form. More recently, in the authors’ laboratories, flavodoxin has been assayed by its ability to couple NADPH oxidation to cytochrome c reduction in the presence of purified ferredoxin-NADP+-reductase ( 4 7 ) . This procedure still suffers from the lack of discrimination between flavodoxin and ferredoxin. Enzymic assay is of only limited use during routine purification, and purity can usually be more reliably estimated from the absorption spectrum (Section II,D,l) . However, estimates of the quantities of flavodoxin and ferredoxin in crude extracts have necessitated their preliminary separation, and are therefore uncertain, particularly in the case of the more unstable ferredoxins. The lack of a suitable catalytic assay might be circumvented by the use of immunochemical techniques. It has been found that antibodies to flavodoxins from P. elsdenii and Clostridium M P do not cross react with ferredoxins from these sources (48)*

4. Regulation by Iron I n certain microorganisms the synthesis of flavodoxin occurs only during growth in iron-poor media. This regulation by iron is the first of its kind observed with a flavoprotein, though similar effects on the synthesis The effect of iron of flavin have been known for about 30 years (49,50). on C. pasteurianum was noted by Knight and Hardy ( 7 ) , who showed that little if any ferredoxin is produced by this organism under iron-deficient conditions. Knight and Hardy ( 7 ) concluded that the flavoprotein 46. R. M. Smillie and B. Entsch, “Methods in Enzymology,” Vol. 23, Part A, p. 504, 1971. 47. M. Shin, “Methods in Enzymology,” Vol. 23, Part A, p. 440, 1971. 48. H. J. Somerville and S. G. Mayhew, unpublished. 49. R. J. Hickey, Arch. Biochem. 8, 439 (1945). 50. A. L. Demain, Annu. R e v . Microbial. 26, 369 (1972).

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STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

is synthesized as a replacement for ferredoxin when the metal is in short supply. Iron has a similarly pronounced effect on the synthesis of ferredoxin and flavodoxin in P , elsdenii. The use of immunochemical techniques to estimate flavodoxin in cell-free extracts of P. elsdenii has permitted the conclusion that iron-rich cells do not contain flavodoxin or its apoprotein, and that iron deficiency brings about de nouo synthesis ( 4 8 ) .I n other organisms control by iron is less dramatic, and it is more difficult to obtain cells either free of ferredoxin in iron-poor media (51) or of flavodoxin in iron-rich media (39,40). In still a third group of organisms, E. coli and A . uinelandii, flavodoxin synthesis appears to be independent of iron (16‘,19,24). It is not clear, however, whether the ratio of ferredoxin to flavodoxin in these two organisms is sensitive to iron, nor is it known whether flavodoxin and ferredoxins from A . vinelandii, and other organisms in which proteins of both types are synthesized simultaneously, share the same functions. The picture is further complicated in the case of A . uinelandii by the presence of more than one type of ferredoxin (19,25,62). The iron concentration which favors flavodoxin synthesis varies with the organism, but is in the range of 0.01-0.5 pg/ml. The iron restriction markedly limits the total growth of certain organisms [ e.g., Clostridium MP (13)] and in such cases it is clear that flavodoxin synthesis depends on a true iron deficiency. The total growth of other organisms is much less affected (e.g., 7,12).However, batch cultures have been used in investigations on the relationship between iron levels and flavodoxin synthesis, and it is not known whether the protein is synthesized throughout growth or only when the available iron is depleted. There is surprisingly little information about the overall effects of iron deficiency on organisms which require low concentrations of iron for the production of flavodoxin. Clostridium pasteurianum is less rodlike, more ovoid, and almost white in iron-deficient media, and anaerobic cells of P. elsdenii from iron-poor media are gray in contrast to cells from ironsufficient media, which, depending on the excess of iron, are green-brown or black ( 5 3 ) . Cells of C . pasteurianum which contain flavodoxin still catalyze the reduction of molecular nitrogen (79, and in P. elsdenii the overall fermentation of lactate to fatty acids, hydrogen, and carbon dioxide (54) is not appreciably influenced by iron deprivation ( 5 3 ) .It is pos51. H. Bothe, P. Hemmerich, and H. Sund, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 211. Univ. Park Press, Baltimore, Maryland, 1971. 52. Y. I. Shethna, D. V. Dervartanian, and H. Beinert, BBRC 31, 862 (1968). 53. S. G. Mayhew, unpublished. 54. S. R. Elsden, B. E. Velcani, F. M. C. Gilchrist, and D. Lewis, J. Bacterial 72, 681 (1956).

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FLAVOPROTEINS

65

sible therefore that iron proteins such as nitrogenase (55), hydrogenase (56),and pyruvate dehydrogenase (41) are not drastically affected under these conditions, but this point has not been tested with the purified proteins. However, there is evidence that some iron proteins are more critical than others ; for example, several iron-deficient organisms contain the iron protein rubredoxin (12,lS), and Chlorella fusca from iron-poor medium still contains cytochromes and nitrite reductase (15). I n view of the profound effects of iron on flavin synthesis by some microorganisms (49,50),it is interesting that the total flavin content of P. elsdenii (57) is not appreciably affected by mild iron deficiency; F M N is increased, but FAD is correspondingly lower (53). 5. Some Properties of the Bound Flavin Mononucleotide In all the flavodoxins the oxidation-reduction potentials of bound F M N differ significantly from those for the free prosthetic group. The two oneelectron steps have distinct potentials with the consequence that the semiquinone form can be obtained in essentially quantitative yields under appropriate conditions. Redox potentials for the semiquinone-fully reduced couple are in the range typical of ferredoxins (Table IV, Section II,E), and are the lowest known for any flavoprotein. Potentials for the oxidized-semiquinone couple are frequently higher than for free FMN. From these data it seemed reasonable to postulate that in vivo the flavodoxins may act as one-electron carriers, shuttling between the fully reduced and semiquinone states (19,58), and there is some evidence to support this suggestion ( 3 6 ) . The prosthetic group is bound tightly but not covalently by apoflavodoxins. The measured association constants are of the order of lo8 or greater, but the holoproteins are reversibly dissociated by a number of procedures used to prepare other apoflavoproteins, such as low p H or dialysis against concentrated KBr (5,59-61). A limited number of modified flavins can be bound instead of FMN, with the specificity depending upon the species from which the flavodoxin is derived (59,61) (Section II,C,3). Spectroscopic differences among the flavodoxins suggest that the environment of the flavin chromophore varies with the species. It has been proposed that the flavodoxins be classified into two spectral groups, one that resembles C. pasteurianum flavodoxin, and a second group, in55. H. Dalton and L. E. Morteneon, Bacterial. Rev. 36, 231 (1972). 56. G. Nakos and L. E. Mortenson, Biochemistry 10,2442 (1971). 57. J. L. Peel, BJ 69, 403 (1958). 58. S. G. Mayhew, G. P. Foust, and V. Massey, JBC 244,803 (1969). 59. S. G. Mayhew, BBA 235, 289 (1971). 60. J. W. Hinkson, Biochemistry 7, 2666 (1968). 61. D. E. Edrnondson and G. Tollin, Biochemistry 10, 124 (1971).

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STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

cluding D. vulgaris flavodoxin, that more closely resembles flavodoxin from R. rubrum (62). This division does not coincide with that based on molecular weight. In many respects the flavodoxins resemble the larger flavoprotein dehydrogenases (63).Both classes of proteins form a blue neutral semiquinone rather than the red anion; they resist substitution by sulfite a t N-5; and they are reoxidized by oxygen in two steps with concomitant formation of the superoxide radical.

B . STRUCTURES 1. Introduction At the present time the detailed three-dimensional structures of two flavodoxins, those from D. vulgaris (29,64) and Clostridium MP (65,66), are known, along with the corresponding amino acid sequences (67,68). For Clostridium M P flavodoxin, electron density maps of not only the oxidized (66) but also the semiquinone and reduced states (65,69) have been computed at high resolution. In addition, the complete sequence of P. elsdenii flavodoxin (70) and partial sequences of Clostridium pasteurianum (27,7l) flavodoxin have been reported. From the sequences alone, it appears that one region near the N-terminus (part of the FMN binding site) is highly conserved during evolution, but farther along the chain homologies are more difficult to discern by inspection. Nevertheless, the three-dimensional folding of D. vulgaris and Clostridium hilP flavodoxins clearly demonstrates structural homology. The surprising conclusion from 62. J. A. D’Anna, Jr. and G. Tollin, Biochemistry 11, 1073 (1972). 63. 8. Massey, F. Muller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G. Howell, S. G. Mayhew, R. G. Matthews, and G. P. Foust, JBC 244, 3999 (1969). 64. K. D. Watenpaugh, L. C. Sieker, and L. H. Jensen, Proc. Nut. Acad. Sci. U. S. 70, 3857 (1973). 65. R. D.Andersen, P. A. Apgar, R. M. Burnett, G. D. Darling, M. E. LeQuesne, S. G. Mayhew, and M. L. Ludwig, Proc. Nut. Acad. Sci. U . S. 89, 3189 (1972). 66. R. M. Burnett, G. D. Darling, D. S. Kendall, M. E. LeQuesne, S. G. Mayhew, W. W. Smith, and M. L. Ludwig, JBC 249,4383 (1974). 67. M. Dubourdieu, J. LeGall, and J. L. Fox, BBRC 52, 1418 (1973). 68. M. Tanaka, M. Haniu, K. T. Yasunobu, and S. G. Mayhew, JBC 249, 4393 (1974). 69. M. L. Ludwig, R. M. Burnett, G. D. Darling, S. R. Jordan, D. S. Xendall, and W. W. Smith, in “Structure and Conformation of Nucleic Acids and ProteinNucleic Acid Interactions” (M. Sundaralingam and s. T. Rao, eds.) (in press). 70. M. Tanaka, M. Haniu, K. T. Yasunobu, S. G. Mayhew, and V. Massey, JBC 248, 4354 (1973); 249, 4397 (1974). 71. M. Tanaka, M. Haniu, G. Matsueda, K. T. Yasunobu, S. G. Mayhew, and V. Massey, Biochemistry 10, 3041 (1971).

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67

comparisons of the two structures is that many of the variations between species occur a t the active center, in the vicinity of the isoalloxazine ring. Structural differences in the neighborhood of the flavin ring are compatible with the distinctive spectral properties of the two proteins (Section II,D,l). 2. Determination and Comparison of Chemical Sequences

Total sequence determination has so far been confined t o the smaller flavodoxins with chain lengths less than 150 residues. These proteins have presented no extraordinary problems, yielding tractable peptides after cleavage with CNBr, trypsin, chymotrypsin, and thermolysin (67,68,7O). Sequences as long as 52 residues have been determined by automated Edman degradation (68). The primary structures of Clostridium M P and D. vulgaris flavodoxins, comprising 138 and 148 residues, respectively, are displayed in Fig. 1. Some of the residues determined to be equivalent (Le., structurally homologous) by comparison of the three-dimensional models are also shown. The chain folding in the two flavodoxins (Section II,B,Q) has been compared after applying to one model the rigid-body rotations and translations required to minimize the squares of the distances between related &-carbon atoms (72). (Nonequivalent atoms are eliminated as the calculation proceeds.) Preliminary calculations have established the general similarity of the helical and sheet domains in the two structures ( 6 5 ) . Identification of all residues which occupy equivalent positions in the three-dimensional structures is incomplete a t the time of writing, but homologies based on the correspondence of C, positions have been established for the p sheet and the N-terminal helix (Figs. 1-3). The results indicate the regions in which the extra 10 residues are inserted in the D. vulgaris chain. Three additions occur a t the N-terminus, one more somewhere between residues 38 and 47, two between 57 and 77, three between 89 and 100, and one beyond 123. (Numbers refer to the Clostridium MP sequence.) Superposition of the drawings in Fig. 2 reveals several areas in which the matching of the structures is imperfect, notably in the vicinity of Clostridium MP residues 40, 58, and 90, and along much of the helix surrounding residue 70. The primary sequences of the flavodoxins have been analyzed to determine those residues which are homologous in the evolutionary sense (67,73). Since the C-terminal portions of the flavodoxin chains, from approximately residue 90 (Clostridium M P numbering) onward, are highly variable, alignments have relied in part on the principle of mini72. s. T. Rao and M. G. Rossmann, J M B 76, 241 (1973). 73. W. M. Fitch and K. T. Yasunobu, private communication.

68

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG 1 10 Met Lys Ile Val T y r T r p Ser Gly T h r Gly Asn T h r Glu Lys Met Ala Glu Leu Ile Ala 21 30 40 Lye Gly Ile Ile Glu Ser Gly LYEAsp Val Asn T h r Ile Asn Val Ser Asp Val A m Ile 41 50 60 Asp Glu Leu Leu Asn Glu Asp Ile Leu Ile Leu Gly CyS S e r Ala Met Gly Asp Glu Val

61

80

70

Leu Glu Glu Ser Glu Phe Glu P r o Phe Ile Glu Glu Ile S e r T h r Lys Ile Ser Gly Lys 81 go 100 Lys Val Ala Leu Phe Gly Ser Tyr Gly T r p Gly Asp Gly Lys T r p Met Arg Asp Phe Glu 101 110 Glu Arg Met Asx Gly T y r Gly CyS Val Val Val Glu T h r P r o Leu Ile Val Gln

120

+ Glu

121 130 138 P r o Asp Glu A h Glu Gln Asp CyS Ile Glu Phe Gly Lys Lys Ile Ala Am Ile

(a)

-

I0 CLDST.RlDlUM

Up 1 I I (21

0. VULGARIS C. PASTEURIANUM P. ELSDENll

-8-

-

20

30

M*KVNIIYWSGTGNTEAMAKLIAEGAQEKGAOVKLLNV M*+VEIVYWSGTGNTEAMANEIEAAVKAAGADVESVRF

m

60

x)

-8CLDST!tlPlUM

MP 1 I 1 ( 21

D. VULGARIS C. PASTEURIANUM P. ELSDENII CLOST~IOIUM MP II

I

(21 D. VULGARIS C. PASTEURIANUM P. ELSOENII

D. VULGARIS

YP 1 II ( 21

ED

-a-

VNIDELLNE*DILILGCSAMGDEVLEESEFfPFlEElSTK (761 V DI L I L G C 5 AMGI V E A G G L F E G F D L V L L G C S T W G D D ~ OI * * L Z B B F I P L F D S L 1781 DVVAFGSPSMG DVILLGCPAMG 90 100 I10 I20

**

-B-

-

-a-

I S G K K V A L F G S Y G W G O* ISGKKVALFGSYG

* G K W M R D F E E R M N G Y G C V il091 E E R M N G Y GCV

Z Z T G A Z G R K V A C F G C G B S S Y E Y F C G A V D A I E E K L K N L G A Z 11181 GKKEGAFXXXX GKKVGLFGSYG

8 CLOST.RlDlUM

40

-8-

M*K**IVYWSGTGNTEKMAELIAUGI I E S G K D V N T I N V S D I371 MKIIVYWSGTGNTEKMAELIAKGIIESGKDVIMTINVSD M P K A L I V Y G S T T G N T E Y T A E T I A R E L A ~ A G Y E V D S R D A A1401 S

I30

-

I40

ISD

-a-

VVETPL*IVONEPOEAEQD.CI EFGKKlANl V V E T P L I V O W E P DIE1 I VLBGLR I DGDPRAARBB IVGWAHDVRGA I

(1381 (1481

(b)

FIQ.1. (a) The sequence of Clostridium M P flavodoxin. Sequences forming the FMK binding site are underlined; that does not imply that every underlined residue provides contacts with the prosthetic group (cf. Figs. 4 and 5 ) . (b) Comparison of the chemical sequences of four flavodoxins, employing a numbering scheme which allowa for deletions (*) in each chain. The actual residue number a t the end of each line is given in parentheses. The sequence of C. pasteuriunum flavodoxin is unknown between position 93 and the seven residues at the C-terminus. Lines (1) and (3) through ( 5 ) display the alignments which minimize the number of mutations required to relate the four sequences (75). Lines (2) and (3) show some of the structurally equivalent a-carbons, determined by rigid-body fitting of the coordinates. When the structures are superposed, each C, in line (2) is less than 2.0 A from the corresponding atom in the D. vulgaris structure, except for those few residues enclosed in parentheses. These latter C , atoms are in matching regions and are separated by just slightly more than 2.0 A. The average distance between the equivale'nt atoms shown is 1.1 A. A number of additional equivalent a-carbons have been omitted from line (2), pending comparison of the p-carbon positions. The location of the p structure and of helices in Clostridium M P flavodoxin is indicated by arrows. Residues constituting the F M N binding site of D. vulgaris flavodoxin are shown in Fig. 4b.

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING

FLAVOPROTEINS

69

mum mutation frequency ( 7 4 ) . At many positions, including much of the p sheet, the alignments determined from the structures are consistent with those expected from mutation of a “primordial” flavodoxin gene. The approximate location of gaps and insertions is in agreement. However, the homologies deduced by the two methods do not always coincide. The N-terminus provides a nice illustration of the discrepancies. Althmgh the two methionines presumably initiate both chains and are related according to genetic analysis, i t happens that deletions in the Clostridium MP chain can be accommodated in the structure by placing Met-1 a t the position occupied by the fourth residue of the D. vulgaris chain. Again, in the final strand of p sheet, genetic arguments suggest an insertion in the D. vulgaris chain a t position 127 of Fig. lb, following Leu-115 of Clostridium M P flavodoxin. There is no corresponding deviation of the superposed structures at this point in the p sheet; instead the chains remain in register for this and several succeeding residues. Correspondence of the two structures is poor for at least seven residues of the D. vulgaris chain, beginning with Ser-96, even though this sequence constitutes part of the active site and the residues may be derived from the same ancestor. The disparity between structural equivalences and the alignments predicted from genetic considerations recalls similar difficulties in assignment of homologies in cytochrome c ( 7 5 ) . Comparison of the sequences of C . pasteurianum and P. elsdenii flavodoxins with those of Clostridium M P and D. vulgaris flavodoxins demonstrates that the first three proteins are more closely related t o one another than to D. vulgaris flavodoxin (67,7’3).Only portions of the known sequences of C . pasteurianum and P. elsdenii flavodoxins have been selected for Fig. l b ; these correspond to the most highly conserved parts of the structure, where the relationships of the sequences are often evident by inspection. At present there is no information on the location of the extra residues in the flavodoxins of molecular weight near 20,000. It will be fascinating to see whether these longer chains contain additional elements of secondary structure (cf. Volume XI, Chapter 2). Evolutionary conservation of most of the residues a t positions 6 through 16 is obvious from Fig. lb. Several other positions, ie., Ala-19, Ile-22, Gly-30, Val-33, Asp-51, Gly-56, Gly-61, Gly-87, Lys-89, and Phe-93, are invariant in the portions of the four different flavodoxins shown in Fig. lb. The region from position 10 to 15, containing several hydroxyamino acids, constitutes part of the binding site for the phosphate portion of the flavin mononucleotide. However, many of the other active center residues of Clostridium M P and D . vulgaris flavodoxin differ. The 74. W. M. Fitch, J M B 49, 1 (1970). 75. R.E. Dickerson, J M B 57, 1 (1971).

70

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

relative arrangements of polypeptide and FMN in each molecule are described more explicitly in the following discussion of the three-dimensional structures. Although modification of cysteine residues is known to interfere with FMN binding in the four flavodoxins included in Fig. l b (8,13,40,69), each cysteine is replaceable. 3. Three-Dimensional Structures

a. Structure Determination. The pertinent crystallographic data for Clostridium M P and D. vulgaris flavodoxins are given in Table I. Despite their conformational similarities, the molecules of Clostridium M P and D. vulgaris flavodoxins pack quite differently in their respective unit cells, with the crystals of the D . vulgaris species containing relatively more solvent. Phases for Clostridium MP flavodoxin were calculated using SrnIII (76) and Au1 (66,66)derivatives and incorporating anomalous scattering from each, but Watenpaugh et al. (29,64) have demonstrated that the Sm"' derivative alone, thanks to the large anomalous scattering TABLE I CRYSTALLOQRAPHIC DATAFOR FLAVODOXINS

Space group

Flavodoxin

D. vulgaris oxidized

Clostridium MP oxidizedd

Clostridium MP,

Cell dimensions

P43212 a = b = 51.6 c = 139.6 P3121 a = b = 61.56 c = 70.36

P3121

a

P3121

a

semiquinone

c

Clostridium MP, reduced!

c ~~

Resolution

(b)

(m).

Heavy atoms

2.0

0.74-0.68* Sm"1 at Glu-25, ASP-63" 1.9' 0.81 Sm"1 at Glu-123, ASP-124 AuI a t Cys-53, CYS-128 = b = 61.63 2.08 0.72 = 70.98 = b = 61.68 2.5 = 71.05

~~~~

Mean figure of merit. The variation from the innermost to outermost resolution ranges. From Jensen and Watenpaugh (78). Partial refinement of this structure has reduced the crystallographic R factor to 0.27. The smallest intensities, representing about 15% of the scattering, have not been included in the maps. Difference map, with coefficients m(lFlr.d - IFI.,) exp ia., (79).

76. M. L. Ludwig, R. D. Andersen, P. A. Apgar, R. M. Burnett, M. E. LeQuesne, and S. G. Mayhew, Cold Spring Harbor Symp. Quant. Biol. 36, 369 (1972).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

71

by this element, can suffice for phase determination. Sm"' has provided a suitable heavy atom derivative for still another highly acidic protein, bacterial ferredoxin (7'7), and is observed in all three structures to bind at Asx and Glx residues. However, the Sm positions are not identical in the two flavodoxin structures (Table I). One-electron reduction of both crystalline flavodoxins is accompanied by rather large changes in the diffracted intensities (28,'78). R I = 2110x- I s q l / 2 ( I o x I s , ) is 0.33 for the Clostridium M P data to 2 A resolution. Because of the magnitude of the intensity differences, the phases for the semiquinone form of Clostridium M P flavodoxin have been determined independently, utilizing the same heavy atom substituents (65,699).The average difference in phase angle, determined by isomorphous replacement, is 5 9 O . (Uncorrelated phases would give a value of goo.) Examination of the structures shows that for Clostridium M P flavodoxin the intensity differences arise from a combination of conformational changes with an overall (rigid-body) motion of the molecules in the unit cell. The X-ray and chemical sequence studies of Clostridium M P flavodoxin were complementary. In the map of the oxidized form obtained by isomorphous replacement phasing of 1.9 A data, more than 20 residues were insufficiently defined to permit their identification. Six of these residues occur in the region 36-48, adjacent to a large channel of solvent, and five others at 109-114 and 122-124, also at the surface of the molecule. Identification of residues 88-96 in the map clarified the sequence of this segment and observation of the branch points on the side chains discriminated between Leu and Ile a t residues 49 and 50. At the present time there are no known discrepancies between the chemical sequence and the electron density map. b. Molecular Conformation. The drawings of Fig. 2 depict the folding of the polypeptide chain and the relative orientation of the prosthetic group in oxidized Clostridium MP and D. vulgaris flavodoxins. The close similarity of the two structures is evident. Both flavodoxins are characterized by a high proportion of secondary structure; in each molecule a central parallel p sheet is flanked on either side by pairs of helices. No antiparallel sheet is found, but the chain changes direction a t a num-

+

77. E. T. Adman, L. C. Sieker, and L. H. Jensen, JBC 248, 3987 (1973). 78. L. H. Jeneen and K. D. Watenpaugh, private communication. 79. IFI.., IFlaq, I F I I C d , structure factor amplitudes for oxidized, mmiquinone, and reduced crystals, respectively, referred to as IFIObs when the oxidation state is phases for the oxidized or semiquinone structures, deterobvious; a,, or as,,, mined by isomorphous replacement; IFIc.lc and a,.le, structure factor amplitude and phase, respectively, computed from the atomic positions; m, figure of merit.

72

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

FIQ.2. Drawings of the C, and FMN atoms of (a) oxidized Clostridium MP flavodoxin and (b) oxidized D . vuZguriS flavodoxin. In (a), C. of residue 115 eclipses C, 114. Residue 1 of the D. vulgaris chain is omitted since its image does not appear in the electron density (78). The models are shown in approximately the same orientation with respect to the protein atoms, and hence the different arrangement of the isoalloxaaine ring is emphasized. (a) is from Burnett et ul. (66).

ber of hydrogen-bonded 3,0 bends (80). The hydrogen bonding schemes for the p sheet, shown in Fig. 3, are based on coordinates obtained from the isomophous replacement maps. I n Clostridium MP flavodoxin, the following residues appear to contribute a t least one hydrogen bond for helix formation: 10-27, 66-74, 93-107, and 124-138. Deviations from the helix have been noted for certain of these residues (66). According to the skeletal model constructed from the 1.9 A map, the number of residues assigned to helices, 310 bends, or p structure is 115 of the total 138 in Clostridium MP flavodoxin (66). Examination of the p sheets and flavin binding regions of the two molecules reveals the importance of water in maintenance of the structures. In two locations the regular sheet hydrogen bonding is interrupted by bonding to solvent or side chains (Fig. 3). The D.vulgaris molecule sub80. C . M. Venkatachalam, Biopolymers 6, 1425 (1968).

2. FLAVODOXINS

AND ELECTRON-TRANSFERRING FLAVOPROTEINS

73

FIG.3. Hydrogen bonding schemes proposed for the parallel /3 sheet found n Clostridium MP and D. vulgaris flavodoxins (66,SS). The residues of Clostridium M P flavodoxin are shown above, in larger type; the equivalent D. vulgaris residues are beneath in smaller lettering. Water molecules and hydrogen bonds found only in D. vulgaris flavodoxin are represent.ed by smaller letters and dotted bonds, respectively. Dashed hydrogen bonds are common to both structures with the exception of the bond from Asx-122 to water, which does not occur in D . vulgaris flavodoxin.

stitutes a solvent interaction for the hydrogen bond to Trp-95 which occurs in Clostridium M P flavodoxin. Similar exchanges of solvent for side chain interactions can be observed in Fig. 4. A water near 0-1 in Clostridium M P flavodoxin is "replaced" by Trp-60 in the D. vulgaris molecule; similarly, a solvent molecule bridging the ribityl 0-4' and the carbonyl o f residue 128 in D. vulgaris flavodoxin is the counterpart of the side chain of Ser-87 in the Clostridium M P structure. Comparisons of the folding of D. vulgaris and Clostridium M P flavodoxins with several pyridine nucleotide dehydrogenases have suggested that the flavodoxins may be members of a larger family of nucleotidebinding proteins (81,82). Appropriate superposition of the parallel sheets of flavodoxin and lactate dehydrogenase brings the F M N phosphate into approximate correspondence with the adenine phosphate of NAD' and aligns the first, second, and third helices of flavodoxin with helices a B , CrE, and cwF of LDH, respectively. However, the positions of the flavin and nicotinamide rings do not quite coincide in this superposition. The 81. M. G. Rossmann and A. Liljas, J M B 85,177 (1974). 82. M. G. Rossmann, D. Moras, and K. W. Olsen, Nature (London) 250, 194 (1974).

74

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

A

3

(b) FIG.4. Stereo view of the FMN-binding sites of (a) oxidized Clostridium M P and (b) oxidized D. vulgaris flavodoxins. The two drawings have been oriented to provide approximately the same view of equivalent protein atoms. Some bound solvent atoms appear in both drawings. The hydrogen bonding scheme for D. vulgaris flavodoxin, included in (b) can be compared with that for Clostridium M P flavodoxin, shown in Fig. 5.

homologies between the flavodoxins and dehydrogenases are presented and discussed in more detail in Chapter 2, Volume XI. c. T h e Flavin Mononucleotide Binding Site. Stereo views of the two FM N binding sites are presented in Fig. 4, and the probable hydrogen bonding interactions between F M N and protein in Figs. 4 and 5 and Table I1 (69, 83-86). In both structures the isoalloxazine ring is found 83. K. D. Watenpaugh, L. C. Sieker, J. R. Herriott, and L. H. Jensen, Acta Crystallogr., Sect. B 29, 943 (1973). 84. R. Diamond, Acta Crystallogr. 21, 253 (1966). 85. R. Diamond, Acta Crystallogr., Sect. A 27, 436 (1971).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

75

FIG.5 . Proposed hydrogen bonding contiibutions to the FMN-protein interactions in Clostridium MP flavodoxin. The orientation is shifted slightly from Fig. 4a. For the flavin ring and the ribityl side chain, the bonds indicated by (-.-) are those which seemed most likely according to the initial model, before refinement. The bonds to phosphate oxygens were selected to illustrate the similarity t o the interactions in D. vulgaris flavodoxin. As can be seen from Table 11, alternative or additional hydrogen bonds are possible.

at the periphery of the molecule with the dimethylbenzene end accessible to solvent and the pyrimidine portion “buried” in the protein. Two segments of the polypeptide chain, residues 56-59 and 89-91 in Clostridiurn M P or 60-62 and 95-102 in D. vulgaris flavodoxin provide interactions with the isoalloxazine ring. The ribityl side chain extends toward the interior of the protein (P:N-10 = 8.5 A in Clostridium M P flavodoxin) , permitting OH-2’ and OH-4’ to form hydrogen bonds with protein atoms. The predominant contribution to the binding of the phosphate moiety is made by residues 7-12 (1C15 in D. vulgaris flavodoxin), which include the initial turn of an a-helix. In both structures the resolution is sufficient to determine the P-0 directions. The conformation of the protein-bound FMN, in terms of the torsion angles along the ribityl-phosphate side chain, is similar to that observed in model structures (69,86). A difference in the torsion angles at N-10 to C-1’ correlates with the dissimilar orientations of the isoalloxazine rings in D. vulgaris and Clostridiurn M P flavodoxins (see below) ; otherwise, the FMN conformations in the two structures are very similar. From C-1’ to C-4’ the chain is in a trans, extended conformation, but close contact of 0-2’ and 0-4’ is avoided by a rotation of the (2-3’ to C-4’ bond (x = 60° rather than B O O ) . The dihedral angles for C-4’ to 86. K. D. Watenpaugh and L. H. Jensen, in “Structure and Conformation of Nucleic Acids and Protein-Nucleic Acid Interactions” (M. Sundralingam and S. T. Rao, eds.) (in press).

76

STEPHEN G . MAYHEIW AND MARTHA L. LUDWIG

DISTANCES BETWEEN FMN

AND

TABLE I1 PROTEIN ATOMS”IN Clostridium M P FLAVODOXIN ~

FMN atom

0-1

0-11

0-111

0-4’

~~

~~

~

~~~

Distance

FMN atom

Protein or solvent atom

0-3’

W-2 w-3

2.7 3.0

w-lb

2.5 2.8 2.7

0-2’

Ala-55 CO

2.9

Ser-7 OH Thr-12 NH Asn-11 NH Thr-12 OH Gly-8 NH

2.7 2.8 3.6 3.3 3.7

N-1

Gly-89 NH

2.9

0-2

Thr-9 OH Thr-9 NH Gly-10 NH Asn-11 NH Ser-7 OH Asn-11 (NH1,O) Gly-8 NH

2.9 3.0 3.4 3.2 3.4 3.7 3.7

Gly-91 NH Gly-89 NH Trp-90 NH w-4

2.8 2.9 3.4 3.3

N-3

Glu-59 COO-

2.8

0-4

Glu-59 NH Asp-58 NH

2.9 3.4

N-5

Asp-58 NH

4.2

Ser-87 OH Asn-11 (NH2,O) Asn-119 (NH,,O)

2.6 3.0 3.6

Protein or solvent atom Gly-8 NH Ser-54 OH

(A)

Distance

(A)

Selected heteroatoms in the vicinity of flavin atoms. Most of the listed atoms are displayed in Fig. 5, but every pair does not necessarily form a hydrogen bond. Distances were calculated from protein coordinates obtained after four cycles of difference FMN coordinates Fourier refinement (83),followed by real space refinement (69,86). were determined by real space refinement (89,86). Although designated &B water, this peak could represent an NHI+ ion.

C-5’ and C-5’ to 0-5’ produce approximately trans conformations about these bonds, and the phosphate oxygens are partly staggered with respect to the C-5’ to 0-5’ bond. As in certain riboflavin structures (87,88), the ribityl 0-2’ is approximately cis to the flavin N-1. The isoalloxazine ring appears to be planar, as expected for the oxidized state (89). When the 87. T. D. Wade and C. J. Fritchie, Jr., JBC 248, 2337 (1973); W. T. Garland, Jr., and C. J. Fritchie, Jr., JBC 249, 2228 (1974). 88. D. Voet and A. Rich, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 23. Univ. Park Press, Baltimore, Maryland, 1971. 89. P. Kierkegaard, R. Norrestam, P.-E. Werner, I. Csoregh, M. von Glehn, R. Karlsson, M. Leijonmarck, 0. Ronnquist, B. Stensland, 0. Tillberg, and L. Torbjornsaon, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 1. Univ. Park Press, Baltimore, Maryland, 1971.

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extent of folding about the N-5:N-lO direction was estimated from the electron density of oxidized Clostridium M P flavodoxin, using real space refinement ( 8 5 ) ,the angle between the dimethylbenzene and pyrimidine planes was found to be less than 2 O (69). The orientation of the protein about the phosphate group appears identical in the two flavodoxins. Four homologous hydroxyamino acids and five backbone NH groups are near the phosphate oxygens. The environment of the phosphate is remarkable in several respects. First, the phosphate is partially buried in a region devoid of countercharges. I n neither structure are there any neighboring basic residues to compensate for the charge on the phosphate, presumably bound as either the mono- or dianion (90). The environment is quite unlike that observed for NAD’ in lactate dehydrogenase (91), or for nucleotides bound to ribonuclease A (9.2)or to staphylococcal nuclease (9S),where arginine or lysine residues are adjacent to the phosphate. The solvent or counterion access to phosphate oxygens has not been systematically computed (94), but appears to be limited. I n Clostridium MP flavodoxin, only one of the three oxygens seems able to form hydrogen bonds to solvent without displacing protein atoms. A solvent molecule is observed about 2.7 A from this oxygen (Table 11).In the D.vulgaris structure this position is occupied by atoms of Trp-60. Second, the phosphate binding site is conserved, despite the differences in the isoalloxazine interactions in the two structures. Finally, the phosphate group appears to be essential for association of flavins with Clostridium M P and P. elsdenii flavodoxins, whereas D.vulgaris flavodoxin readily binds riboflavin (Section II,C13). The ribityl side chain interactions are similar though not identical in the two structures. A backbone carbonyl to OH-2’ hydrogen bond occurs in each molecule, as does an interaction between 0-3’ and solvent and between Asn-11 (or Asn-14) and 0-4’. The other 0-4’ interactions differ slightly (Figs. 4 and 5). Contrasts in the isoalloxazine-protein interactions are striking. I n Fig. 4 the drawings are oriented to optimize matching of the protein atoms; when the protein “overlaps” are maximized, the flavin rings are found to be inclined to one another at an angle of - 2 4 O (96). Both flavin rings 90. M. L. MacKnight, J. M. Gillard, and G . Tollin, Biochemistry 12, 4200 (1973). 91. M. J. Adams, M. Buehner, K. Chandrasekhar, G. L. Ford, M. L. Hackert, A. Liljas, M. G. Rossmann, I. E. Smiley, W. S. Allison, J. Everse, N. 0. Kaplan, and S.G. Taylor, Proc. Nat. Acad. Sci. U . S. 70, 1968 (1973). 92. F. M. Richards and H. W. Wyckoff, “The Enzymes,” 3rd ed., Vol. 4, p. 647, 1971. 93. F. A. Cotton and E. Hazen, Jr., “The Enzymes,” 3rd ed., Vol. 4, p. 153, 1971. 94. B. Lee and F. M. Richards, J M B 55, 379 (1971). 95. The estimated error in positioning each ring is 2 3 ” .

78

STEPHEN G . MAYHEW AND MARTHA L. LUDWIG

are sandwiched between hydrophobic residues, but these residues differ in the two flavodoxins (Figs. 4 and 5). In Clostridium M P flavodoxin, they are Met-56, toward the interior of the molecule, and Trp-90, which partially shields the flavin ring from solvent. I n D. vulgaris flavodoxin Trp-60 is inside and Tyr-98 occupies the outside of the flavin ring. The indole and isoalloxazine planes are not parallel in either structure, but Tyr-98 is stacked with the flavin ring in D. vulgaris flavodoxin. All the hydrogen bonds between the flavin ring and the protein appear to be different in the two structures. The only possible similarity would be the interaction of the flavin 0-2 with a backbone N H (89 in Clostridium MP, 95 in D. vulgaris). This bond is not drawn in Fig. 5, because the angular orientation is poor. The N-3 and 0 - 4 interactions utilize dissimilar segments of the polypeptide chain in the two molecules. A backbone NH is the nearest neighbor of the flavin N-5 in both cases, being about 4 A from that flavin atom. While this distance appears too great for formation of a hydrogen bond, the accuracy of the initial models is insufficient to preclude its formation, and Watenpaugh et al. (64) have included this interaction in their hydrogen bonding scheme (Fig. 4b). Burnett et al. (66) concluded that this hydrogen bond is unlikely, both because N-5 in oxidized flavins is not a very good acceptor (96,973 and because the distance between N-5 and the Asp-58 amide is found to be 4.2 A after difference Fourier (83)and real space refinement (85) of oxidized Clostridium M P flavodoxin. In both structures two acidic groups are in the neighborhood of the flavin: residues 58 and 59 in Clostridium M P flavodoxin and 62 and 99 in D. vulgaris flavodoxin. They are oriented somewhat differently in the two molecules. Assuming that electron transfer occurs within a molecular complex between flavodoxin and a “reductase,” one might have expected the three-dimensional arrangement in the neighborhood of the flavin ring to be more highly conserved. The differences ought to be reflected in the relative efficiencies of transfer from the two flavodoxins to a given acceptor. d. Distv-ibution of Residues. The results of chemical modification of cysteine, tyrosine, and tryptophan have suggested that integrity of these side chains is essential for maintenance of the F M N binding site (Section II,C,4). Hence, the position of these residues in each structure is of special interest, In Ctostridium MP flavodoxin, none of the cysteine residues is in direct contact with FMN. The nearest Cys, a t position 53 in the parallel sheet, is adjacent to Ser-54, which forms a hydrogen bond to the flavin phosphate, but the side chain of Cys-53 necessarily protrudes 96. M. Sun and P.4.Song, Biochemistry 12,4663 (1973). 97. F. Miiller, P. Hemmerioh, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Press, Baltimore, Maryland, 1971.

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from the opposite side of the sheet, away from the FMN. The remaining cysteine sulfurs a t residues 128 and 108 are at distances of 12 and 22 A, respectively, from the F M N phosphorus. The relative positions of the tyrosine and tryptophan residues of Clostridium M P flavodoxin can be seen in Fig. 9. All the tyrosines are separated from the F M N by intervening atoms. Tryptophan-90, on the other hand, is an immediate neighbor of the isoalloxazine ring and would be expected to influence the properties of the protein-bound flavin. The five phenylalanine rings are all inside the molecule, with 66, 69, and 99 forming a central hydrophobic cluster. Desulfovibrio vulgaris flavodoxin contains four cysteines ; as in the Clostridium M P molecule, none of these is involved in a disulfide linkage. Cysteine-57 is in the same position as Cys-53 of the Clostridium M P protein. The backbone amide N of Cys-102 forms a hydrogen bond to the isoalloxazine 0-2, and Cys-93, in the neighborhood of the phosphate, corresponds approximately to Ser-87 of Clostridium M P flavodoxin. However, neither of these -SH groups interacts with atoms of FMN. Cysteine90 of D . vulgaris flavodoxin is more distant from the FMN binding site. Both Tyr-98 and Trp-60 are clearly involved in flavin binding in the D. vulgaris protein (64,78). According to the sequence analysis, there are nine aspartate and 19 glutamate residues in Clostridium M P flavodoxin, but only two arginines and 10 lysines. Four of the acidic groups are clearly paired with basic residues, and many of the remaining carboxylates are within 5-7 A of the charges on arginines or lysines. However, two clusters of uncompensated negative charge occur on the surface of the molecule. One acidic region, residues 62-67, is near the flavin; and the other, residues 120-125, includes the ligands which bind samarium. The arrangement of charged areas may be functionally important since the binding of flavodoxin to ferredoxin-NADP+-reductase is very dependent on ionic strength (42-45). e. Structure as a Function of Oxidation State. Comparisons of independently phased oxidized and semiquinone structures (Table I) have been made for crystalline Clostridium M P flavodoxin (69). The molecular displacement accompanying reduction complicates direct comparison of the two electron density maps. To detect possible conformational changes, small rotations and translations had to be applied to the map of the semiquinone form to maximize its correspondence with the oxidized density (98). The resulting density was then optically superposed on the skeletal model of oxidized flavodoxin. In nearly all regions the agreement between the semiquinone map and the oxidized structure was close. I n particular, the conformation of the ribityl side chain appeared to be unchanged, and the isoalloxazine ring was essentially planar. Real space refinement of 98. J. M.Cox, JMB 28, 151 (1967).

80

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

111

Ibl

FIO.6. The proposed conformations of residues 58-59 in (a) the oxidized form and (b) the semiquinone form of Clostridium MP flavodoxin. The drawings are based on coordinates obtained by real space refinement (86). The view is not quite perpendicular to the flavin ring. Except for the N-5 proton in the semiquinone, hydrogens have been omitted, nor are the full electronic structures shown Ccf. formula (I), Section II,D,51.

the F M N (69,86)yielded a bending angle similar to that found for oxidized flavodoxin (98~). Several discrepancies between the oxidized model and the semiquinone density have been observed in the FMN binding site and attributed to conformational differences (69).Some of the changes proposed to accompany electron transfer are illustrated in Fig. 6. The density suggests that a movement of the indole ring of Trp-90 and a rearrangement of the bend involving residues 56 through 59 result from one-electron reduction of the FMN. The tryptophan ring motion is complex, involving rotations about both the C,Cp and C,g-C, bonds, but the angle between the indole ring and the mean flavin plane changes only slightly as a result. In the map of oxidized Cbstridium M P flavodoxin, the density suggests that the carbonyl oxygen of Gly-57 points down, away from the FMN, and in nearly the same direction as the p-carbon of Asp-58. The observed conformation resembles the energetically unfavorable Type I1 3,, bend (80). However, in the semiquinone structure the bend appears to revert to the more stable Type I arrangement by a reorientation of the peptide connecting residues 57 and 58, allowing the formation of a hydrogen bond between N-5 and the backbone oxygen 57, which is now about 3.0A from the flavin nitrogen. Hydrogen bonding in the bend may also be perturbed in the semiquinone structure since the peptide planes shift slightly (Fig. 6). Conformational changes as subtle as those suggested by the electron density maps are di5cult to prove by crystallographic methods especially 98a. The bending angles for the flavin ring in reduced and radical forms of Clostridium MP flavodoxin have been found to be about 9" and 6', respectively.

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when accompanied, as in this case, by the small movement of many other atoms. A difference synthesis using terms m(lFI,, - IF,,) exp iaox ( 7 9 ) , with phases determined by isomorphous replacement, contained features consistent with the above interpretation but was not definitive. Therefore, refinement and extension of the data have been undertaken. If the data and phases can be improved sufficiently, then difference Fourier maps with coefficients ( lFlobs- (Flcale) exp iaeslc(79) should reveal the conformation of residues 56-59 when these atoms are omitted from the structure factor calculation. For oxidized flavodoxin, the use of calculated structure factors with R = 0.27 and data to 1.9 A resolution results in densities corresponding to the 0-57 arrangement deduced from the isomorphous replacement maps ( 6 9 ) . Final verification of the proposed changes will require further refinement of the models for both oxidation states. The structural differences indicated by the two isomorphous replacement maps, if proved valid, have several chemical implications. The flavin semiquinone is bound more firmly by the protein than is oxidized F M N ; the change in I(, reflects the perturbation of the redox potentials of F M N by the protein (Sections II,C,2 and E) . Additional FMN-protein interactions and formation of the more stable Type I bend in the semiquinone state would both tend to account for the larger K,, for association of the FMN semiquinone. Undoubtedly other phenomena, such as the altered charge distribution in the flavin ring, also contribute to the net change in free energy of association. Further, the formation of a hydrogen bond between the protonated N-5 and a peptide oxygen provides a means of stabilizing the neutral form of the flavin radical. The pk’ of the N-5 proton is displaced upward by more than two units in the presence of the protein (Section 11,DJ). The ionization of this proton may also be affected by the negative charges on Glu-59 and Asp-58. Finally, if a conformational change must occur during the oxidation-reduction reaction, then the potential energy barrier for rearrangement might limit the rate of electron transfer. It should be emphasized that the proposed structural changes, associated with formation of the semiquinone of Clostridium M P flavodoxin, are unlikely to be duplicated in all flavodoxins. The Clostridium M P and D. vulgaris structures are folded differently in the loops adjacent to the isoalloxazine ring. I n particular, the oxidized D. vulgaris chain does not form a 310bend in the region corresponding approximately to residues 56-59 of Clostridium M P flavodoxin. Thus, although the D. vulgaris protein stabilizes the neutral F M N semiquinone and has an E2 redox p6tential somewhat different from that of free FMN, the structural “explanations” for these phenomena cannot be precisely the same as those offered for Clostridium M P flavodoxin.

82

STEPHEN G. MAYHEW AND MARTHA L. LmwIa

X-Ray intensities for fully reduced crystals of Clostridium M P flavodoxin have been measured to 2.5 A resolution. The intensities are almost identical with those for semiquinone crystals ( R I = 0.06) ; difference Fourier maps comparing the semiquinone and reduced states do not suggest any large rearrangements involving the F M N or its surroundings (69). Other techniques affirm that the conformations of the semiquinone and reduced states of flavodoxins are very similar. Neither the NMR spectra of Clostridium M P flavodoxin (99) (Section II,D,5) nor temperature-jump studies of the reduction of A . vinelandii flavodoxin (100) provides evidence for conformational changes accompanying formation of the fully reduced states. The crystallographic results suggest that the flavin ring in reduced Clostridium M P flavodoxin is nearly planar (98a). This is presumably not its most stable conformation ; structural analyses of fully reduced flavins (89) indicate that the dihydroisoalloxazine ring prefers a conformation which is folded along the N-5:N-lO line.

C. FLAVIN-PROTEIN INTERACTIONS: CHEMICAL AND PHYSICAL STUDIES IN

SOLUTION

1. Preparation and Properties of the Apoprobein FMN is released from flavodoxins by treatment with TCA (15,46,61) or other acids (5,11,59,60), 2 M KBr at pH 3.9 (59), guanidine hydrochloride a t pH 7 (8,11), and, in certain cases, by reaction with mercurials (8,13,40). Solutions of apoflavodoxins are colorless, with a single absorption maximum in the near UV (A, 280 nm, c = 25,000-26,000 M-l cm-l) and fluoresce upon excitation of tyrosyl and tryptophanyl residues (59,62,101). Circular dichroism spectra in the far UV show that the secondary structure of apoflavodoxins is different from that of native flavodoxins (62). In A . vinelandii flavodoxins, this change in conformation on removal of the flavin does not affect the overall exposure of tryptophan residues, but, as judged by the effects of ethylene glycol on the UV absorption spectrum, it may decrease the exposure of tyrosine residues to solvent (101). Nevertheless, the tyrosines titrate normally in the apoprotein whereas in the holoprotein their pK values are displaced upward and the titration becomes partly irreversible (106,103). CI

99. T. L. James, M. L. Ludwig, and M. Cohn, Proc. Nat. Acad. Sci. U. S. 70, 3292 (1973). 100. B. G.Barman and G. Tollin, Biochemistry 11,4755 (1972). 101. J. A. D’Anna, Jr., and G. Tollin, Biochemistry 10,57 (1971). 102. D.E.Edmondson and G. Tollin, BiochemCtry 10, 133 (1971). 103. G. Tollin and D. E. Edmondson, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 153. Univ. Park Press, Baltimore, Maryland, 1971.

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FLAVODOXINS AND ELECTRON-TRANSFERRING

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Reconstitution of the holoproteins can be achieved with good yields provided precautions are taken to avoid oxidation of sulfhydryl groups, which appear to be more accessible in the apo- than in the holoproteins (59,104).The CD spectra of native and reconstituted proteins are very similar ( 6 2 ) . Complete regeneration of the structure has been demonstrated for Clostridium M P flavodoxin ; after reconstitution this protein yields crystals whose diffraction patterns are identical with crystals of untreated protein (105). 2. Themnodynamics and Kinetics of Flavin Binding

Association constants for FMN and apoflavodoxins from A . vinelandii (60,61), P. elsdenii (59,104), and other flavodoxins (59,100)have been

determined by equilibrium titrations monitored by fluorescence or absorbance measurements (see Table 111). In the case of the A . vinelandii protein, the constant is almost independent of p H from p H 4.5 to 8.0 (90).Since the second ionization of the F M N phosphate occurs near p H 6 (106) these results imply that both the moao- and dianion forms of the phosphate are bound with approximately equal affinity. Below p H 4.5 the association constant decreases abruptly, suggesting that protonation of two groups affects the FMN-protein interactions (90). The equilibrium constants for association of F M N semiquinone and hydroquinone with apoflavodoxins can be calculated from the association constants of the oxidized protein and the measured shifts of the redox potentials of F M N (59,100). Such calculations show that F MN semiquinone is usually bound very much more tightly than either the oxidized or fully reduced flavin. The calculated K , differences are especially dramatic for flavodoxin from A . vinelandii (Table 111). During F M N binding the protein and flavin fluorescences are quenched in parallel and the quenching reaction appears to be second order (59,61,101). For P. elsdenii flavodoxin a maximum rate of association occurs near pH 4.5, but for A . vinelandii flavodoxin the reaction rate continues to increase as the pH is lowered to about 4 (90).Barman and Tollin ( l o r ) ,employing temperature-jump techniques, have shown that association of F MN with apoflavodoxin from A . vinelandii actually proceeds in two steps. After applying the temperature perturbation, they observed a decrease in flavin fluorescence during about 5 sec followed 104. S. G. Mayhew, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 185. Univ. Park Press, Baltimore, Maryland, 1971. 105. M. L. Ludwig, R. Andersen, P. A. Apgar, and M. LeQuesne, in ‘(Flavins and Flavoproteins” (H. Kamin, ed.), p. 171. Univ. Park Press, Baltimore, Maryland, 1971. 106. H. Theorell and A. P. Nygaard, Acta Chem. Scand. 5, 1649 (1954).

TABLE I11 BINDINGOF FLAVINS BY FLAVODOXINS' Apoflavodoxin

A. vinelandii Flavin FMN derivatives FMN, oxidized semiquinone reduced ZPropyl%Methyl3-CHzCOO5DeaeaIsoDWXYRiboflavin derivatives Riboflavin DeoxyISO-

K., M-1

(24", pH 7) 2.0 x l o a d 5.8 X lolac 1.4 x 1091

3.7

x x x

ki," M-I sec-' 2

x

106

107

4 107 107 4.8 1 . 3 X 108

4

x

104

1.8 X 1 0 6 2.4 X i o a 1.7 X 106 6.3 x 107

8.9 X

lo6

K . (20", PH 7) 2.3 x 2.9 X 1.1 x 1.3 x 3.3 x 1 x

109 10'' 108 104 108 107

++ 2.3 X 10' -(
klJc(0.5" pH 7) 1.4 X

lo6

6.8 X lo4 3.2 X 108 6 X 10' 4.9

x

Radical at half reduction (%) 95

D. vulgaris K., pH 8.2 8.2 x 107 5 X looe 5 x 10k

Ref. 13,69, 61,100 65 6a,61

66 Small

63 107,108

79

63,61

104 1.3 X 10''

63,61

?

61,69,107 61 63.61 61.

Eic

Riboflavin SO4 Other flavins N-10 w-Carboxvbu t,ylisoalloxkzine 3.1 X 106 63,61 FAD 1 3 x / isA1 ,. in6 ---,-Likiflavin 2.2 x 105 4 x 107 63,61,107 a No entry indicates not measured; indicates compound binds, K . not measured; and - indicates binding not detectable under experimental conditions. !This protein also binds 6- and &OH-FMN, and 7,&dichloro-FMN. For the related Clostridium MP flavodoxin, using K. for oxidwed P. elsdenii flavodoxin and redox potentials at pH 7 (Table IV), K . for the semiquinone = 7.1 X 10" and K,, for the hydroquinone = 1.0 x 108. Overall bimolecular rate constant for formation of the flavin-protein complex. Reported values range from 1.7 to 2.2 X 108. Redox potentials at pH 8.2 were used to calculate these values. f pH 7.

+

m

u

E

e

P

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING

FLAVOPROTEINS

85

by a slower increase during a further 25 sec. The faster relaxation was independent of concentration. They concluded that in F M N binding, the initial second-order reaction of flavin and protein precedes tt faster rearrangement in which fluorescence is further quenched. For A . vinelandii flavodoxin at loo and pH7: Apoprotetn

+ FMN

kl =

5.3

X

k-,= 8.1

X

lo4 M-' sec-' (Apoprotein - FMN)

sec-l

(2)

k, = 0.16 sec-'

a_,= 4.2

X

sec-I

Holoprotein

Because T~ GS r2 a t the concentrations used, the two relaxations had to be resolved by analog fitting to the experimental traces. The forward bimolecular rate constant, k,, is in reasonable agreement with values obtained by stopped-flow measurements (61).Rate constants were not evaluated for P . elsdenii flavodoxin, which displays similar behavior, but the first relaxation appears to be more rapid for this protein. The occurrence of a first-order process is consistent with the shifts in CD spectra resulting from addition of FMN (62),but the observed relaxation should not be ascribed t o changes in the secondary structure of the protein until rates of the CD spectral changes have been determined. Binding of FMN to the apoprotein of A . vinelandii flavodoxin involves a large positive entropy of activation; the activation energy for the reaction is 15.8 kcal/mole (lor),in contrast with the value of 8.3 kcal/mole determined for P . elsdenii flavodoxin (59). 3. Binding of Modified Flavins

The flavin binding site of flavodoxins has been explored by studying the effects of substituting other flavins for FMN. The energies of binding and the properties of the complexes with different flavins vary widely (Table 111).The most extensive studies of the equilibria and kinetics of binding of flavin analogs have been carried out with A . vinelandii flavodoxin by Tollin and co-workers (61,107,108). More limited data are available for the P . elsdenii and D. vulgaris proteins. The free energies and kinetics of binding are affected by substitutions or deletions a t any of a number of positions in the prosthetic group. Deletion of a substituent does not always provide an estimate of its contribution to the overall energy of interaction. Some effects appear explicable only if one assumes 107. B. G. Barman and G. Tollin, Biochemistry 11, 4746 (1972). 108. D. E. Edmondson, B. Barman, and G. Tollin, Biochemistry 11, 1133 (1972).

86

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

that the conformation of the protein and/or the flavin in analog complexes differs from that found in the FMN holoproteins (61). The effects on K, of substitution in the isoalloxazine moiety are generally consistent with the multiple interactions between the flavin ring and the protein observed in the known structures (Table 111).Substituents at N-3 and C-2 uniformly decrease the association constants. Although the dimethylbenzene end of the flavin ring is relatively accessible to solvent, the introduction of a methyl group a t position 6 (iso-FMN) can be shown by model building to produce close contacts with residue 57 in Clostridium M P flavodoxin. The redox properties of FMN derivatives of A . vinelandii flavodoxin modified a t positions 2, 3, and 6 differ from those of native flavodoxin; the potentials and rates of reduction by dithionite are significantly altered (102). Nevertheless, in many derivatives of A . vinelandii or P. elsdenii flavodoxin a semiquinone form is stabilized a t half-reduction (102) and biological electron transfer reactions can proceed, albeit with diminished efficiency, when modified flavins are incorporated into P. elsdenii flavodoxin (63).Deaza-FMN is isosteric with F M N and its different behavior must be attributed to altered electronic properties (96,108). Bound deaza-FMN does not form a semiquinone, but is instead partially converted to the fully reduced form by reaction with dithionite (108). From the effects of this analog on the rates and equilibria of binding, an interaction between the protein and N-5 of the F M N in oxidized A . vinelandii flavodoxin has been inferred (108). Such an interaction is not predicted from the structure of Clostridium M P flavodoxin (Section II,B,3). The results of modification of the ribityl side chain vary with the species of flavodoxin. The hydrogen bonding interactions of OH-2’ and OH-4’, observed in the structure of Clostridium M P flavodoxin, would be expected to favor association. Removal of these groups ought to reduce K,. Assuming isomorphism of the Clostridium M P and P. elsdenii structures (109), comparison of the constants for deoxy-FMN and FMN binding to P . elsdenii flavodoxin suggests that these interactions contribute approximately 4 kcal of binding energy. Similar contributions for the D . vulgaris protein are predicted by its structure. Surprisingly, the constants for F M N and deoxy-FMN are reported to be almost identical for A . vinelandii flavodoxin ; however, there must be some stereochemical restrictions on the conformation of the ribityl side chain, since tetra-0109. Because of their similar properties and the homology of the sequences constituting their active sites, Clostridium MP and P . etsdenii flavodoxins have been considered equivalent when the chemical results are compared with the structural data. The term “clostridial flavodoxin” is used to include both C. pasteurianum and Clostridiurn MP flavodoxins.

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FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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acetyl riboflavin fails to bind to the A . vinelandii apoprotein (61). Derivatives with four or six carbon atoms in the side chain do not bind to P. elsdenii apoflavodoxin ( 11 0 ) . The role of the phosphate group in stabilizing the F M N complexes is most intriguing. The apoflavodoxins from A . vinelandii, R. rubrum, and D. vuEgaris all bind riboflavin. I n contrast, apoflavodoxins from Clostm'diu and P. elsdenii are specific for flavin a t the level of F MN (59). Even though the latter proteins have higher affinities for F M N than do the D. vulgaris or A . vinelandii proteins, they do not bind riboflavin (or lumiflavin or FAD) to a detectable extent ( K , < lo3),These observations do not necessarily imply that ring interactions contribute little to the binding energy. Nervertheless, the apparent contribution of the phosphate-protein interactions is very much greater for the clostridial and P. elsdenii flavodoxins than for A . vinelandii or D. vulgaris flavodoxins. Yet the structures show these interactions to be essentially identical in the Clostridium M P and D. vulgaris proteins. To resolve this dilemma, differences in the apoprotein structures have been invoked. From the biphasic binding kinetics and other observations it has been postulated that the ,iif phosphate of FMN is necessary to trigger a conformational change in the apoprotein (62,107). D'Anna and Tollin ( 6 2 ) have proposed that with the FMN-specific proteins this conformational change is much larger. In support of this suggestion, they showed that when flavin is bound by the FMN-specific apoproteins, there is a relatively large change in the ellipticities of the C D bands in the f a r UV region (WS,62). On the other hand, the kinetics of riboflavin binding to A . vinelandii apoprotein show no evidence of a structural rearrangement (10'7). Conceivably, then, the structures of some of the complexes with dephospho analogs are different from those of the FMN proteins. It may be significant that complexes of deoxyriboflavin and N-10-o-carboxybutyl isoalloxazine with A . vinelandii apoflavodoxin fluoresce and differ in other respects from the native flavodoxin and that the riboflavin complex forms negligible amounts of semiquinone (61). Unfortunately, the far UV circular dichroism spectra of riboflavin and related complexes have not been reported. The crystal structures of suitable analog complexes would also be of great assistance in evaluating some of these data. 4. Protein Modifications That AffectFlavin Binding

Chemical modification of cysteines seemed to implicate these residues in maintenance of the flavin binding site of certain flavodoxins. Knight and Hardy (8) found that F M N dissociated from C. pasteurianum flavodoxin upon reaction with sodium mersalyl and proposed that the single 110. R. Gast and F. Miiller, private communication.

88

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

sulfhydryl group in this protein might be important for flavin binding. Subsequent work with Clostridium M P (59), P. elsdenii (59), and D. vulgaris (40) flavodoxins supported the view that one or more sulfhydryls were involved in flavin binding in these proteins. On the other hand, flavin binding by A. vinelandii flavodoxin is not prevented by reaction of the apoprotein with DTNB (111). The possibility that -SH groups bind directly to flavin atoms in Clostridium M P and D. vulgaris flavodoxins has been ruled out by the two structures (Section II,B,3) and it must be concluded that the observed inhibition of F M N binding by N-ethylmaleimide and mercurials (59,104) arises indirectly from effects on the secondary or tertiary structure of these proteins. Chemical evidence for the proximity of tyrosines to FMN in A. vinelandii and P. elsdenii flavodoxins is based on modification by tetranitromethane. Hinkson (60) observed that nitration of four of the five tryosines in A . vinelandii apoprotein eliminated most of its capacity to bind FMN and that in the native protein two tyrosines are protected against nitration. Flavin binding by P . elsdenii apoflavodoxin is similarly inhibited after reaction with TNM (59) and also by reaction with 5 moles of iodine (118).The two -SH groups were protected during iodination and consequently a modification of tyrosines seemed likely. I n the absence of a structure, conclusions about the interaction of tyrosine and F M N in A. vinelandii flavodoxin cannot be assessed, but assuming the similarity of P. elsdenii and Clostridium M P flavodoxins (109) then tyrosyl modifications must exert an indirect effect on flavin binding in both of these proteins since no tyrosines are in contact with FMN. On the basis of results of experiments with N-bromosuccinimide, McCormick (113) proposed that a tryptophan might be complexed with the flavin of C. pasteurianum flavodoxin, a suggestion consistent with the known structures. Ryan and Tollin (111) conducted similar experiments with A. vinelandii flavodoxin, concluding that reaction of a single tryptophan was responsible for loss of F M N binding capacity.

D. SPECTROSCOPIC PROPERTIES 1. Optical Absorption Spectra of Solutions

In oxidized flavodoxins the bound flavin exhibits two broad absorption bands in the visible (Fig. 7) and a third band in the near UV (A,,, 272-275), overlapping the aromatic absorbances of the protein. The peak 111. J. Ryan and G . Tollin, Biochemistry 12, 4550 (1973). 112. E. A. Gowie and S.G. Mayhew, unpublished. 113. D. B. McCormick, Experientia 15, 243 (1970).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING I

I

I

I

I

89

FLAVOPROTEINS I

I

I

I

Wovelength (nm)

FIO.7. Absorption spectra of flavodoxins in solution. Curves 1 and 3: oxidized and semiquinone forms, respectively, of A . vinelaiidii flavodoxin; curves 2, 4, and 5 : oxidized, semiquinone, and fully reduced forms, respectively, of P. elsdenii flavodoxin.

positions, shapes, and intensities of the visible bands vary considerably with the source of the protein and differ from those of free F M N (Table IV) ; for example, the maximum of F M N a t 445 nm is red-shifted by 5-22 nm in most flavodoxins, slightly blue-shifted to 443 nm in flavodoxin from C . pasteurianum, but unchanged in flavodoxins from P . elsdenii and Clostridium MP. The extinction coefficient a t this maximum is usually about 85% of that of free FMN, and in addition the band has more or less pronounced shoulders on each side of the maximum as a result of vibronic splittings. The position and structure of the 373-nm band of F M N is likewise altered in the proteins. Temperature difference spectra of several flavodoxins reveal the vibrational structure of the two visible bands in more detail (114). Peak shifts and vibrational splittings are commonly observed in other flavoproteins and also occur with model flavins when the polariaability or hydrogen bonding capacity of the solvent is varied or when intermolecular complexes are formed with compounds such as indoles (116,116).The inference that in flavodoxins the 114. F. Miiller, S. G . Mayhew, and V. Massey, Biochemistry 12, 4854 (1973). 115. H. A. Harbury, K. F. LaNoue, P. A. Loach, and R. M. Amick, Proc. Nut. Acad. Sci. U.S . 45, 1708 (1959). 118. G. Weber, BJ 47, 114 (1950).

90

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

flavin lies in a relatively nonpolar environment, perhaps containing aromatic residues, is borne out by the structures of the Clostridium M P and D . vulgaris proteins, and the considerable differences in the visible spectra of these two flavodoxins (Table IV) may be attributed to the more aromatic environment of the flavin chromophore in the D. vulgaris molecule. Upon addition of one reducing equivalent, free isoalloxazines are converted to mixtures of the three oxidation states in which the semiquinone constitutes a minor fraction (117). Because of the shifts in redox potential (Table IV) the distribution a t equilibrium in flavodoxin solutions favors the semiquinone in yields approaching 100% a t neutral pH (118). Flavodoxin radicals always display the blue spectrum assigned to the neutral species with a proton a t N-5 (118-121) [formula (I), Section II,D,51; they are not converted to the red anionic form even at pH values above 10 (12,13,102), although the pR of the free FMN radical is approximately 8.6 (117,121).The absorption spectrum of the semiquinone of P . elsdenii flavodoxin (Fig. 7) has maxima at 350, 377, 505, and 580 nm, and a broad shoulder at 627 nm. Spectra of semiquinones of clostridial flavodoxins are similar (7,lS) ; other flavodoxins absorb less around 500 nm (14,16,23,51); and in A . vinelandii flavodoxin the long wavelength shoulder becomes a peak in the region of 615 nm ( 2 3 ) . Muller et al. (122) have proposed that spectra of the latter type indicate less accessibility of the flavin radical to water. Edmondson and Tollin (102) found that a number of flavodoxin semiquinones show a large decrease of absorption around 285 nm, and an increase around 270 nm, compared with the absorption of the oxidized protein. The decrease a t 285 corresponds to a similar change in the spectrum of free FMN on reduction to the semiquinone (102,123); it is suggested that the increase a t 270 nm is either the result of a red shift of the UV absorption maximum of the semiquinone, which lies below 260 nm in free FMN semiquinone (123), or of a charge transfer interaction between the semiquinone and an aromatic amino acid. 117. A. Ehrenberg, F. Muller, and P. Hemmerich, Eur. J . Biochem. 2, 286 (1967). 118. V. Massey and G. Palmer, Biochemistry 5,3181 (1966). 119. G. Palmer and V. Massey, in “Biological Oxidations” (T. P. Singer, ed.), p. 263. Wiley, New York, 1968. 120. G. Palmer, F. Muller, and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 123. Univ. Park Press, Baltimore, Maryland, 1971. 121. F. Miiller, P. Hemmerich, A. Ehrenberg, G. Palmer, and V. Massey, Eur. J. Biochem. 14, 185 (1970). 122. F. Miiller, M. Briistlein, P. Hemmerich, V. Massey, and W. Walker, Eur. J . Biochem. W, 673 (1972). 123. E. J. Land and A. J. Swallow, Biochemistry 8, 2117 (1969).

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FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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Fully reduced flavodoxins from P. elsdenii and Clostridia are pale yellow with Amax near 365 and 315 nm and a broad shoulder centered around 450 nm (Fig. 7). Resemblances to the low temperature spectrum of the anionic form of reduced tetraacetyl-riboflavin suggest that the reduced protein-bound flavin is the anion, and the presence of the long wavelength absorbance appears consistent with a planar conformation for the reduced isoalloxazine in these proteins (1244).Unfortunately, there is little similarly detailed information about the spectroscopic properties of other flavodoxin hydroquinones. However, some differences from P. elsdenii flavodoxin seem likely; for example, the hydroquinone of A. nidulans flavodoxin appears to have less absorbance a t 450 nm (125) and flavodoxin from Chlorella fusca has a peak a t 395 nm rather than near 370 nm (15). 2. Optical Absorption Spectra of Single Crystals

Single crystal spectroscopy offers a means of comparing dissolved and crystalline states. An isotropic ‘‘solution” spectrum can be reconstructed by appropriate averaging of the spectra obtained when a crystal is oriented so that incident light is polarized either parallel or perpendicular to selected crystal axes. Calculation of the flavin semiquinone spectrum from measurements on crystals of Clostridium M P flavodoxin was of special interest since the reddish color of these crystals, noted on examination in unpolarized light (105), suggested that changes in the flavin spectrum might have occurred as a result of crystallization. Spectra of single crystals of Clostridium MP flavodoxin are reproduced in Fig. 8 along with the calculated isotropic spectra (126). The reconstructed semiquinone spectrum is typical of the blue neutral flavin radical. Hence, the radical does not undergo tautomerization or conversion to the anionic form upon crystallization. These conclusions from optical spectroscopy are supported by measurements of the linewidth (19 G) of the E P R spectrum of a crystalline powder sample of flavodoxin semiquinone (Section

II,D,5). From the optical measurements on crystals of Clostridium RiIP flavodoxin (Fig. 8) and the known orientation of the isoalloxazine ring with respect to the crystal axes, i t has also been possible to assign the transition moment directions of the r-r* transitions responsible for the absorbance of the oxidized flavin a t 376 and 445 nm (13,126). Components 124. S. Ghisla, V. Massey, J.-M. Lhoste, and S. G. Mayhew, Biochemistry 13, 589 (1974). 125. B. Entsch and R. M. Smillie, A B B 151, 378 (1972). 126. W. A. Eaton, J. Hofrichter, M. W. Makinen, R. D. Andersen, and M. L. Ludwig, Biochemistry 14, 2146 (1975).

92

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

350

WAVELENGTH (nml 4w

WAVELENGTH (nml

500

'

L a

* I

Ibl

la)

It1

Fro. 8. Single crystal, solution, and polarization ratio spectra of flavodoxin from Clostridium MP. The crystal spectra were measured with the electric vector of the polarized light parallel and perpendicular to the c axis. The polarization ratio, P , is defined as qc/eIc, and is related to 6,the angle between the transition moment directions and the c axis of the crystal. (a) Oxidized flavodoxin. The isotropic spectrum was calculated assuming equal extinction coefficients of 10.4 mM-1 cm-1 for the peaks of the solution and isotropic crystal spectra near 22,000 cm-I (13). (b) Flavodoxin semiquinone. The calculated isotropic spectrum assumed the extinction coefficient of 4.6 mM-1 om-' a t 17,250 cm-I (13). (c) Possible transition moment directions for the a (445 nm) and @ (376 nm) transitions of the oxidized form of Clostridium MP flavodoxin. The angles are defined relative to the y axis, which lies in the flavin plane and is perpendicular to the line joining N-5 and N-10. Directions determined experimentally are the solid arrows, while the dotted arrows are the directions predicted from theory (167). The values of (I = 15" and @ = -5' are considered to be the correct assignments. From Eaton et al. (166).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

93

of these moments parallel and perpendicular to the c axis are derived from the polarization ratios Ell/4,. Loci of the transition moment directions then describe cones about c, whose intersections with the isoalloxazine plane represent the two permissible directions for each in-plane transition. The polarized emission spectra of model flavins, measured b y Sun et al. (127) suggest that the angle between the two transition directions must be about 20°, and measurements on stretched films (128)indicate that both transitions are nearly parallel to the long axis of the isoalloxazine system. This information further restricts the assignment of the moments. The directions shown in Fig. 8c are in good agreement with theoretical calculations for model oxidized isoalloxazines (127). The low energy transitions for the semiquinone, centered near 580 and 500 nm, have directions very similar to those for the 445- and 376-nm transitions of the oxidized form. As a result it has been postulated that the *-orbital charge density is very similar in the oxidized and radical states of the protein-bound flavin (126). Electron transfer involving these oxidation states would therefore require no gross rearrangement of the electronic structure of the flavin. 3. Fluorescence

The greenish yellow fluorescence of unbound oxidized FMN is quenched in the flavodoxins. The residual fluorescence observed in solutions of several flavodoxins is less than 1% of the fluorescence of F M N (69) and is probably the result of traces of flavin dissociated from the holoprotein; a M solution of a flavodoxin with an association constant of lo@ M-l, as is typical of these proteins, would contain about lo-' M free FMN. For all practical purposes these flavoproteins are nonfluorescent. In D. vulgaris and Clostridium MP flavodoxins, the quenching may be attributed to the presence of neighboring aromatic residues. Though the precise mechanism is unclear, quenching of FMN fluorescence by complex formation with aromatic compounds has been demonstrated in various model systems (129). In contrast to a widely held view, ffavin hydroquinones are intrinsically fluorescent ( l Z 4 ) . In some reduced flavoproteins, this emission is quenched, but in flavodoxin from P. elsdenii a weak fluorescence is observed with a peak a t 530 nm (19.4). The fluorescence of the aromatic residues of apoflavodoxins from C h tridia, P. elsdenii, and D. vulgaris is about 99% quenched upon addition of FMN (69,62). I n flavodoxins from A . vinelandii and R. m b r u m , only 127. M. Sun, T.A. Moore, and P.-S. Song,JACS94, 1730 (1972). 128. J.-M. Lhoste, Proc. Eur. Biophys. Congr., I s t , 4, 221 (1971). 129. R.E.MacKenzie, W. Fory, and D. B. McCormick, Biochemhstry 8,1839 (1969).

94

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

FIG.9. A stereo view showing the relationship of the tyrosine, tryptophan, and FMN rings in Clostridium M P flavodoxin. From Burnett et al. (66).

partial quenching of the aromatic fluorescence occurs ; the holoproteins are, respectively, 13 and 60% as fluorescent as the apoproteins (62,101). D'Anna and Tollin (101) have suggested that the quenching of the protein fluorescence may result from two effects: a direct molecular overlap of the side chains of some aromatic amino acids and the flavin, and Forster energy transfer between the different aromatic systems in the molecule. The first of these suggestions is confirmed in the X-ray structures of two flavodoxins. Resonance transfer from tyrosine to tryptophan and from tryptophan or tyrosine to FMN (130) may explain quenching of the emission of those residues more distant from the flavin in Clostridium M P flavodoxin (Fig. 9 ) . Measurements of the fluorescence of the tryptophans in A . vinelandii flavodoxin support the interpretation that quenching is dependent on a single tryptophan, probably located in the vicinity of the F M N (111,131). 4. Circular Dichroism

The measured far ultraviolet CD spectra of the flavodoxins, despite some differences in detail, all provide evidence for the occurrence of highly ordered secondary structure similar to that observed in the Clostridium M P and D. vulgaris molecules (62). In the visible region of the spectrum, free F M N has rather weak circular dichroism; the bands become much more intense in the flavodoxins, and 130. P.S. Song, T. A. Moore, and W. E. Curtin, 2.Naturforsch. B.27, 1011 (1972). 131. L. Andrews, M. L. MacKnight, J. Ryan, and G. Tollin, BBRC 55, 1165 (1973).

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FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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their signs are reversed (14,23,62). The C D spectra of six flavodoxins have in common a positive band a t about 470 nm and a more intense negative band around 370 nm. There are qualitative and quantitative differences between the several flavodoxins, however, and these are particularly marked in the region 380-450 nm. By manual fitting of gaussian curves, Edmondson and Tollin resolved the C D spectra and visible absorption spectra into six vibronic bands, three being associated with each of the two electronic transitions of FMN ( 2 3 ) . [Other data (114,132) suggest the presence of at least seven bands in the visible region.] One C D band, a t about 450 nm, is positive in P. elsdenii and clostridial flavodoxins, but negative in flavodoxins from A . vinelandii, R. rubrum, and D. vulgaris. The distinctive C D spectra and variations in the resolution of the visible spectra (Table IV and Fig. 7) prompted D’Anna and Tollin to divide these flavodoxins into two groups (62). Their division correlated with the ability to bind riboflavin, and additional correlations were proposed. The spectral properties seemed to indicate a more apolar environment, possibly involving more aromatic residues, for the FMN in flavodoxins from D. vulgaris, R. rubrum, and A . vinelandii. Now that the structures of representative flavodoxins from each class appear to verify these predictions, it is tempting to speculate that the FMN binding sites of A . vinelandii and R. rubrum flavodoxins bear a closer resemblance to D. vulyaris than to Clostridium M P flavodoxin. 5. Magnetic Resonance Spectroscopy

Because of their low molecular weights, the ability to exchange the prosthetic group, and the convenience of microorganisms as a source for isotopically substituted proteins, flavodoxins have been favorite subjects for magnetic resonance experiments. a. EPR and ENDOR Spectra of Semiquinones. The electronic structures of model flavin semiquinones have been established by a combination of E P R and optical spectroscopy, utilizing isotopic substitution and alkylated derivatives (117,121). Comparison of the magnetic hyperfine interactions in EPR spectra of flavins variously substituted with 15N or 2H has permitted determination of the spin density distribution in the isoalloxazine nucleus (121). Negligible spin density is found in the pyrimidine ring (positions 1 to 4) ; for the neutral radical, the maximum spin density occurs at N-5, followed in decreasing order by N-10, C-8, and C-6. The magnitude of the coupling constant (about 7.5 G ) for an exchangeable proton in neutral flavosemiquinones leads to assignment of formula (I),with the dissociable proton of pK 8.5 a t N-5 (191).

-

132. G. Scola-Nagelschneider and P. Hemmerich, 2. Naturforsch. B 27, 1044 (1972).

96

STEPHEN 0. MAYHEW AND MARTHA L. LUDWIG

H

O

(1)

In proteins the detailed structure of the EPR spectrum disappears as a result of the slow tumbling of the protein-bound radical. Nevertheless, the coupling of the N-5 proton is sufficiently large to alter the line width of the spectral envelope ( l a 0 ) .Thus, the spectra of neutral semiquinones, such as the flavodoxins, typically have a line width of 19 G, which decreases to 15 G upon exchange with ‘H20 (120).The EPR line widths correlate precisely with the optical spectra in distinguishing neutral from anionic protein semiquinones. The EPR spectrum of fully deuterated flavodoxin from S. lividus in ‘H,O contains some hyperfine structure which allows estimation of the coupling to N-5 and N-10 (133-135). Recently, ENDOR spectroscopy of flavodoxins from P. elsdenii and A . vinelandii has yielded additional information on the electronic structure and geometry of the protein-bound F M N radical. ENDOR signals demonstrate the presence of spin density a t positions C-8, C-6, and N-10 in flavodoxin semiquinones (136,137). Furthermore, the magnitude of the hyperfine coupling to CH, (8), which can be correlated with the ionization state of flavin radicals (137),is consistent with the presence of the neutral radical (1) in A . vinelandii flavodoxin. A signal corresponding to the interaction of the spin density a t N-5 and the N-5 proton was not detected (137).Nevertheless, the ENDOR results, in combination with the EPR spectra of deuterated flavodoxins, indicate that the spin density distribution in flavodoxin radicals is similar to that found in model flavin semiquinones. ENDOR spectra of several flavoproteins haZle been interpreted as signifying that the flavin radical adopts a planar conformation (137). b. N M R Spectra. The NMR spectra of flavodoxins from C . pasteurianum (I%), S. lividus (18), Clostridium MP, and P. elsdenii (99) have 133. H. L. Crespi, J. R. Norris, and J . J . Katz, BBA 253, 509 (1971). 134. H. L. Crespi, J. R. Norris, J. P. Bays, and J . J . Katz, Ann. N . Y . Aead. Sci. 222, 800 (1973). 135. J. S. Hyde, L. E. G. Eriksson, and A. Ehrenberg, BBA 222, 688 (1970). 136. J. Fritz, F. Muller, and S. G. Mayhew, Helv. Chim. Acta 56, 2250 (1973). 137. L. E. G. Eriksson and A. Ehrenberg, BBA 293,57 (1973). 138. C. C. MacDonald and W. D. Phillips, in “Fine Structure of Proteins and Nucleic Acids” (G. D. Fasman and S. N. Timasheff, eds.), p. 1. Dekker, New York, 1971.

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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been reported; for the latter two proteins, spectra of all oxidation states have been recorded. The spectra of the diamagnetic oxidized and fully reduced forms are nearly identical for either Clostridium MP or P. elsdenii flavodoxins, and therefore, for each of these flavodoxins, the conformation cannot be appreciably changed by two-electron reduction. I n the semiquinone state, certain resonances are selectively broadened by the proximity of the flavin paramagnet. However, the general resemblance of the spectra of the radical and diamagnetic forms, along with the X-ray results, argues against any large conformational changes associated with formation of the semiquinone. I n mixtures of partially reduced forms, the spectra appear additive. This observation permits a limit of <50 sec-I to be placed on kerchange, in agreement with values for rates of cornproportionation and disproportionation established by kinetic studies (Section II,F,l). Only certain aromatic resonances and several lines shifted upfield from the aliphatic region are sufficiently well resolved in spectra of P. elsdenii and Clostridium MP fiavodoxins to permit their assignment to individual residues (99). Some of these resonances are not only ring-shifted but also must represent protons in the neighborhood of the flavin ring since they are absent from the spectrum of the semiquinone form; for example, aromatic proton peaks observed at 5.95 and 6.39 ppm in the spectrum of Clostridium MP flavodoxin are upfield from the shifts expected for tyrosine or tryptophan protons and disappear in the presence of the radical. These are likely to be protons of Trp-90. The FMN proton resonances cannot be identified unambiguously in these spectra. Selective proton labeling of deuterated flavodoxin from S. lividus has afforded simplified spectra which are amenable to more detailed interpretation (134) ; for example, these spectra furnish evidence for the presence of two alternative conformations for a leucine residue. Two NMR studies have been specifically concerned with the environment of the protein-bound FMN. Using fully deuterated apoflavodoxin from S. lividus, reconstituted by addition of ['HIFMN, Crespi and coworkers obtained the spectrum of bound oxidized FMN (18). Not all of the peaks could be assigned unequivocally. Changes in the chemical shifts of FMN protons were attributed to the presence of an aromatic residue stacked parallel to the isoalloxazine ring (18,134). The line widths of the ribityl protons and of one aromatic proton were consistent with rigid attachment to the protein, i.e., with the rotational correlation time predicted for flavodoxin (18). The authors ascribed the narrower resonances of the methyl protons and the remaining benzene proton to effects other than rapid motion of the isoalloxazine system relative to the protein, although some motion of the flavin ring is not excluded (134). The

98

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

structural results (Section II,B,3) similarly suggest restrictions on the mobility of the protein-bound flavin. Palmer and Mildvan (139)observed a dramatic effect of the flavin radical of P. elsdenii flavodoxin on the relaxation rate of water protons, denoting exposure of the radical to solvent. I n the crystal structures, the dimethylbenzene end of the isoalloxazine ring is clearly accessible to solvent and several peaks of density near the FMN ring have been interpreted as representing bound solvent molecules (64,667.

E. OXIDATION-REDUCTION POTENTIALS Oxidation-reduction potentials for several flavodoxins are included in Table IV. I n every case, the two one-electron steps prove to be well resolved. The magnitude of the potential difference, E , - El,varies from 0.17 to 0.545 V, depending on the pH and the species of flavodoxin, but E, (oxidized-semiquinone) is always more positive than El (semiquinone-reduced) (140). This behavior distinguishes the protein-bound flavin from free FMN, whose potentials, E , = -0.238 V and El= -0.172 V a t pH 6.95 ( l d l ) , imply that the equilibrium for comproportionation FH2-

+ F H + H+ + 2FH2’

(3)

lies far to the left. In contrast, addition of one reducing equivalent to the flavodoxins produces predominantly the semiquinone. The shifts of potential for the protein-bound flavin, relative to FMN, facilitate measurement of the individual Emidpoint values for the flavodoxins since only two oxidation states are observed during most of the titration. For P . elsdenii flavodoxin, E , was originally determined by titration with NADPH in the presence of catalytic amounts of ferredoxin-NADP+-reductase, using indigodisulfonate as an indicator ; Elwas calculated from the concentration of semiquinone and hydroquinone in equilibrium with excess NADPH in the presence of reductase, or by equilibration with hydrogen and hydrogenase (13,58). Similar procedures, utilizing various redox indicators, as well as potentiometric titrations, have been employed with the other flavodoxins (references in Table IV) . The pH dependence of the redox potentials has been examined, over 139. G. Palmer and A. S. Mildvan, in “Structure and Function of Oxidation Reduc-

tion Enzymes” (A. Akeson and A. Ehrenberg, eds.), p. 385. Pergamon, Oxford, 1972. 140. The potential for the semiquinone-reduced couple in flavodoxins is designated El to be consistent with the assignments in FMN, where El is the higher potential but refers to the semiquinone-reduced couple. 141. R. D. Draper and L. L. Ingraham, ABB 125, 802 (1968).

TABLE IV DATAFOR FLAVODOXINS Redox potentials

MW of Source

holoprotein

C . pasteurianum

14,600

Absorption maxima nm (mM-1 cm-1) 272(40) 272(45.8)

372(7.9) 374(8.47)

443(9.1) 443(10.4)

P . elsdenii

15,0005

272 (47.6)

377(8.75)

445(10.2)

Clostridium M P

1.5,800a

272(46.8)

376(9.1)

445(10.4)

pH

7 8 7 8 7

8

D. vulgaris D. gigas R. rubrum A . vinelandii

16, 300a 16,000 22,800 23,000

E. coli A . nidulans

14,500 20,300

273(48) 273(47) 272(54.2)

375.5(8.7) 374(8.2) 376(11.3) 371(9.5)

456.5(10.7) 456.5(10.2) 460(11.2) 450(10.6)

274(50) 275

370(6.6)

377

467(8.25) 465(9.2)

270

377

465 (10.2)

7.8 -

8.2 7.7 7.7

7 8

S. lividus C . fusca FMN

17,000 22,000

-

7 275(54.6) 266(31.8)

379(9) 373(10.4)

464(10 rt 0.2) 445( 12.5)

-

7

Ei (mV)

-419 -429 -372 - 375 - 399 -408 -440 -

-495 -464 -410 -447 -450 -

-450 - 172

Ef

(mV)

- 132 - 192 -115 - 175 - 92 - I52 - 150 -

-

+50

References 6-8 13 12,13,58 13

11,100 11

14 22,61,100

-270 -240 -221 -281 - 50 -

-238

144 16 &,IS5

51 17,18,1S1 15 141

~

Calculated from the amino acid sequence. The remaining values are determined from amino acid compositions (27).

co co

100

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

a limited range of pH, for P . elsdenii (68),C. pasteuhnum (1S11@), Clostridium M P (IS), and A. nidulans (186) flavodoxins. The results are consistent with the formulation of the reactions at pH 8 as

where the oxidieed and semiquinone flavins are neutral and the reduced form exists as the anion. As expected from the failure to observe any ionization of the oxidized or semiquinone FMN in P. elsdenii flavodoxin over the pH range 5.8 to 9.2 by spectroscopic techniques, the slope of the line relating E , to pH is -0.059 V in accord with the addition of a proton and an electron in reaction (4). Values of E, are independent of pH in the high pH region but become pH dependent at the lower extreme (Fig. 10) (143).The change in slope corresponds to the titration of a redox linked group with a pK of 5.8 in fully reduced P. elsdenii flavodoxin (68) and a pK of 6.7 in C. pasteurianum and Clostridium M P flavodoxins ( I S ) . These pK values have been identified with the formation of the hydroquinone anion ; in model flavins, this ionization occurs a t N-1 with a pK of 6.7 (181,146). A similar ionization, with pK = 7.0, has been proposed for A . vinelandii flavodoxin (B), implying that the pH dependence of El for this flavodoxin is probably similar to those shown in Fig. 10 (143). Because of the dependence of E2on pH, the equilibrium constant for comproportionation changes with pH; for example, a t pH 6.7 the semiquinone formation constant [K = (FH2')2/(FH)(FH2- FHa)] for P . elsdenii flavodoxin is 43,000,while a t pH 9.3 it decreases to 110 (b8). As noted in Table I11 (Section II,C,%), changes in the reduction potentials of FMN upon combination with the apoprotein require that the association constants for FMN binding vary with oxidation state. Thus, the effect of the protein on the redox potentials can equally well be described in terms of the variable affinity of the protein for each redox species of FMN. The available structural and spectroscopic data have led to some hypotheses regarding the molecular basis for the shifts in

+

142. The redox potential first reported for this protein ( 8) was based on extinction measurements at 443 nm which were assumed to change linearly during reduction; the changes are not linear because of semiquinone formation and the calculation was therefore in error. 143. Failure to consider the different effects of pH on Eland E, has led to inaccurate comparisons of the redox potentials of flavodoxins from different sources (18,100,14464).

144. D. C. Yoch, BBRC 49, 336 (1972). 145. H. J. Lowe and W. M. Clark, JBC 221, 983 (1956).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

101

-0.3

-0.4 -

I

I

I

I

5

6

7

1

a

I

9

PH

FIG.10. Effect of pH on the redox potentials of flavodoxins. (I, 0 ) CEostridium MP flavodoxin, (E, 0 ) C. pasteurianunz flavodoxin, and (0, 0, A ) P . elsdenii flavodoxin. From Mayhew (13).

K , (69). Conformational changes accompanying reduction to the semiquinone state, suggested by electron density maps of Clostridium M P flavodoxin, are consistent with the tighter binding of F M N semiquinone. An additional hydrogen bond between F M N and the protein and the rearrangement of atoms in the bend 56-59 to produce a lower energy conformation would both act to increase the relative stability of the semiquinone complex. However, the existence of these conformational changes remains to be conclusively established and they are unlikely to occur in all flavodoxins (Section II,B,3,e). The oxidized-semiquinone potential, E,, is highly dependent on the source of the flavodoxin and does not seem to correlate well with the classification of flavodoxins according to their spectral properties (Table IV). Hence, examination of the structures of the semiquinone state of D. vulgaris or other flavodoxins may reveal additional mechanisms for stabilization of the semiquinone form. The strength of nonbonded interactions between isoalloxazine and the adj acent tryptophan, tyrosine, or methionine residues may also vary with

102

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

oxidation state. Draper and Ingraham (146) found that the extent of complex formation between riboflavin and tryptophan, tyrosine, or glutathione decreases in the order semiquinone > oxidized > fully reduced. The available X-ray data for fully reduced Clostridium M P flavodoxin indicate that the isoalloxazine ring is nearly planar in the reduced protein (69). The spectra of reduced flavodoxins from Clostridia and P. elsdenii also favor the existence of a planar ring, and the fluorescence of reduced flavodoxins is compatible with the restriction of the bending modes typical of reduced isoalloxazines (124). Free reduced flavins, on the other hand, are bent along the N-5:N-lO direction, with the extent of bending dependent on the nature of ring substituents (89),and they undergo rapid ring inversion (147). It is reasonable to suppose that the decrease in K , for the reduced state results in part from constraints imposed by the protein on the conformation and mobility of the dihydroisoalloxazine ring. The relative destabilization of the reduced FMN-protein complex is presumably the physiologically most significant thermodynamic role of the protein since its effect is to lower E l to the range of potentials characteristic of the ferredoxins ( 1 0 0 ~ ) .

F. REACTIVITY 1. Cornproportionation In contrast to certain other flavoproteins ( I 48), most flavodoxins comproportionate readily in the absence of mediators (61,68). I n P. elsdenii flavodoxin, the rate of reaction (3) depends on pH and ionic strength, varying a t pH 8.5 from 100 M-’ min-I a t zero ionic strength to almost lo6 M-’ min-’ a t I = 0.24 (1.69). This marked dependence on ionic strength probably reflects the large net negative charge on the protein. The reverse, or disproportionation, rates are smaller for flavodoxins, since the equilibrium in reaction (3) favors the semiquinone (68). A rate constant of 5 X lo3 M-l min-1 has been determined for the disproportionation of A . vinelandii flavodoxin a t pH 11; a t pH 8.3 in 0.05 M pyrophosphate the constant for P . elsdenii flavodoxin is only 47 M-I min-’ (68). Not surprisingly, rates for the comparable reactions of free flavins are much greater. Disproportionation of neutral FMN radicals proceeds with a rate constant of the order of lo5 M-I sec-1 (160). To obtain valid redox 146. R. D. Draper and L. L. Ingraham, ABB 139, 265 (1970). 147. L. Tauscher, S. Ghisla, and P. Hemmerich, Helv. Chim. Acta 56, 630 (1973). 148. V. Massey, in “Flavim and Flavoproteins” (H. Kamin, ed.), p. 231. Univ. Park Press, Baltimore, Maryland, 1971. 149. S. G. Mayhew and V. Massey, BBA 319, 181 (1973). 150. S. P. Vaish and G. Tollin, Bioenergetics 2, 61 (1971).

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FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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potentials for the flavodoxins, it is, of course, essential that the comproportionation reaction attain equilibrium during titrations, but that may be accomplished by addition of mediators if necessary. 2. Sodium Dithionite The individual flavodoxins vary in their reactivity with dithionite and occasionally display peculiar behavior upon titration with this reagent. In view of the wide usage of dithionite as a reducing agent for redox proteins, its reactions with the flavodoxins will be described in some detail. Flavodoxins from P. elsdenii and Clostridium MP react rapidly with equimolar dithionite at p H 7, and full reduction is achieved within a few minutes (IS).The kinetics of the reduction of P. elsdenii flavodoxin a t p H 8.5 have been investigated by stopped-flow spectrophotometry at ionic strengths where comproportionation does not complicate the results (149). The reduction of both oxidized and semiquinone forms of the protein is first order with respect to protein concentration, but the semiquinone is reduced much more rapidly than is the oxidized form. The reaction rates depend on the square root of dithionite concentration. From these observations, Mayhew and Massey (149) concluded that P. elsdenii flavodoxin is reduced by the dissociation product of dithionite, SOZ', and that full reduction of the oxidized protein involves two sequential one-electron transfers according to the following scheme: K1 szo2- s 2SOZ'

Oxidized flavodoxin Flavodoxin semiquinone

+ SO2' + SO1'

+ SO2 f flavodoxin hydroquinone + SO1 k 2 flavodoxin semiquinone k

(6)

(7) (8)

The ratio of the rate constants, k z / k l , is approximately 450; consequently, it is not possible to detect the semiquinone during reduction of the oxidized protein by an excess of dithionite. The values of kl and kz depend on determination of K 1 ; use of the value given by Lynn et al. (161) leads to the estimate that kz is about 4 x lo7 M-l sec-1. I n a similar study of the dithionite reduction of additional redox proteins and small molecules, Lambeth and Palmer (166)found that one-electron reduction by SOZ' is the dominant pathway in several other reactions. Although ffavodoxin from P. elsdenii is fully reduced by one mole of dithionite per mole of flavin a t pH 7 and above, this stoichiometry does not obtain at lower pH; for example, a t pH 5.2,where dithionite is still reasonably stable under anaerobic conditions, about 12% of the flavin 151. S. Lynn, R. G. Rinker, and W. H. Corcoran, J. Phys. Chem. 88,2363 (1964). 152. D. 0. Lambeth and G . Palmer, JBC 248, 6095 (1973).

104

STEPHEN Q. MAYHJGW AND MARTHA L. LUDWIG

remains as the semiquinone in the presence of a twelvefold excess of the reductant (149).Further studies of the effect of pH on the reduction of this protein by dithionite strongly suggest that a pH-dependent equilibrium, described in part by Eq. (8) and subsequent hydration of SO,, is established (153).The ability of dithionite to achieve full reduction of the protein a t high pH is explicable if the potential of the dithionite redbx system remains pH-dependent and continues to decrease in the region where E , for flavodoxin becomes pH-independent. At pH 5, the redox potential of the dithionite acceptor-donor is probably similar to El for P . elsdenii flavodoxin (-323 mV) ; at pH 8, where El is -375 mV and independent of pH, the potential of the dithionite system is evidently more reducing. At pH 7 and p H 8.3, flavodoxin from C. p a s t e ~ T ~ u nbehaves u~ somewhat like P. elsdenii flavodoxin a t p H 5.2. Addition of one equivalent of dithionite generates the semiquinone as expected, but further additions do not produce stoichiometric yields of reduced protein. Some side reactions which consume dithionite may intervene (13). Flavodoxins from A . nidulans, A . vinelandii, and R. rubrum are much less reactive with dithionite than are the clostridial or P . elsdenii flavo; methyl viologen can be used to mediate the doxins (19-22,39,51,125) reduction (100). Reduction of R . rubrum and A . vinelandii flavodoxin is incomplete a t pH 7 even in the presence of large excesses of the reagent (14,102). Full reduction of A . vinelandii flavodoxin can be achieved a t pH 8 with an excess of dithionite or alternatively a t pH 7 if the ionic strength is raised [e.g., in 3 M ammonium sulfate ( I O S ) ] . Tollin and co-workers (100,lOS) have proposed that the neutral flavin hydroquinone cannot be accommodated by the protein and that the extent of reduction is therefore controlled by the ionization of the reduced flavin (pK = 7.0). They attributed the effect of ionic strength to a shift in this pK. An alternative explanation would invoke the pH dependence of the dithionite system as described above. The higher pH values and dithionite concentrations required for full reduction of the A . vinelandii protein would then be a result of its very low E l ;the ionic strength effect would have to be ascribed to a salt-dependent increase in this potential. 153. After P . ekdenii flavodoxin is fully reduced by ti stoichiometric amount of dithionite at p H 8, anaerobic addition of acid to pH 5 converts half of the protein to the semiquinone, with a concomitant increase in absorption at 329

nm, an isobestic point for the protein, but a wavelength where dithionite absorbs. These changes can be reversed by raising the pH. The extent of reduction of the protein is influenced by the concentration of dithionite or bisulfite but not by sulfate or thiosulfate (164) 154. E. J. F. van Arem and S. G. Mayhew, unpublished.

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3. Oxygen and Ferricyanide

Oxidation of reduced flavodoxins by molecular oxygen is a nonphysiological reaction; in contrast to the flavoprotein oxidases and hydroxylases, the flavin-containing dehydrogenases presumably do not reduce oxygen during in vivo catalysis. The oxidation of dehydrogenases (and flavodoxin) by oxygen in vitro differs in a characteristic manner from the oxygen reactions of flavin-containing oxidases and hydroxylases (63). Flavodoxin is an interesting case since the reactivity of the fully reduced protein with oxygen is similar to that of model flavins, whereas the reactivity of the semiquinone is orders of magnitude less than that observed for comparable models. Kinetic studies have shown the following reactions to be significant in the reoxidation of reduced flavodoxin (165,156) :

The reactions of model flavins with oxygen proceed via intermediate flavin-02 adducts (167-159). Whether the reactions of flavodoxin with 0 2 involve similar adducts is not known. Determination of the kinetic parameters is complicated by reaction (3) and b y disproportionation of OaT to yield oxygen and peroxide. Furthermore, oxidation of both the anionic and neutral species of the fully and partially reduced flavins must be considered. [Equations (9)-(12) show only the forms predominating at neutral pH.] Stopped-flow spectrophotometry with P. elsdenii flavodoxin (166)has shown that the semiquinone is formed directly b y oneelectron oxidation of the hydroquinone since the rate of semiquinone formation is too great to be attributed t o full oxidation followed by comproportionation. The contribution of reaction (10) has been assessed by addition of superoxide dismutase to catalyze breakdown of 0 2 ’ . I n the presence of this enzyme, the rate of formation of the semiquinone a t pH 7.2 is decreased (Fig. 11). Decay of the semiquinone absorbance in 155. D . P. Ballou, Ph.D. Thesis, University of Michigan, Ann Arbor, 1971. 156. D. P. Ballou and S. G. Mayhew, unpublished. 157. Q . H. Gibson and J. W. Hastings, BJ 68,368 (1962). 158. V. Massey, G. Palmer, and D. P. Ballou in “Oxidases and Related Redox Systems’’ (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 25. Univ. Park Press, Baltimore, Maryland, 1973. 169. V. Massey, G. Palmer, and D. P. Ballou, in Flavins and Flavoproteins” (H. Kamin, ed.), p. 349. Univ. Park Press, Baltimore, Maryland, 1971.

106

STEPHEN G . MAYHEW AND MARTHA L. LUDWIG

"

0

1

2 3 Time lsecl

L

FIG.11. Effect of superoxide dismutase on the reaction of reduced flavodoxin with oxygen. P . elsdenii flavodoxin (3.5x lo-' M in 0.05 M K phosphate buffer, pH 7.5) was titrated to full reduction with sodium dithionite (13) and then mixed in a stopped-flow spectrophotometer (149) with oxygen-containing buffer (0.15 M K phosphate, pH 7.2, and 6.26 x lo-' M 0 2 , curves 1 and 2; 0.15 M Na pyrophosphate, p H 9, and 2.5 x M 02,curves 3 and 4). Bovine erythrocyte superoxide dismutase was present in the experiments of curves 1 and 3.

the absence of dismutase (Fig. 11) provides evidence for reaction of 02' with the semiquinone [reaction (12)]. Reaction of the products of reaction (9), in a second one-electron step, is an interesting aspect of the oxidation scheme and may indicate that O 2 and 02' react at different sites. A minor pathway involving two-electron transfer has not been rigorously excluded. The rate constant for oxidation of P . elsdenii flavodoxin hydroquinone to the semiquinone at room temperature, in the presence of superoxide dismutase, is 3.5 x l o 4 M-1 sec-l and does not vary significantly with pH between 7 . 2 and 9.0 (155,156). The lack of pH dependence is consistent with a pK of 5.8 for formation of the anionic form of fully reduced P. elsdenii flavodoxin (58) (Section 11,E). For reoxidation of tetraacetylriboflavin a t pH 8.4 in the presence of dismutase, the rate constant is estimated to be ca. 1 X 104 M-I sec-l (158,169).For some other flavoprotein dehydrogenases, the comparable rates are of the order of lo2 RII-' sec-I (63).Because of the difficulty of determining the concentration of 02' in the reaction mixtures (168-160), constants for reaction (10) can160. V. Massey, s. Strickland, s. G. Mayhew, L. G. Howell, P. C. Engel, R. G. Matthews, M. Schuman, and P. A. Sullivan, BBRC 36, 891 (1909).

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not readily be evaluated, but it is evident from Fig. 11 that the secondorder rate constant for reaction (10) a t p H 7.2 must be larger than that for reaction (9). At p H 9, oxidation by 02' appears t o be slower than a t p H 7.2. For oxidation of the semiquinone, the rate constants vary considerably among the flavodoxins and in addition depend on oxygen concentration (58) and p H (58,102). At p H 7, k,,, for oxidation of the semiquinones of P. elsdenii or Clostridium MP flavodoxin is approximately 7 M-' min-', about 20 times greater than the value measured for A . vinelandii flavodoxin (102). [The contributions of reactions (11) and (12) have not been separated in these determinations.] All of the protein semiquinones are more rapidly oxidized a t higher pH, an effect attributed t o the presence of increasing concentrations of the more reactive semiquinone anion species (150).Edmondson and Tollin found a linear relationship between the reciprocal of the rate and the hydrogen ion concentration for oxidation of A . vinelandii flavodoxin radical (102).Assigning a p K of 11.5 for the ionization of the bound FMN radical, they calculated a rate constant of 2.4 X lo3 M-' sec-' for oxidation of the anion. Corresponding plots for P. elsdenii flavodoxin are not linear, and there are other indications that the pH effects with this protein are more complicated (155). A significant fraction of this semiquinone is oxidized via reaction (12), and the relative rates of dismutation of 02' and of its reaction with the protein may be very pH-dependent. Despite such individual variations, the flavodoxin semiquinones are all far less reactive with oxygen than might be predicted from models. The rate constants for reaction of oxygen with FMN or FAD radicals, generated by flash photolysis (150,161), are los M-' sec- for the anion and 104 M-1 sec-' for the neutral species ( 1 6 1 ~ ) . The factors which control the reactivity of fully reduced flavoproteins with oxygen are not completely understood. Hemmerich (16'2) has proposed that reactivity reflects hydrogen bonding a t N-1 or N-5 of the flavin and that the more planar the reduced flavin, the more readily will it be oxidized. Detailed structure analyses of fully reduced flavodoxins may suggest additional explanations for the effect of the protein on flavin reactivity. Oxidation of flavodoxins by ferricyanide probably provides a more appropriate model for the in vivo electron transfer reactions than does oxidation by oxygen. The ferricyanide and oxygen reactions respond differently to substitutions in the flavin ring (150,16'1),and the two reagents may react a t different sites in the isoalloxazine system. The oxidation of reduced flavodoxin by ferricyanide has been reported to be "very 161. J. M. Gillnrd and G. Tollin, BBRC 58, 328 (1974). 161a. R. A. Lazaarini and A. San Pietro, BBA 62, 417 (1962).

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STEPHEN G . MAYHEW AND MARTHA L. LUDWIG

fast” (63); rate constants for oxidation of flavodoxin semiquinone by ferricyanide are of the order of 104 M-1 sec-1 (63).Free neutral lumiflavin radical is oxidized much more readily with a constant of 5 X lo8 M-l sec-1 (161). Gillard and Tollin (161) have found the rate of transfer from lumiflavin radical to ferricyanide (and to oxygen) to be decreased, by a factor of 10 or less, in the presence of tryptophan concentrations sufficient for complex formation. They suggested that the tryptophan-FMN interaction in flavodoxins may be partly responsible for the lower reactivity of protein-bound FMN. Conformational changes accompanying the reaction could make a major contribution to the activation energy for oxidation of the semiquinone (Section II,C,5). 4. Redox Proteins

No complete kinetic studies of electron transfer between flavodoxin and individual acceptor proteins have been reported. While it has been assumed from the values of the redox potentials (Table IV) that flavodoxins function in vivo as one-electron carriers, alternating between the semiquinone and hydroquinone states, definitive evidence for this mechanism is not easy to obtain. Flavodoxins could in principle mediate transfer from either one- or two-electron donors to one- or two-electron acceptors over a wide range of potentials. Of the known flavodoxin-dependent reactions, the best characterized are reduction of ferredoxin-NADP+-reductase (FNR) , reduction of nitrogenase, and transfer of electrons between FNR and cytochrome c. Reduction of mammalian cytochrome c by reduced F N R does not proceed unless flavodoxin or ferredoxin is present (48,161~).Both mediators are known to form molecular complexes with F N R (48-46), and electron transfer between ferredoxin and FNR has been demonstrated (163). Under aerobic conditions the reduction of cytochrome c catalyzed by flavodoxin is partially inhibited by superoxide dismutase (160).Thus cytochrome c is reduced by 02’ in the presence of oxygen, but a direct reaction between flavodoxin and cytochrome c must also occur. For the overall reaction, NADPH + cytochrome c, turnover numbers have been determined for F N R but not for the mediators (164). Hence, the efficiency of flavodoxin in catalysis cannot yet be compared with the transfer rates in reactions involving dithionite, oxygen, or ferricyanide. 162. P. Hemmerich, A. P. Bhaduri, G. Blankenhorn, M. Briistlein, W. Haas, and W.-R. Knappe, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.) p. 3. Univ. Park Press, Baltimore, Maryland, 1973. 163. J. Siedow, unpublished. 164. G. Forti and E. Sturani, Eur. J. Biochem. 3, 461 (1968).

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An EPR signal attributable to flavodoxin semiquinone can be detected in fully deuterated cells of S. Zividus. The concentration of radical increases in the dark and diminishes when electron flow is initiated by illumination (165). These observations suggest that the principal oxidation states of flavodoxin in photosynthesizing systems are the fully reduced and semiquinone forms. However, they do not demonstrate unequivocally that flavodoxin acts as one-electron carrier in the photosynthetic chain. Yates (36) has studied the reaction of flavodoxin from A . chroococcum with purified nitrogenase. Substrate amounts of reduced flavodoxin are converted to the semiquinone by catalytic quantities of nitrogenase in the presence of nitrogen and an ATP-generating system. Reduction of nitrogenase substrates requires fully reduced flavodoxin and will not proceed if only the semiquinone form is added. As in the photosynthetic reactions, these results appear to establish the kinetic importance of the semiquinone. Formation of the flavodoxin radical seems too rapid to be ascribed to two-electron oxidation and subsequent comproportionation, but additional experiments may be necessary to exclude the two-electron pathway. Determination of the precise mechanism of catalytic electron transfer by flavodoxins remains a challenge. The structure analyses suggest some restrictions on allowed mechanisms. Since the disposition of side chains adjacent to the isoalloxazine ring is so different in D. vulgaris and Clostridium MP flavodoxins, it is difficult to envision a universal pathway of electron transfer which utilizes these residues (Section II,B,3). The simplest suggestion offered by the structures is that the redox reactions involve the dimethylbenzene moiety of the flavin, which appears readily accessible for direct electron transfer. With the wealth of data on the structure and properties of the flavodoxins, the time seems ripe for incisive investigations of the oxidation-reduction reactions. 111. Electron-Transferring Flavoprotein

A. INTRODUCTION In the second edition of “The Enzymes” Beinert (1) described the properties of electron-transferring flavoprotein (ETF), an enzyme from mammals and mycobacteria which mediates the transfer of electrons from acyl-CoA and sarcosine dehydrogenases to the mitochondria1 cytochrome chain and also to nonphysiological electron acceptors. The mammalian enzyme is not readily obtained in high purity (1) and perhaps as a conse165. J.

R. Norris, H. L. Crespi, and J . J. Kata, BBRC 49, 139 (1972).

110

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

quence only a few further studies on its molecular properties have been reported (166-168). This section is therefore concerned mainly with a bacterial flavoprotein which has recently been isolated from P. eZsdenii (169,170) and whose function resembles that of mammalian ETF. Peptostreptococcus elsdenii excretes short-chain fatty acids (C, to C,) into the growth medium during the fermentation of lactate and carbohydrates (54,171). The final reductive step in the synthesis of butyrate, the conversion of crotonyl-CoA to butyryl-CoA, is catalyzed by a flavoprotein (172,173) that is closely similar to butyryl-CoA dehydrogenase (BCD) of mammals. Electrons for this reduction can be provided by lactate and two further flavoproteins. One is a D-lactate dehydrogenase (pyridine nucleotide independent) in which FAD is the only known chromophore (174,175); the other is an enzyme which mediates electron transfer from D-lactate dehydrogenase to BCD (174,175). Although this mediator enzyme is concerned in the synthesis of fatty acids rather than their oxidation, it is functionally similar to ETF of mammals, and accordingly the term “electron-transferring flavoprotein” has been extended to include it (170,175) . NADH

D-

Lactate Lactate dehydrogenase D-

-

t

ETF-BCD

Crotonyl- CoA

I

(13)

BUtyWl-CoA

Peptostreptococcus elsdenii ETF also oxidizes NADH which therefore serves as an alternative source of electrons for the reduction of BCD and nonphysiological acceptors such as 2,6-dichlorophenolindopheno1 (170,17$,175). A similar diaphorase activity is associated with preparations of ETF from mammalian sources (166,176), but it has been attributed to contamination by other enzymes (1,177). 166. 167. 168. 169.

D. D. Hoskins and R. A. Bjur, JBC 240, 2201 (1965). D. D. Hoskins, JBC 241, 4472 (1966). C. L. Hall, Fed. Proc., Fed. Amer. SOC.Ezp. Biol, 32,596 (1973). C. D. Whitfield, S. G. Mayhew, and V. Massey, Fed. Proc., Fed. Amer. SOC. Ezp. Biol. 31, 447 (1972).

170. 171. 172. 173. 174.

C. D. Whitfield and S. G. Mayhew, JBC 249, 2801 (1974). S. R. Elsden and D. Lewis, BJ 55, 183 (1953). R. L. Baldwin and L. P. Milligen, BBA 92, 421 (1964). P. Engel and V. Massey, BJ 125, 879 (1971). H. L. Brockman and W. A. Wood, Fed. Proc., Fed. Amer. SOC. Exp. Biol. 29,

862 (1970). 175. H. L. Brockman, Ph.D. Thesis, Michigan State University, E. Lansing, 1971. 176. F. L. Crane and H. Beinert, JBC 218, 717 (1956). 177. W. R. Frisell, J. R. Cronin, and C. G. Mackenrie, in “Flavins and Flavoproteins” (E. C. Slater, ed.), p. 367. Elsevier, Amsterdam, 1966.

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B. MOLECULAR PROPERTIES 1. Purification, Molecular Weight, and Amino Acid Composition Highly purified preparations of ETF are obtained from extracts of P. elsdenii by a combination of ion-exchange chromatography on DEAEcellulose, salt fractionation with ammonium sulfate, and gel filtration in Sephadex G-100 (170).I n some preparations, ETF is associated with a larger unidentified flavoprotein and is separated from this protein only during the final purification step. Purified ETF is stable for long periods in frozen solution at -200. The molecular weight of the enzyme, determined by gel filtration in calibrated columns of Sephadex, is about 73,500. The protein is composed of two nonidentical subunits which can be separated by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate. From their electrophoretic mobilities and a comparison with standard proteins, the molecular weights of the two subunits were estimated to be 41,200 and 33,200. Amino acid and flavin analyses gave a minimum molecular weight of 37,800, a value which is in close agreement with the average molecular weight of the two subunits. The subunits evidently form a tight complex since strong denaturing agents such as sodium dodecyl sulfate or guanidine HC1 are required for their dissociation (170). Early estimates of the molecular weight of mammalian ETF ranged from 30,000 to 70,000 (176). A more recent determination with protein from pig liver gave a molecular weight of 35,000 (168). The amino acid composition of the bacterial protein has been determined and found to be unusually low in tryptophan (170). 2. Properties of the Flavin Chromophore

Purified preparations of P. elsdenii ETF contain about 1.4 moles of flavin per mole of protein, but they bind additional FAD to give a total of 2 moles per mole of protein (170); evidently some flavin is lost from the protein during its isolation. It is not known whether the two flavins are bound to the same or different subunits. Although FAD is the major chromophore in most preparations of the enzyme, all preparations contain at least traces of two modified flavins ( 170,178). When a protein-free extract of the enzyme is fractionated on a column of DEAE-cellulose, three colored bands are separated. In order of their elution from the column, the bands are yellow (FAD), green, and orange. After further purification, the chromophores in the green and orange bands were characterized and identified by comparison 178. C. D. Whitfield and S. C. Mayhew, JBC 249, 2811 (1974).

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STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

TABLE V SPECTROSCOPIC PROPERTIES OF 6- AND 8-OH FLAVINS ~~

~~

.A.,

Compound

pH

(4

nm (mM-l cm-l)

PK

422(19,6) 7.1 323(19.6) 427(22.6) 600(3.41) 420(21.2) 6-OH-FMN 5.5 7.1 320(23) 423(24.9) 600(3.45) 9.0 asO(60.5) 446(26.9) SOH-FAD 3.1 4.8 6.7 252(80) 263sh(58.5)' 300(15.1) 478(36.7) 262(31) 435(26) SOH-FMN 3.0 4.8 7.0 252(51) 267sh(25.5)" 300(10.7) 472(41) 6-0H-FAD

5.6 9.0

262(50.5) 260(51.7)

Here, sh refers to a shoulder.

with chemically synthesized model flavins (178-183). The green chro(ribityl-5'-ADP) 4soalloxazine mophore is 6-hydroxy-7,8-dimethyl-10(181,183,184) and the orange chromophore is 7-methyl-S-hydroxy-10(ribityl-5'-ADP) -isoalloxazine (179,I8O,185,186), The 6-hydroxyl and 8-hydroxyl groups are in direct conjugation with the chromophoric system and cause drastic changes in the spectrum of FAD; moreover, each of them gives rise to an additional pK in the flavin (Table V) . The spectra of both compounds are notable for their very intense peaks in the 400-480-nm region and, in the case of the anion of 6-OH-FAD, for a broad band of absorption centered at 600 nm. The proportions of the hydroxyflavins in ETF vary in different preparations of the enzyme (5-30 and 1-35% of the total flavin for 6-OH-FAD and 8-OH-FAD, respectively). At the highest levels observed, they together comprise about 50% of the total flavin (178). Such preparations differ in a number of properties from preparations which contain mainly FAD and only traces of the two modified flavins. First, and as detailed later, they have markedly different spectroscopic properties. Second, they 179. S. G. Mayhew and V. Massey, BBA 235, 303 (1971). S. Ghisla and S. G. Mayhew, JBC 248, 6568 (1973). 181. S. G. Mayhew, C. D. Whitfield, 9. Ghisla, and M. SchumanJorns, Eur. J . Biochem. 44, 579 (1974). 182. G. Schollnhammer and P. Hemmerich, Eur. J. Biochem. 44, 561 (1974). 183. G. Schollnhammer and P. Hemmerich, 2. Naturforsch. B 27, 1030 (1972). 184. A similar flavin but a t the level of FMN has been found in glycollate oxidase 180.

from pig liver (181),

185. N. A. Polyakova, L. S. Tul'chinskaya, L. G. Zapesochnaya, and V. M. Berekovskii, Zh. Obshch. Khim. 42, 465 (1972). 186. R. Addink and W. Berends, Tetrahedron 30, 75 (1974).

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fail to reduce BCD although they catalyze the oxidation of NADH with 2,6-dichlorophenolindophenol as electron acceptor. Third, they have a higher electrophoretic mobility and isoelectric point and can therefore be separated from ETF by electrophoresis or isoelectric focusing (178). Initially it was concluded that the modified flavins are associated not with ETF but with a different enzyme which also displays NADH dehydrogenase activity (179). However, more recent observations (178) have suggested that the protein moiety to which 6-OH-FAD and 8-OH-FAD are bound is very similar to apo-ETF, and that the different properties of the proteins separated by electrophoresis simply reflect differences in the proportions of the three flavin prosthetic groups. The molecular weight and subunit composition of the protein in the two bands was found to be the same; their amino acid compositions are very similar; protein with a high level of the hydroxyflavins shows a reaction of identity with antibody prepared against E T F ; and complexes made from apo-ETF and the separated hydroxyflavins catalyze the oxidation of NADH but they fail to reduce BCD (178,181). The mechanism of formation of the modified flavins is not known. However, i t appears that they are not present i n vivo in P. elsdenii since they are not detectable in crude extracts of the organism until after exposure of the extracts to oxygen (187), and it therefore seems likely that they are formed in ETF during the purification procedure. Attempts to generate them from FAD in the purified enzyme have failed. About 96% of the flavin of P . elsdenii ETF can be dissociated by chromotography of the holoprotein on a column of Sephadex G-25 equilibrated with guanidine HCI (170).After removal of the guanidine HC1, the apoprotein binds FAD with full restoration of catalytic activity (units per mole of bound flavin) ; it does not bind FMN. When freshly prepared, the apoprotein binds 2 moles of FAD per mole of protein ( K , = 5 X lo6 M-l, pH 6.2 and 16O). However, it is unstable and the amount of flavin which can be bound decreases progressively during storage a t 4 O or -2OO. The apoprotein also binds the hydroxyflavins ( K , = 6.7 X los M-' for 6-OH-FAD) and causes marked changes in their visible and near UV spectra (178,181).The spectra of the complexes of apo-ETF with the three separate flavins were used to determine the flavin composition of native ETF. Unlike normal flavins, the chromophores of 6-OH-FAD and 8-OH-FAD contain groups which ionize in the physiological pH range, and whose pK values can be studied after binding of the flavins to apoflavoproteins. When 6-OH-FAD is bound by apo187. C. D. Whitfield, private communication; I. Dekker and S. G . Mayhew, unpublished observations.

114

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

WMLENGM. nm

FIG.12. Absorption spectra of ETF. Curve 1, oxidized enzyme as isolated; curve 2, isolated enzyme after reduction with dithionite and reoxidation with potassium ferricyanide ; curve 3, enzyme fully reduced with dithionite.

ETF, the pK at 7.1 of the free flavin is decreased, and, in addition, the pH titration curve for the complex extends over a range of pH that is wider than expected for the ionization of a single group. It has been proposed that these effects might result from a positively charged group (pK 5-7) in ETF that is close to N-l:C-2:0-2of the flavin (181). These and similar observations with other apoflavoproteins (179-181,188) indicate that the hydroxyflavins can be used as reporter groups for the flavin binding sites of flavoproteins to obtain information not readily available from studies with unmodified FAD and FMN. As will be evident from the previous discussion, the absorption spectrum of ETF depends on the mixture of flavins in a particular preparation. Most preparations have a low content of the modified flavins and display absorption maxima at 275, 375, 450, and 660 nm, with relative intensities of 5.9:0.81:1 :0.01,respectively (Fig. 12). The band a t 660 nm is the result of a small amount of 6-OH-FAD; preparations with a higher content of this flavin have more absorption a t 660 nm and addi188. A similarly charged group in the flavin binding site of glycollate oxidase, but with a higher pK, has been inferred from the effects of pH on 6-OH-FMN-apoglycollate oxidase (181). In contrast, a negatively charged group near the flavin of flavodoxin, as observed in X-ray crystallographic studies (64,661, would account for the increase of p K observed when 6-OH-FMN and %OHF M N are bound by apoflavodoxin (180,181).

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FLAVOPROTEINS

115

tional peaks a t 350 and 430 nm (181).In contrast, the absorption spectrum of preparations of the enzyme with a high content of 8-OH-FAD has a relatively intense peak a t 475 nm, marked shoulders on both sides of this peak, and further peaks at 318, 356, and 375 nm (178-180) ; such preparations are orange colored rather than yellow. The spectrum of ETF with FAD as the predominant flavin is altered when more FAD is added t o saturate a11 of the flavin binding sites (approximately 0.6 mole of added FAD per mole of protein). A large increase of absorption occurs around 400 nm, and the separation of the peaks a t 450 and 375 nm becomes correspondingly less distinct (170).Holoprotein prepared from apo-ETF and FAD shows a similar high absorption at 400 nm, and the extinction coefficient a t 450 nm of the newly bound flavin is low (10,500 M-l cm-l) by comparison with that of the FAD present in the enzyme as it is isolated (12,500 M-l cm-l). Reduction of the isolated enzyme and subsequent reoxidation also leads to a change in the spectrum, and again the most obvious effect is a large increase in the absorption a t 400 nm. The explanation for these changes is not known. The flavin prosthetic group of mammalian ETF is also FAD (1,168,189).Crane and Beinert (176) reported that the pig liver enzyme contains 1 mole of FAD per 83,500 g protein, but a higher value (1 FAD per 35,000 g protein) has been observed more recently (168).The absorption spectrum of this ETF, and ETF from other mammalian sources, differs considerably from those described above for P . elsdenii ETF, particularly in the region 400-500 nm. Highly purified ETF from pig liver and beef heart has absorption maxima a t 270, 375, 437.5, and 460 nm, and there appears to be very little absorption a t wavelengths greater than about 520 nm ( 1 ) . Electron-transferring flavoprotein from ra t (177,189) and monkey (166,167) liver on the other hand has a high peak of absorption in the region of 410 nm and very poor resolution of the absorption bands expected for FAD. These mammalian enzymes have not yet been analyzed for hydroxyflavins, and in particular for 6-OH-FAD which has an intense absorption band between 400 and 440 nm. The flavin of P. elsdenii ETF shows considerable fluorescence, a property which is shared by mammalian ETF (1,168) but by very few other flavoproteins (164.The fluorescence intensity is roughly twice that of free FAD, and it depends on the treatments described earlier which alter the absorption spectrum. The protein of this ETF is also fluorescent, and the positions of the fluorescence excitation and emission maxima suggest that there are contributions from both tyrosine and tryptophan ( 17 0 ) . 189. W.

R. Frisell, J. R. Cronin, and C. G. Mackenzie, JBC 237,2975 (1962).

116

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

3. Oxidation-Reduction Preparations of P. elsdenii ETF with FAD as the major chromophore are reduced by NADH, D-lactate and catalytic amounts of D-lactate dehydrogenase (175),and sodium dithionite (Fig, 12). They can also be reduced photochemically with EDTA (170). Enzyme reduced with NADH shows a band of long wavelength absorption which has been attributed to a complex between the reduced enzyme and NAD' (170). Titration of the enzyme with dithionite showed that complete reduction requires two electrons per molecule of flavin; hence, there are no colorless redox-active groups in the protein. A red intermediate which is generated during photochemical reduction appears to be the flavin semiquinone anion. Formation of the anion, which also occurs in mammalian ETF (I), is unexpected since ETF functions as a dehydrogenase, and flavoprotein dehydrogenases usually form the blue neutral semiquinone ( 6 3 ) . Preparations of ETF with a high content of the hydroxyflavins, and complexes of apo-ETF with purified 6-OH- and 8-OH-FAD, are similarly reduced by NADH, dithionite, or EDTA and light, but no stable semiquinone intermediates have been observed (178,179,181). The reduced enzyme is rapidly oxidized by air, anaerobic ferricyanide, or crotonyl-CoA and catalytic quantities of BCD (170,176). The rapid oxidation in air contrasts sharply with the slow rate of reaction observed with mammalian ETF (176).As noted earlier, the spectrum of the reoxidised enzyme differs under some circumstances from that of the starting material (Fig. 12). The spectral change is observed when enzyme which contains less than the full complement of flavin (1.4 moles of FAD per mole of protein) is reduced for times as long as those required for anaerobic titrations with reducing agents. The changes do not occur if the enzyme is fully reduced by a single addition of NADH or dithionite and immediately reoxidized, nor are they observed with enzyme that is saturated with flavin (which already has high absorption a t 400 nm). The spectral change is independent of the method used to reduce and oxidize the enzyme, and it is not accompanied by significant changes in catalytic activity (170). C. CATALYTIC PROPERTIES Electron-transferring flavoprotein from P . elsdenii mediates the oxidation of D-lactate dehydrogenase or NADH and the reduction of BCD. The intermediary role of ETF in these reactions has been established in coupled assays with catalytic amounts of the three enzymes and also by studying partial reactions with substrate levels of ETF or BCD as

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

117

the final electron acceptor (170,1?'2,17.5). Detailed kinetic studies with the purified enzyme and physical studies on its interactions with BCD and D-lactate dehydrogenase have not yet been carried out. D-Lactate dehydrogenase and NADH are the only known physiological electron donors for ETF, but a variety of compounds, including 2,6-di. The chlorophenolindophenol, will serve as electron acceptors (176,175) NADH dehydrogenase activity does not result from contamination by another enzyme as seems probable for ETF from pig (I) and rat (177) liver. Several protein fractionation techniques were used to analyze the purified protein (electrophoresis in acrylamide gel or Sephadex G-100, isoelectric focusing, gel filtration in Sephadex G-100,and analytical ultracentrifugation), but they neither separated the NADH dehydrogenase activity from ETF nor gave any indication for inhomogeneity other than that resulting from the hydroxyflavins (Section 1II1B,2). The modified flavins influence the activities of the enzyme since, as noted earlier, their complexes with apo-ETF display NADH dehydrogenase activity but they do not reduce BCD. Under defined assay conditions with 2,6dichlorophenolindophenol as electron acceptor, the relative NADH dehydrogenase activities of FAD-apo-ETF, 6-OH-FAD-apo-ETF1 and fI-OH-FAD-apo-ETF, are 1 :2.9: 0.31, respectively. The observed NADH dehydrogenase activity of enzyme preparations with the highest content of the hydroxyflavins agrees well with the theoretical value calculated from the activities of the separate complexes and the flavin composition of the enzyme (178).Although such preparations lack ETF activity, about half of their flavin content is unmodified FAD. It appears that modified flavin at one site in the enzyme influences the enzymic activity of FAD a t the other site since addition of 1 equivalent of 8-OH-FAD to apo-ETF that is 50% saturated with FAD causes a complete loss of ETF activity; NADH dehydrogenase activity in such mixed complexes is as expected for the binding of both flavins (178).This inhibitory effect of the hydroxyflavins is not understood. It is not yet clear whether the two molecules of FAD in ETF are equivalent in the catalytic reactions of the enzyme. When FAD is added to the isolated enzyme to saturate the flavin binding sites, the content of bound flavin increases by 40%, but the ETF activity is doubled and the NADH dehydrogenase activity with 2,6-dichlorophenolindophenolis unchanged (170).These observations, and the accompanying changes in the absorption spectrum (Section 111,B12), suggest that there may be differences between the two flavin binding sites. However, when increments of FAD are added to apo-FTF, the two activities increase in parallel until the end point at 2 moles of flavin per mole of protein; the changes in the absorption spectrum are also linear and provide no evidence for

118

STEPHEN G. MAYHEW AND MARTHA L. LUDWIG

differences in either the spectroscopic properties of the sites or their affinities for FAD (17'0). Beinert (1) has discussed the catalytic properties of mammalian ETF and experiments in which ETF from pig liver was shown to substitute for ETF prepared from other mammalian sources and mycobacteria. Similar exchange experiments with the protein from P. elsdenii have not yet been attempted.

Oxygenases: Dioxygenases OSAMU HAYAISHI MITSUHIRO NOZAKI MITCHEL T. ABBOTT ( I ) I . Introduction . . . . . . . . . . . . . . . . A . History and Definition . . . . . . . . . . . B . Classification . . . . . . . . . . . . . . C . Biologicai Function and General Properties of Dioxygenases I1. Heme-Containing Dioxygenases . . . . . . . . . . A . Tryptophan 2, 3-Dioxygenase . . . . . . . . . . B . Indoleamine 2, 3-Dioxygenase . . . . . . . . . I11. Nonheme Iron-Containing Dioxygenases . . . . . . . A . Phenol Dioxygenases . . . . . . . . . . . . B . Cysteamine Oxygenase . . . . . . . . . . . . C . Cysteine Oxygenaae . . . . . . . . . . . . D . Lipoxygenase . . . . . . . . . . . . . . E . Concluding Remarks . . . . . . . . . . . . IV . a-Ketoglutarate Dioxygenases . . . . . . . . . . A . Introduction . . . . . . . . . . . . . . . B. Prolyl Hydroxylase . . . . . . . . . . . . C . Lysyl Hydroxylaae . . . . . . . . . . . . . D . 7-Butyrobetaine Hydroxylase . . . . . . . . . E. Dioxygenase Reactions of Pyrimidines and Nucleosides . . F. p-Hydroxyphenylpyruvate Hydroxylase . . . . . . G . Mechanism . . . . . . . . . . . . . . .

.

.

120 120 . 121 . 123 . 127 . 127 . 130 . 132 . 133 . 148 . 149 . 150 . 150 . 151 . 151 . 152 . 165 . 167 . 169 . 179 . 183

.

1 Sections I and I1 prepared by Osamu Hayaishi. Section I11 prepared by Mitsuhiro Nozaki. and Section IV prepared by Mitchel T . Abbott .

119

120

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

1. Introduction

A. HISTORY AND DEFINITION In 1950, pyrocatechase was isolated from cells of a pseudomonad which catalyzed oxidative ring cleavage of the benzene ring of catechol forming cis,&-muconic acid as the reaction product [Eq. ( l ) ]( 2 ) . This enzyme

exhibited properties unlike those of an “oxidase” or “dehydrogenase,” because it was not associated with any of the previously known coenzymes or electron carriers, and none of the dyes or artificial electron acceptors tested could replace oxygen as an oxidant. According to the generally accepted belief, biological oxidation proceeded exclusively by the removal of electrons or hydrogen atoms from substrates, and direct addition of molecular oxygen was excluded from consideration. Oxygen might still be incorporated into substrates by hydration reactions involving water, but prior or subsequent dehydrogenation process would remove hydrogen or electrons as in the case of aldehyde oxidases. I n 1955, a heavy oxygen isotope was used as a tracer in l8OZand HZlsO and it was demonstrated that the two oxygen atoms incorporated into the product of the above reaction were derived exclusively from molecular oxygen rather than from water (3). Concurrently and independently, Mason and his collaborators, using the same isotope, found that during the oxidation of 3,4-dimethylphenol to 4,5-dimethylcatechol catalyzed by phenolase the oxygen atom incorporated into the substrate molecule was derived exclusively from molecular oxygen but not from the oxygen of water [Eq. (2) I ( 4 ) .

6

CHS

CHS

+ $0,-

Ho9 CH,

CHS

(2)

These findings conflicted sharply with the current concept that oxygen could act only as an ultimate electron acceptor in biological oxidation and that all oxygen atoms incorporated into substrates are derived from 2. 0. Hayaishi and Z. Hashimoto, J. Biochem. (Tokgo) 37, 371 (1950). 3. 0. Hayaishi, M. Katagiri, and 8. Rothberg, JACS 77,5450 (1955). 4. H. S. Mason, W. L. Fowlks, and E. Peterson, JACS 77, 2914 (1955).

3.

OXYGENASES

121

DIOXYGENASES

the oxygen atoms of water. These two newly discovered reactions may be schematically represented by Eqs. (3) and (4) :

x + oz+ xoz x + 4 0 2 - f xo

(3) (4) Because these two new types of reactions both involve “oxygen fixation” into a substrate molecule, they are different from the classic oxidase reactions, but are similar to the oxygenation reactions known to occur in chemical or photochemical processes. We therefore proposed a new term “oxygenase” to designate enzymes which catalyze such oxygen fixation reactions ( b ) , and the terms “di” and “mono” oxygenases are generally assigned, respectively, to the enzymes catalyzing these two types of reactions ( 6 ) .

B. CLASSIFICATION Dioxygenases are defined as enzymes catalyzing reactions in which both atoms of molecular oxygen are incorporated into substrates. I n many instances where one substrate can act as the oxygen acceptor [Eq. ( 3 ) ] , the term “intramolecular dioxygenases” may be used. The dioxygenases acting upon two acceptor substrates, which have recently been reported from a number of laboratories, may be referred to as “intermolecular dioxygenases.” One of the two substrates for the latter type has so far been found invariably to be a-ketoglutarate, and the overall reaction may be schematically shown by Eq. ( 5 ) . More detailed discussions of this type of enzyme will be found in Section IV. a-Ketoglutarate

+ O2 + X

+ succinate

+ C o t + XO

(5) A third class of dioxygenases includes various types of enzymes which are not yet well characterized but play important roles in the metabolism of sulfur-containing compounds, prostaglandins, fatty acids, vitamin A, and other compounds. These enzymes are tentatively classified as “Miscellaneous” in Table I but will be described below in more detail. The other subclass of oxygenases is “Monooxygenases,” which are defined as a group of enzymes catalyzing the incorporation of one atom of molecular oxygen into a substrate while the other oxygen is reduced to water. The simplest type of monooxygenase catalyzes the incorporation of a single atom of molecular oxygen concomitant with the reduction of the other oxygen atom by electrons derived from the substrate. 5. 0. Hayaishi, S. Rothberg, and A . H. Mehler, Abstr. 130th Meet. Amer. Chern. SOC.p. 53C (1956). 6. 0 . Hayaishi, Proe. Plen. Sess., Znt. Congr. Biochem., 6th, 1964 p. 31 (1964).

122

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

TABLE I CLASSIFICATION OF OXYGENASES EC NO.^

Oxygenase

A. Dioxygenase 1. Intramolecular dioxygenase a. Hemoprotein b. Nonheme iron protein c. Copper protein d. Flavoprotein 2. Intermolecular dioxygenase 3. Miscellaneous

1.13.11 1.13.11 1.13.11 1.14.12 1.14.11 1.13.99

B. Monooxygenase

1. Internal monooxygenase 2. External monooxygenase a. Pyridine nucleotide-linked flavoprotein b. Flavin-linked hemoprotein c. Iron-sulfur protein-linked hemoprotein d. Pteridine-linked monooxygenase e. Ascorbate-linked copper protein f . With another substrate as reductant

1.13.12 1.14.13 1.14.14 1.14.15 1.14.16 1.14.17 1.14.18

Enzyme Commission number (EC No.) refers to the new numbering system introduced in 1972 by the International Union of Biochemistry, Enzyme Nomenclature Commission. 0

Thus, the overall reaction may be expressed by the following equation:

+

XHz + 02+ XO Hz0 (6) Since the reducing agent is internally supplied, these enzymes may be referred to as internal monooxygenases. While the internal monooxygenases do not require external reducing agents, the more common types of monooxygenases require various kinds of electron donors. The overall reactions are schematically represented by Eq. ( 7 ) . One of the atoms of molecular oxygen is incorporated into a substrate molecule and the other is reduced to H,O in the presence of an appropriate electron donor, (DH2),such as NADH, NADPH, tetrahydrobiopterin, or ascorbic acid [Eq. (7) 1.

X

+

0 2

+ D H a + XO

+ HzO + D

(7)

Because information concerning the mechanism of action of oxygenases is still limited even though the field is progressing rapidly, any classification scheme would necessarily be arbitrary and perhaps temporary. Two subclasses, di- and monooxygenases, have been employed by the majority of workers in this field, although the distinction between the di- and

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OXYGENASES : DIOXYGENASES

123

monooxygenase subclasses, discussed above, is phenomenological and historical and may not be so permanent and meaningful.

C. BIOLOGICAL FUNCTION AND GENERAL PROPERTIES OF DIOXYGENASES The significance of biological oxygen fixation in medicine, agriculture, microbiology, and also in food technology, cosmobiology, public health problems, and biochemistry in general has now been well established. Dioxygenases play important roles in biosynthesis, transformation, and degradation of essential metabolites such as amino acids, lipids, sugars, nucleic acids, porphyrins, vitamins, and hormones. They also play a crucial role in the degradation of various aromatic compounds such as drugs, insecticides, and carcinogens. Furthermore, they participate in the degradation of v'arious natural and synthetic compounds by soil and airborne microorganisms in nature and are therefore of great significance in environmental science. 1. Dioxygenases Involved in the Cleavage of a C=C Bond The major reaction catalyzed by dioxygenases is the cleavage of an aromatic double bond, which may be located (a) between two hydroxylated carbon atoms, (b) adjacent to a hydroxylated carbon atom, or (c) in an indole ring. However, a similar type of reaction also occurs with aliphatic substrates such as p-carotene, which yields vitamin A as the product. A large number of enzymes which catalyze the ring cleavage of various phenols have been isolated and characterized from animals, plants, and microorganisms. Some of them have been purified extensively and crystallized. Most of these enzymes contain either ferrous or ferric inorganic iron as a sole cofactor. A detailed account of their molecular and catalytic properties is given in Section 111. Indole derivatives including tryptophan, serotonin, and melatonin are degraded by enzymes containing protoheme I X as a sole prosthetic (see Section 11). 2-Methyl-3-hydroxypyridine-5-carboxylatedioxygenase was isolated from a pseudomonad and was crystallized by Sparrow and his co-workers (7 ) . It contains approximately 2 moles of FAD per mole of enzyme protein and catalyzes the following reaction: __

7. L. G. Sparrow, P. P. K. Ho, T. K. Sundaram, D. Zach, E. J. Nyns, and E. E. Snell, JBC 244, 2590 (1969).

TABLE I1 ENZYMES PRODUCING CATECHOL OR Im DERIVATIVES Enzyme

Reaction

Cofactor

Ref.

Anthranilate hydroxylase (decarboxylating)

Fez+, NADH

8

Anthranilak hydroxylase

Fez+, NADPH

9

Benzoate oxygenase

-aoH 0 .-aon +

+ co,

0,

Fe*+, NADH

10

Fe*+, NADH

11

on

Benzene oxygenase

+

OH

3.

125

OXYGENASES: DIOXYGENASES

This is probably the only dioxygenase which has been characterized as a flavoprotein. 2. Dioxygenuses Involved in the So-called Double H ydroxy lation Reactions Several enzymes are known which produce catechol or its derivatives from either anthranilate, benzoate, or benzene. These are listed in Table I1 (8-11). 3. Dioxygenases Involved in the Metabolism of Sulfur and Sulfur-Containing Compounds

Thiobacillus thiooxiidans contains an oxygenase which produces H2Sz03 from inorganic sulfur and molecular oxygen in the presence of reduced glutathione [ Eq. (9) ] (12J3). 2S

+ + H?O -+ 0 2

HZSZOS

(9)

Cysteamine (14-16) and cysteine (17-19) are also metabolized by individual dioxygenases to form hypotaurine and cysteine sulfinic acid, respectively. The details concerning these enzymes will be described in Section 111.

4. Miscellaneous Dioxygenases a. Quereetine Dioxygenase. This unique enzyme has been isolated and partially purified from Aspergillus flavus and was shown to contain copper as its sole prosthetic group (20,21). 8. S. Kobayashi, S. Kuno, N. Itada, and 0. Hayaishi, BBRC 16,556 (1964). 9. R. P. Kumar, S. D. Ravindranath, C. S. Vaidyanathan, and N. A. Rao, BBRC 49, 1422 (1972). 10. H. Takeda, M. Mori, N. Itada, H. Taniuchi, Y. Kojima, and 0. Hayaishi, unpublished. 11. D. T. Gibson, J. R. Koch, and R. E. Kallio, Biochemist-ry 7, 2653 (1968). 12. I. Suzuki, BBA 104, 359 (1965). 13. I. Suzuki, BBA 110, 97 (1965). 14. D. Cavallini, R. Scandurra, and C. De Marco, JBC 238, 2999 (1963). 15. D. Cavallini, R. Scandurra, and F. Monacelli, BBRC 24, 185 (1966). 16. G. Rotilio, G. Federici, L. Calabrese, M. Costa, and D. Cavallini, JBC 245, 6235 (1970). 17. B. Sorbo and L. Ewetz, BBRC 18, 359 (1965). 18. J. B. Lombardini, P. Turini, D. R. Biggo, and T. P. Singer, Physiol. Chem. Phys. I (1969). 19. J. B. Lombardini, T. P. Singer, and P. D. Boyer, JBC 244, 1172 (1969). 20. H. G. Krishnamurty and F. J. Simpson, JBC 245, 1467 (1970). 21. T.Oka and F. J. Simpson, BBRC 43, 1 (1971).

126

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

/

OH

HO

0

b. p-Carotene 15,15’-Dwxygenase. This enzyme has been isolated and partially purified from rat liver and intestine. p-Carotene is cleaved by the insertion of two atoms of molecular oxygen to produce 2 moles of retinal (92,dS) as follows:

c. Prostaglandin Synthetase. Prostaglandins are synthesized from unsaturated fatty acid precursors by the action of an enzyme complex containing a t least two oxygenases and an isomerase (24). The overall reaction is as follows: 22. J. A. Olson and 0. Hayaishi, Proc. Nat. Acad. Sci. U. S. 54, 1364 (1965). 23. D. S. Goodman, H. S. Huang, M. Kanai, and T. Shiratori, JBC 242, 3543 (1907). 24. M. Hamberg, B. Samuelsson, I. Bjorkhem, and H. Danielson, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 29. Academic Prem, New York, 1974.

3.

127

OXYGENASES : DIOXYGENASES

0

Prostaglandin E, -I-

(12)

Prostaglandin F1,

The oxygenase fraction and the isomerase fraction have been isolated from microsomal particles of bovine vesicular gland and the endoperoxide intermediate has been characterized ( 6 5 ) . This oxygenase is inhibited by anti-inflammatory agents such as indomesathine and aspirin. d. Lipoxygenase. Lipoxygenase is prevalent in plants and catalyzes the hydroperoxidation of polyunsaturated fatty acids and esters containing a cis,ci-s-1,4-pentadiene system. As discussed in Section 111, it now appears that lipoxygenase is no longer an exception t o the generalization that all of the dioxygenases obtained in crystalline form contain either iron or copper as the sole cofactor. The only exception to this rule is a flavoprotein, 2-methyl-3-hydroxypyridine-5-carboxylatedioxygenase, mentioned above ( 7 ) .

II. Heme-Containing Dioxygenases

A. TRYPTOPHAN 2,3-DIOXYGENASE 1.

Introduction

I n 1936, Kotake and Masayama described the conversion of L-tryptophan t o L-kynurenine in vitro by crude extracts of rabbit liver (66).Since the reaction involved the cleavage of the pyrrole ring of tryptophan, the 25. T. Miyamoto, S. Yamamoto, and 0. Hayaishi, Proc. N u t . Acad. Sci. U. S. (in press). 26. Y. Kotake and T. Masayama, Hoppe-Seyler’s 2.Physiol. Chem. 243,237 (1936).

128

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

enzyme was named “tryptophan pyrrolase.” About 15 years later, Knox and Mehler purified the enzyme system and separated the enzyme catalyzing the conversion of tryptophan to formylkynurenine and a second enzyme catalyzing the hydrolysis of the latter to kynurenine and formic acid (27). I n 1957, experiments carried out with 1 8 0 2 and H,180 showed that molecular oxygen, but not the oxygen atom of water, was incorporated into the reaction products (28). Thus, the enzyme was renamed “tryptophan 2,3-dioxygenase” [ Eq. (13)1.

Thus far tryptophan 2,3-dioxygenase has been found only in the livers of mammals and in some microorganisms. This enzyme catalyzes the first, and probably the rate-limiting, step in the metabolic conversion of tryptophan to pyridine nucleotides ( g 9 ) . The enzyme is induced by L-kynurenine in Pseudomonas (30) and by glucocorticoids in the livers of animals ($1). Tryptophan also elevates the enzyme level in liver, but the effect of tryptophan seems to result from the diminished rate of breakdown of enzyme protein rather than from an increased rate of synthesis (32-34). 2. Molecular Properties

a. Purification. Either cells of Pseudomonas, grown in the presence of tryptophan (36),or rat liver of animals previously injected with tryptophan and glucocorticoid hormones serve as a starting material for purification. These enzymes appear to be in the soluble fraction and can be purified by conventional procedures (3637). The enzyme has not yet been obtained in a crystalline form, but final preparations of the enzyme appear to be homogeneous as judged by acrylamide gel electrophoresis. b. Prosthetic Group. Both microbial and hepatic enzymes contain protoporphyrin IX. Highly purified hepatic enzyme is catalytically inert and 27. W. E. Knox and A. H. Mehler, JBC 187,419 (1950). 28. 0. Hayaishi, S. Rothberg, A. H. Mehler, and Y. Saito, JBC 229, 889 (1957). 29. Y. Nishizuka and 0. Hayaishi, JBC 238, 3369 (1963). 30. N. J. Palleroni and R. Y. Stanier, J . Gen. Microbiol. 35, 319 (1964). 31. W. E. Knox, J . Exp. PathoE. 32, 462 (1951). 32. R. T. Schimke, E. W. Sweeney, and C. M. Berlin, BBRC 15, 214 (1964). 33. R. T. Schimke, E. W. Sweeney, and C . M. Berlin, JBC 240, 322 (1965). 34. R. T. Schimke, E. W. Sweeney, and C. M. Berlin, JBC 240, 4609 (1965). 35. 0. Hayaishi and R. Y. Stanier, J. Bacteriol. 62, 691 (1951). 36. W. N. Poillon, H. Maeno, K. Koike, and P. Feigelson, JBC 244, 3447 (1969). 37. G. Schutz and P. Feigelson, JBC 247, 5327 (1972).

3.

129

OXYGENASES : DIOXYGENASES

TABLE I11 OF PSEUDOMONAD A N D HEPATIC ENZYMES MOLECULAR PROPERTIES ~

~~

~

Property ~~

Molecular weight Subunit Subunit molecular weight Subunit structure Specific activity (pmoles of product formed per minute per mg protein at 20") a

Pseudomonada

Rat live+

122,000 4 31,000 ? 17.0

167,000 4 43,000 a2 8 2 2.5

~

From Poillon et al. (36).

* From Schutz and Feigelson (37). full catalytic activity can be restored upon addition of exogenous ferriprotoporphyrin IX (38,39). The presence of 2 g-atoms of copper and 2 moles of heme per mole of enzyme has been reported by Feigelson and co-workers (40,dl). On the other hand, Poillon et al. (36) and Ishimura and Hayaishi (42) found only minute quantities of copper in their preparations and regarded these as adventitious. Since the specific activities of the enzyme preparations used by these investigators are in the same order of magnitude, the possible presence of copper in L-tryptophan 2,3-dioxygenase deserves further investigation. c. Homogeneous Preparations. Some of the molecular properties reported with homogeneous preparations of pseudomonad and hepatic enzymes are listed in Table 111. 3. Catalytic Properties

a. Reductive Activation of the Enzyme. The purified enzyme exhibits absorption spectra in the visible and the ultraviolet regions typical of ferriheme proteins. This form of enzyme is catalytically inactive unless the heme is reduced by some reducing agent such as H,O, (43),ascorbic acid (43), or superoxide anion (42,44).Feigelson and co-workers asserted that copper plays an essential role in the oxidation-reduction of the 38. 39. 40. 41.

P. Feigelson and 0. Greengard, JBC 236, 153 (1961). P. Feigelson and 0. Greengard, BBA 50,200 (1961). H. Maeno and P. Feigclson, BBRC 21, 297 (1965). F. 0. Brady, M. E. Monaco, H. J. Fornian, G . Schuts, and P. Feigelson, JBC

247, 7915 (1972). 42. Y. Ishimura and 0. Hagaishi, JBC 248, S610 (1973). 43. T. Tanaka and W. E. Knox, JBC 234, 1162 (1959). 44. 0. Hayaishi and M. Nozaki, Science 164, 389 (1969).

130

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

heme, since the copper complexing agents selectively inhibited tryptophan dioxygenase activity (45). I n contrast, Ishimura and Hayaishi (42) found that the inhibitory effect of copper chelating agents such as sodium diethyldithiocarbamate and bathocuproine sulfonate on enzymic activity was the result of their specific chelating action but was attributable to their unforeseen properties as a hydrogen peroxide-trapping reagent and as a nonspecific inhibitor, respectively. The role of copper in this enzyme still remains to be elucidated. b. Kinetic Properties. A large body of evidence reported from the Kyoto group ( 4 )and the Columbia group (45) is consistent with an ordered bi-uni mechanism in which the substrate, L-tryptophan, binds with the active (reduced) form of enzyme and then with molecular oxygen. When oxygen binds with the enzyme-substrate complex, there appears a spectral species possessing maxima a t 418, 545, and 580 nm, similar to those of the oxygenated ferroheme forms of hemoglobin, myoglobin, and peroxidase. Its spectral properties and conditions for its appearance and decay are all compatible with the interpretation that this spectral species results from a ternary complex of enzyme-substrate-oxygen and that the oxygenated form is indeed the true intermediate of the catalytic process (46-48).Since the oxygenated form of enzyme was discovered in 1967 with tryptophan 2,3-dioxygenase1 similar findings have been made with other dioxygenases such as protocatechuate 3,4-dioxygenase (49,50) and certain monooxygenases such as cytochrome P-450 (51) and others.

B. INDOLEAMINE 2,3-DIOXYGENASE 1. Introduction The presence of an enzyme system that catalyzes the conversion of D-tryptophan to D-kynurenine via D-formylkynurenine was first reported in 1963 from the intestine of rabbit (52).Subsequently, the enzyme was 45. H. J. Forman and P. Feigelson, Biochemistry 10, 760 (1971). 46. Y. Ishimura, M. Noaaki, 0. Hayaishi, M. Tamura, and I. Yamaaaki, JBC 242, 2574 (1967). 47. 0 . Hayaishi, Ann. N . Y . Acad. Sci. 158, 318 (1969). 48. Y. Ishimura, M. Nozaki, 0. Hayaishi, T. Nakamura, M. Tamura, and I. Yamaaaki, JBC 245, 3593 (1970). 49. H. Fujisawa, K. Hiromi, M. Uyeda, M. Nozaki, and 0. Hayaishi, JBC 246, 2320 (1971).

50. H. Fujisawa, K. Hiromi, M. Uyeda, S. Okuno, M. Nozaki, and 0. Hayaishi, JBC 247, 4422 (1972). 51. J. A. Peterson, Y. Ishimura, and B. W. Griffin, A B B 149, 197 (1972). 52. K. Higuchi, S. Kuno, and 0. Hayaishi, Fed. Proc., Fed. Arner. SOC.Exp. Biol. 22, 243 (1963).

3.

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131

partially purified from the ileum of the rabbit intestine and was named %-tryptophan pyrrolase” by Higuchi and Hayaishi (53) since the classic tryptophan pyrrolase acts specifically on the L isomer of tryptophan. Furthermore, it was noted that catalase inhibited the L-tryptophan pyrrolase activity, whereas it stimulated the activity of the new enzyme. These authors also suggested the possible involvement of superoxide anion in this reaction because adenosine and xanthine oxidase stimulated enzymic activity. I n 1967, the enzyme was purified about 100-fold ( 5 4 ) . Methylene blue and ascorbate were shown to be required for maximal activity. Xanthine oxidase and its substrate could replace ascorbate but hydrogen peroxide, added directly or enzymically generated, was not effective. Again catalase was shown t o be completely ineffective. Further studies by Hirata and Hayaishi (55) revealed that highly purified preparations of superoxide dismutase obtained from bovine erythrocytes and green peas inhibited the catalytic activity of intestinal tryptophan 2,3-dioxygenase a t any stage of the reaction. I n contrast, superoxide dismutase failed to inhibit hepatic and pseudomonad tryptophan 2,3-dioxygenases during the steady state of the reaction. With hepatic and pseudomonad enzymes, however, a prolonged lag period was observed when superoxide dismutase was added to the reaction mixture prior to the start of the reaction. Further experiments with superoxide anion, produced by electrolytic reduction of molecular oxygen, indicated that superoxide anion, rather than molecular oxygen, serves as the oxygenating agent for the intestinal enzyme while i t reductively activates pseudomonad and hepatic enzymes ( 6 5 ) . 2. Molecular and Catalytic Properties The enzyme has now been purified about 650-fold from rabbit intestine to near homogeneity but has not yet been obtained in a crystalline form. Molecular weight was estimated to be approximately 120,000 and it contains 2 moles of heme per mole of enzyme protein. The enzyme has been found in the intestine, stomach, lung, and brain but not in liver or kidney of rabbit ( 5 6 ) . A highly purified preparation of intestinal enzyme can degrade both D and L isomers of tryptophan to form corresponding isomers of formylkynurenine. Furthermore, it also catalyzes oxygenative ring cleavage of L53. K. Higuchi and O. Hayaishi, ABB 120, 397 (1967). 54. S. Yamamoto and 0. Hayaishi, JBC 242, 5260 (1967). 55. F. Hirata and 0. Hayaishi, JBC 246, 7825 (1971). 56. 0. Hayaishi, in “Proceedings of the Robert A. Welch Foundation Conferences on Chemical Research” (W. 0. Milligan, ed.), p. 185. Robert A. Welch Found., Houston, Texas, 1971.

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OSAMU HAYAISHI, MITSUHIRO NOZAXI, AND MITCHEL T. ABBOTT

TABLE IV KINETICPARAMETERS OF SUBSTRATES FOR INDOLEAMINE 2,3-DIOXYGENASE V ma*

K,

Substrate

(nmole/mg of protein/min at 24')

(pM)

D-Tryptophan L-Tryptophan D-5-Hydroxytryptophan b5-Hydroxytryptophan Tryptamine Serotonin Melatonin

140 140 9 11 5 3 0.1

300 20 100 6 250 150 40

and D-5-hydroxytryptophan, serotonin, tryptamine, and melatonin (67). Kinetic parameters for these substrates are presented in Table IV. Skatole, indole, indoleacetic acid, and 5-hydroxyindoleacetic acid do not serve as substrate. Although melatonin is a rather poor substrate for this enzyme in vitm, recent evidence from our laboratory demonstrated that a partially purified enzyme from rabbit brain catalyzed the oxygenative cleavage of the pyrrole moiety of melatonin yielding N~-acetyl-NZ-formyl-5-methoxykynurenamine, which was further degraded to Nu-acetyl-5-methoxykynurenamine by the action of formamidase. Authentic samples of these two metabolites were synthesized by ozonolysis of melatonin and shown to be indistinguishable from the enzymic products. When [ '*C] melatonin was injected intracisternally, the major metabolite in the rat brain was shown to be N7-acetyl-5-methoxykynurenamine.The results indicate that these two new metabolites are involved in the major metabolic pathway of melatonin in the central nervous system (68). In view of the broad substrate specificity of this enzyme, it is proposed that this enzyme be designated indoleamine 2,3-dioxygenase (pyrrolase),

111. Nonheme Iron-Containing Dioxygenarer

Among some 35 dioxygenases studied to date, more than 8Q% have nonheme iron built into their structure or require added iron for full activity. These dioxygenases have been discovered in all types of living 57. F. Hirata and 0. Hayaishi, BBRC 47, 1112 (1972). 58. F. Hirata, 0. Hayaishi, T. Tokuyama, and S. Senoh, JBC 249, 1311 (1974).

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133

organisms and have been shown to perform a variety of functions (44,69). The cleavage of the aromatic ring, the oxidation of certain sulfur-containing compounds, and the hydroperoxidation of polyunsaturated fatty acids appear to depend largely or entirely upon this type of enzyme. Thus, extensive studies on the reaction mechanism of dioxygenases have been carried out, especially with phenolic dioxygenases that cleave the phenol or catechol ring with the insertion of two atoms of molecular oxygen. Details of these studies have been reviewed in two recent articles (60,61). A. PHENOLIC DIOXYGENASES Catechol dioxygenases can be classified into two groups from the mode of ring fission. These are intradiol and extradiol dioxygenases. The former cleaves the C-C bond of the catechol ring between two hydroxylated carbon atoms and the latter between a hydroxylated and an adjacent carbon atom (62). As will be discussed below, differences between the intradiol and extradiol dioxygenases include not only their functions but also the valence state of the iron bound to the enzyme. 1. Intradiol Dioxygenases

a. Pyrocatechase Molecular and catalytic properties-Pyrocatechase [ catechol: oxygen 1,2-oxidoreductase (decyclizing), EC 1.13.11.11 is the first enzyme shown by l80studies to be a dioxygenase (3). This enzyme catalyzes the cleavage of the aromatic ring of catechol to form cis,cis-muconic acid with the insertion of two atoms of molecular oxygen. It is a typical example of an intradiol dioxygenase [Eq. (14) 1. The enzyme has been highly puri-

fied from cell-free extracts of Pseudomonas arvilla C-1 (ATCC 23974) grown with benzoate as the sole carbon source (63).The purified preparation has a specific activity of about 30 pmoles/min/mg of protein and is homogeneous as judged by ultracentrifugal and electrophoretic criteria. The enzyme contains 2 g-atoms of iron per mole of enzyme, based on 59. 0. Hayaishi, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 1. Academic Press, New York. 1974.

60. M. Nozaki, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 135. Academic Press, New York, 1974. 61. M. Nozaki and Y. Ishimura, in “Microbial Iron Metabolism” (J. B. Neilands, ed.), p. 417. Academic Press, New York, 1974. 62. M. Nozaki, S. Kotani, K. Ono, and S. Senoh, BBA 220, 213 (1970).

134

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

TABLE V SUESTRATE SPECIFICITY OF

P Y R o c A T E c H A S E s FROM

Pseudomonas

AND

Brevibacterium

Relative activity (%) of pyrocatechase from Substrate

Pseudomonas

Brevibacterium

Catechol 4-Methylcatechol 3-Methylcatechol PChlorocatechol Pyrogallol 4Hydroxycatechol 4-Methoxycatechol

100 95 5 2 0 -

100 95 140-150 85 30-50 85

a molecular weight of 90,000 (63). The apoenzyme is prepared by the addition of o-phenanthroline to the enzyme solution in the presence of sodium dithionite, followed by exhaustive dialysis. The apoprotein thus obtained can be partially reactivated under aerobic conditions by incubating with ferrous ion in the presence of ascorbic acid ( 6 4 ) . Another pyrocatechase which has been studied is one obtained from phenol-induced Brevibacterium fuscum P-13.This enzyme has a molecular weight of 64,000and contains 1 g-atom of iron per mole of enzyme ( 6 5 ) . The iron is easily removed by incubating the enzyme with a nonmetabolizable substrate analog such as ethylprotocatechuic acid. The apoenzyme which retains less than 5% of its original activity can be fully reactivated by treating it with stoichiometric amounts of ferrous ion in the presence of oxygen (66). The substrate specificities of the two pyrocatechases differ somewhat from each other (63,67-69) (Table V). The following compounds do not serve as substrate for Pseudomonas pyrocatechase: adrenolutine, p-aminocatechol, protocatechuic acid, 2,3- and 2,4-dihydroxybenzoic acid, trans-5,6-dihydroxycyclohexadine, pyrogallol, dopa, homogentisic acid, gentisic acid, gallic acid, 2,3-dihydroxyphenylacetic acid, and Tiron (63,67). K, values for catechol and oxygen are 4 and 20 p M , respectively. 63. Y. Kojima, H. Fujisawa, A. Nakazawa, T. Nakazawa, F. Kanetsuna, H. Taniuchi, M. Nozaki, and 0. Hayaishi, JBC 242, 3270 (1967). 64. T. Nakazawa, M. Noaaki, 0. Hayaishi, and T. Yamano, JBC 244, 119 (1969). 65. K. Nagami and Y. Miyake, BBRC 42, 497 (1971). 66. K. Nagami, BBRC 47, 803 (1972). 67. 0. Hayaishi, M. Katagiri, and S. Rothberg, JBC 229, 905 (1957). 68. H. Nakagawa, H. Inoue, and Y . Takeda, J . Biochem. ( T o k y o ) 54,65 (1963). 69. K. Nagami, Doctral thesis, Faculty of Science, Osaka University (1973).

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135

For Brevibacterium pyrocatechase, the catechol analogs which have an electron-donating residue serve as substrates, whereas those having an electron-withdrawing residue do not serve as substrates. The latter compounds include ethylprotocatechuate, 3,4-dihydroxyacetophenone, 3,4-dihydroxybenzaldehyde1 4-nitrocatechol protocatechuic acid, and 3,4-dihydroxybenzene sulfonic acid. When incubated with the enzyme, these compounds remove iron from the protein, which appears to result from their strong reducing ability (69). As previously mentioned, pyrocatechase is believed to cleave the catechol ring exclusively in the intradiol manner. However, recent studies in our laboratory have revealed that the enzyme from Pseudomonas catalyzes not only an intradiol cleavage but also an extradiol cleavage when 3-methylcatechol is used as a substrate. I n contrast, Brevibacteriuw pyrocatechase cleaves this substrate only in the intradiol manner ( 7 0 ) . Spectral properties-A concentrated solution of highly purified pyrocatechase has a pronounced red color with a broad absorption band between 390 and 650 nm (63,71).The apoenzyme shows neither significant absorption in the visible region nor enzymic activity. Upon reconstitution, the visible absorption and enzyme activity are restored. The visible absorption also decreases on adding sodium dithionite and reappears when the solution is exposed to air. These results suggest that trivalent iron bound to enzyme is responsible for the red color and essential for enzymic activity (63,7l). Under anaerobic conditions, the addition of catechol causes the color of the enzyme solution to change from red to grayish blue with a concurrent increase in absorbance a t about 710 nm, indicating the possible formation of an enzyme-substrate complex. Upon the introduction of oxygen and the consequent degradation of catechol to &,&-muconic acid, the original absorption spectrum is again restored. However, the absorption spectrum of the native enzyme is not altered by the addition of oxygen, nor is it changed by exhaustive evacuation (63,71). When the enzyme obtained from Pseudmonas is titrated with catechol under anerobic conditions, 2 moles of substrate per mole of enzyme are required to bring about a maximal absorption change, demonstrating that 2 moles of substrate can combine with the enzyme (7.2). Electron spin resonance properties-Pseudomonas pyrocatechase shows a sharp electron spin resonance (ESR) signal a t g = 4.28 characteristic 70. M . Fujiwara, L. Golovleva, Y. Saeki, M. Nozaki, and 0. Hayaishi, JBC (1975) (in press). 71. K . Nagami and Y. Miyake, BBRC 48, 198 (1972). 72. M. Nozaki, T. Nakazawa, H. Fujisawa, S.Kotani, Y . Kojima, and 0. Hayaishi, Advan. Chem. Ser. 77,242 (1968).

136

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

of the presence of ferric ion. This signal decreases markedly upon the addition of sodium dithionite and reappears when the solution is exposed to air (64,73). When catechol is added to the enzyme under anerobic conditions, the signal disappears instantaneously and returns to the original level after all of the substrate has been degraded to product on the addition of air (64,73). The colorless, inactive apoenayme shows essentially no signal in the vicinity of g = 4.2. Upon incubation of the apoenzyme with ferrous ion in the presence of oxygen, the signal and the red color are partially restored. A similar signal change caused by the addition of catechol has also been reported with Brevibacterium pyrocatechase (74). However, when protocatechuic acid is used as a substrate, the ESR signal increases rapidly and then decreases very slowly. In contrast, ethylprotocatechuate causes a rapid decrease in the intensity of the ESR signal with concomitant removal of iron from the protein ( 6 5 ) . Circular dichroism-The circular dichroism (CD) of Pseudomonas pyrocatechase exhibits several strong positive bands between 250 and 300 nm, a moderate negative band a t 327 nm, and a weak but broad negative band around 500 nm. I n addition, there is a strong negative band a t 222 nm, characteristic of proteins containing an a-helical structure. When the bound iron is removed, the bands a t 327 and 500 nm completely disappear and those between 250 and 300 nm are partially diminished. Upon incubation with ferrous ion under aerobic conditions these bands are restored to almost the original level ( 7 6 ) .Similar, but not identical, CD bands have been reported for Brevibacterium pyrocatechase (71). Since the bands above 300 nm also disappear on addition of reducing agents, there is a possibility that the ferric ion bound to the enzyme may be responsible for the bands. In the presence of catechol under anaerobic conditions, changes in the magnitude and position of the CD bands occur in the visible range, indicating that an alteration in the state of the ligands around the iron may be involved following the binding of substrate (75). b. Protocatechuate 3,4-Dioxygenase Molecular and catalytic properties-Protocatechuate 3,4-dioxygenase [protocatechuate :oxygen 3,4-oxidoreductase (decycliaing) , EC 1.13.11.31 catalyzes the conversion of protocatechuic acid to p-carboxymuconic acid with the incorporation of two atoms of molecular oxygen [Eq. (15)1. The 73. T. Nakazawa, Y. Kojima, H. Fujisawa, M. Nozaki, 0. Hayaishi, and T. Yamano, JBC 240, PC322-2 (1965). 74. H. Kita, Y. Miyake, M. Kamimoto, S. Senoh, and T. Yamano, J . Biochem. (Tokyo) 66, 45 (1969). 75. A. Nakaaawa, T. Nakazawa, S. Kotani, M. Nozaki, and 0. Hayaishi, JBC 244, 1527 (1969).

3.

137

OXYGENASES : DIOXYGENASES

HoJ - J HOOC

+

OH

4

-

HOOC

/

COOH

\

COOH

(15)

enzyme has been obtained in crystalline form from p-hydroxybenzoateinduced cells of Pseudomonas aeruginosa (ATCC 23975) ( 7 6 ) .The molecular weight is approximately 700,000 and the molecular activity is calculated to be 45,500 min-* a t 24O. The enzyme contains about 8 g-atoms of iron per mole, eight substrate binding sites, and appears to consist of eight subunits (77). The iron is tightly bound, but recently its removal from the enzyme has been achieved by prolonged anaerobic dialysis against a buffer containing both o-phenanthroline and NazS,O,. The apoenzyme thus obtained can be fully reactivated by incubation with Fe(I1) and NazS,Oa (78). Among catechol derivatives, the following are cleaved by oxygen in the presence of enzyme, with relative rates indicated in parentheses: protocatechuic acid (loo), pyrogallol (2.4), catechol (0.4) , 3-methylcatechol (0.4), 4-methylcatechol (0.2), 3,4-dihydroxyphenylacetic acid (0.2), 3,4dihydroxymandelic acid (0.1), and 3,4-dihydroxyphenylpropionic acid (0.03). The following compounds are either not oxygenated or oxygenated at rates less than 0.01% of that of protocatechuic acid: protocatechualdehyde, 2,3-dihydroxybenzoic acid, 3,4-dihydroxyacetophenone,3,4-dihydroxyphenylalanine, vanillic acid, and 4,nitrocatechol ( 7 6 ) . Initial velocity plots of the enzyme with one substrate as the fixed substrate and the other as the variable substrate give lines with intersecting patterns, indicating the formation of a ternary complex of enzyme, oxygen, and catechol. A dead-end inhibitor for the enzyme, protocatechualdehyde, inhibits the enzyme competitively with respect to protocatechuic acid and noncompetitively with respect to oxygen. These results suggest an ordered bi-uni mechanism in which an organic substrate first combines with the enzyme, which then reacts with oxygen to form a ternary complex ( 7 9 ) . As will be discussed below, catalytic properties as well as the appearance of the enzyme are very similar to those of pyrocatechase. Spectral properties-Like pyrocatechase, the enzyme has a red color which results from binding by the ferric ion and shows a broad absorption peak between 400 and 650 nm. The red color disappears upon the addition of sodium dithionite and reappears when the solution is exposed to air 76. H. Fujisawa and 0. Hayaishi, JBC 243, 2673 (1968). 77. H. Fujisawa, M. Uyeda, Y. Kojima, M. Noeaki, and 0. Hayaishi, JBC 247, 4414 (1972). 78. M.Fujiwara and M. Nozaki, BBA 327, 306 (1973). 79. K. Hori, T.Hashimoto, and M. Noeaki, J. Biochem. (Tokyo) 74, 375 (1973).

138

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T . ABBOTT

or when potassium ferricyanide is added to the solution under anaerobic condition (77). When the substrate, protocatechuic acid, is added under anaerobic conditions, the visible absorption shows an increase with a shift in the absorption peak from 465 to 480 nm, suggesting the formation of an enzyme-substrate complex (77). The spectrum of the enzyme-substrate complex decreases markedly when sodium dithionite is added, but returns to that of the original complex upon the addition of ferricyanide. Further addition of oxygen converts the spectrum to that of the original enzyme. These results suggest that the iron in the enzyme-substrate complex is also in the trivalent state (77). A series of stopped-flow experiments indicate the presence of a shortlived, new spectral species in the early stage of the reaction. This species is characterized by a broad absorption band with a maximum between 500 and 520 nm, which is distinctly different from those possessed by the enzyme and by the enzyme-protocatechuic acid complex. The band can be demonstrated only in the presence of both protocatechuic acid and molecular oxygen ( 4 9 ) . The rate constant for the decomposition of the new spectral species is calculated to be 5,640 min-l per active site of the enzyme. Assuming that the enl;yme contains eight active sites, the rate constant per mole of enzyme is calculated to be 45,120 min-l, which agrees quite well with a turnover number of 45,500 min-l for the enzyme ( 4 9 ) . A similar spectral species is also observed during the steady state of the reaction when enzymically active analogs such as 3,4-dihydroxyphenylacetic acid or 3,4-dihydroxyphenylpropionic acid are used (Fig. 1). Detailed analysis of the kinetics of the reaction reveals that the new spectral species observed with the enzyme is really an obligatory intermediate and is a ternary complex of oxygen, substrate, and enzyme, i.e., an oxygenated intermediate. The decomposition of the oxygenated intermediate is the rate-limiting step for the overall reaction. In order to form the oxygenated intermediate, the presence of an organic substrate is necessary. Therefore, it seems reasonable to assume that the organic substrate combines with the enzyme first and then reacts with oxygen to form the oxygenated intermediate [Eq. (16) ] (60). This is consistent E

k +IS

koOt\

.ES k-l k-r ESO*%E

+P

(16)

with the steady-state analyses mentioned above. If the enzymic reaction proceeds via the mechanism given by Eq. (16), K,,, values for organic substrate and oxygen should be expressed by k+3/k+land (k-2 k + , ) / l ~ + ~ , respectively, as the concentration of the other substrate in each case approaches infinity. The value of k,, is in good agreement with the turn-

+

3.

139

OXYGENASES: DIOXYGENASES

0.2'

500 Mlo Wavelength (nm)

400

700

Fxo. 1. Absorption spectra of the native enzyme, enzyme-substrate complex, and oxygenated intermediate of protocatechuate 3,4-dioxygenase with 3,4-dihydroxyphenylpropionate as substrate. The substrate lacks a visible absorption spectrum (60).Curve A. Spectrum of the native enzyme recorded without the substrate; curve B, spectrum of the enzyme-substrate complex recorded without oxygen; and curve C, spectrum of the enzyme recorded during the steady state of the reaction. (Reproduced with the permission of the J. B i d Chem.)

over number based on the active site of the enzyme. The values of lc+3/k+l and (k2 k+3)/lc+2also coincide roughly with the K,,, values for the organic substrate and oxygen, respectively (60). Electron spin resonance properties-Protocatechuate 3,4-dioxygenase also shows a sharp ESR signal a t g = 4.31, which is the result of the ferric ion bound to the enzyme. The signal height a t g = 4.31 diminishes instantaneously upon addition of substrate under anaerobic conditions, but the signal never completely disappears. These results suggest that the addition of substrate causes a modification in the ligand field of ferric ion in the enzyme rather than a change in valency from the ferric to the ferrous state. The decreased signal is restored to that of the original level when the substrate is completely converted upon the introduction of air (77). From a temperature dependence study of the ESR property, Peisach et al. (80) suggested that the ligands of iron are sulfur atoms that are arranged in a tetrahedron around the metal. However, the number of SH groups unmasked by the removal of iron is far less than that expected (78). Upon the addition of substrate or substrate analogs under anaerobic

+

80. J. Peisach, H. Fujisawa, W. E. Blumberg, and 0. Hayaishi, Fed. Proc., Fed. Amer. Soc. E x p . Biol. 31, 448 (1972).

140

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

conditions, new resonances a t g = 6.4 and 5.6 are observed which apparently result from Fe3+in a nearly tetragonal environment. The situation is analogous to the geometric environment of ligands that exists in heme, suggesting that the binding of substrate to the enzyme causes a change in ligand symmetry, which makes the iron accessible for 0, binding. 2. Extradiol Dioxygenases

a. Metapyrocatechase Molecular and catalytic properties-M etapyrocatechase [ catechol : oxygen 2,3-oxidoreductase (decyclizing) , EC 1.13.11.21 catalyzes the conversion of catechol to a-hydroxymuconic r-semialdehyde with the insertion of two atoms of molecular oxygen [Eq. (17)]. The enzyme has been obtained in crystalline form from extracts of Pseudomonas arvilla (ATCC

aoHflWH OH

+

02-

OH

(17)

CHO

23973) grown with benzoate as the major carbon source. The enzyme is extremely unstable in the presence of air (81,82). However, low concentrations of organic solvents such as acetone and ethanol protect the enzyme from inactivation. Hence, buffer solutions containing 10% acetone are used in all purification procedures. The first crystallization of a dioxygenase, metapyrocatechase, was achieved with this procedure (83,84). Crystalline metapyrocatechase has a specific activity of about 110 pmoles/min/mg of protein at 24O and contains 1 g-atom of iron per mole of enzyme, based on a molecular weight of 140,000 (86). The enzyme is activated about 3-fold following treatment with a combination of cysteine and ferrous ion under anaerobic conditions. The fully activated preparation has a specific activity of about 300 pmoles/min/mg of protein and appears to contain 3 4 g-atoms of iron per mole of enzyme (86,87). 81. Y . Kojima, N. Itada, and 0.Hayaishi, JBC 236,2223 (1961). H. Taniuchi, Y. Kojima, F. Kanetsuna, H. Ochiai, and 0. Hayaishi, BBRC 8, 97 (1962). 83. M. Nozaki, H. Kagamiyama, and 0. Hayaishi, BBRC 11, 65 (1963). 84. M. Nozaki, H. Kagamiyama, and 0. Hayaishi, Biochem. 2. 338, 582 (1963). 85. M. Nozaki, K. Ono, T. Nakazawa, S. Kotani, and 0. Hayaishi, JBC 243, 2682

82.

(1968). 86. M. Nozaki, Y. Kojima, T. Nakazawa,

H. Fujisawa, K. Ono, S. Kotani, 0. Hayaishi, and T. Yamano, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 347. Maruzen, Tokyo, 1966. 87. 8. Takemori, T. Komiyama, and M. Katagiri, Eur. J. Biochem. 23, 178 (1971).

3.

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141

The iron in the enzyme may be easily removed from the protein by dialyzing against buffer without acetone. The specific activity of the enzyme appears to be associated with its iron content. The iron seems to be of the ferrous form and oxidation by air or H,O, leads to inactivation of the enzyme. The inactivated enzyme is fully reactivated by incubation with a combination of ferrous ion and a reducing agent under anerobic conditions (85). The enzyme appears to consist of 3 4 subunits (88). Unlike intradiol dioxygenases, metapyrocatechase is colorless and shows neither significant absorption in the visible range nor ESR signal at around g = 4.3. However, the inactivated enzyme prepared by treatment with H,O, shows a broad signal around g = 4.2, which is characteristic of ferric ion. These results are consistent with the concept that the iron in the native enzyme is in the divalent state and that oxidation of the iron results in inactivation of the enzyme (89). The enzyme has broad substrate specificity ( 6 2 ) . The relative maximum rates (in parentheses) are as follows: catechol (loo), 4-methylcatechol (100), 3-methylcatechol (62) , 4-chlorocatechol (51), pyrogallol (33), protocatechualdehyde (21) , 3,5-dichlorocatechol (0.17), protocatechuic acid (0.15), and dopamine (0.014).The K , values for catechol and oxygen are 3 and 7 pM, respectively. The enzyme activity is competitively inhibited by a variety of nitrogen containing aromatic compounds, including o-phenanthroline, a,d-dipyridyl, m-phenanthroline, a-naphthoquinone, quinoline, and pyridine. Based on these results, a hydrophobic interaction is assumed to be involved in the binding of substrate to enzyme ( 8 5 ) . Double reciprocal plots of initial velocities with one fixed substrate and the other variable give rise to patterns of intersecting lines. o-Nitrophenol and m-phenanthroline, dead-end inhibitors of the enzyme, each inhibits the enzyme competitively with respect to the substrate, catechol, and noncompetitively with respect to the other substrate, oxygen. These results are consistent with an ordered bi-uni mechanism in which the organic substrate first combines with the enzyme, which then reacts with oxygen to form a ternary complex (79). Circular dichroism-The enzyme shows a strong negative C D band at 225 nm in the region expected of peptide backbone absorption and has an ordered structure with a relatively low content of a-helix. This ordered structure is partially destroyed by treatment with urea or alkali but not by removal of the bound iron. I n the vicinity of the side chain 88. M. Nozaki, H. Fujisawa, and S. Kotani, Proc. I n t . Congr. Biochem, Yth, 1967 p. 565 (1968). 89. 0. Hayaishi, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), p. 286. Wiley, New York, 1965.

142

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEZ T. ABBOTT

absorption bands (250-300 nm) , positive dichroic bands are detected at 265, 288, and 300 nm. However, no significant CD bands are observed in the visible range. Following catechol addition under anerobic conditions, a new negative band appears a t 317 nm, accompanied by a decrease in the band a t 300 nm. However, with the apoenzyme, no such change in CD is observed. The negative CD band a t 317 nm may conceivably result from an asynmetric configuration of the tyrosyl residue formed by the interaction between substrate and iron atom (90). b. S14-Dihydroxyphenylacetate 8,s-Dioxygenase. 3,4-Dihydroxyphenylacetate 2,3-dioxygenase [3,4-dihydroxyphenylacetate 2,3-oxidoreductase (decyclizing), EC 1.13.11.71 catalyzes the reaction shown in Eq. (18). The enzyme has been obtained in crystalline form from extracts

+ 0, HOOC&C

-

HOOC&C

fl::

(18)

CHO

of Pseudomonas ovalis grown with p-hydroxyphenylacetate as the major carbon source (91,92).Like metapyrocatechase, the enzyme is stabilized by the presence of low concentrations of organic solvents such as acetone and ethanol (9.1). The enzyme contains 4 5 g-atoms of iron per mole of enzyme, based on a molecular weight of approximately 100,000. On prolonged storage in air or treatment with p-mercuribenzoate, the enzyme is dissociated into subunits with a molecular weight of 30,000, accompanied by release of iron. It appears that most of the iron is involved in the association of subunits, but a t least 1 g-atom of iron is at the active site (93). Like metapyrocatechase, the native enzyme is colorless and the iron atoms are not detectable by ESR. When excess substrate is added under aerobic conditions, a signal a t g = 4.3 develops, which is characteristic of ferric ion and which disappears as oxygen in the solution is used up concomitant with the formation of product. When catechol, a slowly reacting substrate, is added to the enzyme solution under aerobic conditions, a sharp signal is observed at g = 4.3, accompanied by the appearance of a deep violet color. These results suggest that the ESR signal and violet color are characteristic properties of a ternary complex involv90. F. Hirata, A. Nakazawa, M. Noeaki, and 0. Hayaishi, JBC 246, 5882 (1971). 91. H. Kita, J . Biochem. (il’okgo) 58, 116 (1905). 92. H.. Kita, M. Kamimoto, S. Senoh, T. Adachi, and Y . Takeda, BBRC 18, 66 ( 1986). 93. S. Senoh, H. Kita, and M. Kamimoto, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 378. Maruzen, Tokyo, 1966.

3.

143

OXYGENASES: DIOXYGENASES

ing enzyme (Fe 111)-substrate-oxygen as an intermediate in the reaction (74).However, no kinetic analysis has been carried out to show whether or not the complex is an obligatory intermediate in the reaction. c. Protocatechuate 4,6-Dioxygenase. Protocatechuate 4,5-dioxygenase [protocatechuate: oxygen 4,5-oxidoreductase (decyclizing) , EC 1.13.11.81 catalyzes the reaction shown in Eq. (19). This enzyme was first described by Dagley and Patel (94). Its partial purification and properties have (19) HOOC

OH

HOOC

OH

been described by Cain (96). The enzyme has been also purified from extracts of p-hydroxybenzoate-induced Pseudomonas testeroni (96). An almost homogeneous preparation has been obtained from extracts of a pseudomonad grown with protocatechuic acid as the major carbon source (97). The most highly purified preparation has a specific activity of 160 pmoles/min/mg of protein a t 24O and a molecular weight of approximately 150,000. One gram-atom of iron is present per mole of enzyme. Although the enzyme is very unstable, being readily inactivated during storage a t Oo or room temperature, it can be protected by the addition of 10% ethanol. Rapid inactivation of the enzyme is also observed during catalysis (96,973. This inactivation is partially prevented by L-cysteine (96) and appears to result simply from the removal of iron since the inactivation can be fully reversed by the addition of ferrous ion (97). According to Zabinski et al. (98), the enzyme purified from Pseudomonas testosteroni contains four iron atoms per mole of enzyme (MW 140,000). The iron in the enzyme is not detectable by ESR. However, incubation of the enzyme with substrate under anerobic conditions generates a typical high-spin Fe3+signal a t g = 4.3. When the substrate becomes converted to product, the signal disappears. Mossbauer studies on 67Fe?+-reconstitutedenzyme indicate that either the iron atoms are in a low-spin ferrous state or the enzyme has two active sites, each containing two antiferromagnetically coupled high-spin ferric ions. d. Steroid Oxygenase. 3,4-Dihydroxy-9,10-secoandrosta-1,3,5 ( 10)-triene-9,17-dione-4,5-dioxygenase(steroid oxygenase) (EC 1.13.11.25) cata(10)-trienelyzes the conversion of 3,4-dihydroxy-9,10-secoandrosta-l,3,5 (lo), 9,17-dione [ (I) in Eq. (20) ] into 4,9-diseco-3-hydroxyandrosta-l 94. 95. 96. 97. 98.

S. Dagley and M. D. Patel, BJ 66, 227 (1957). R. B. Cain, Nature (London) 193, 842 (1962). S. Dagley, P. J. Geary, and J. M. Wood, BJ 109,559 (1968). K. Ono, M. Nozaki, and 0. Hayaishi, BBA 220, 224 (1970).

R. Zabinski, M. Miinck, P. M. Champion, and J. M. Wood, Biochemistry 11, 3212 (1972).

144

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

OH (1)

(II)

2-diene-5,9,17-trion-4-oicacid [ (11) in Eq. (20) ] (99). The steroid oxygenase also catalyzes the oxidative cleavage of several substituted catechols in addition to its natural substrate. However, substitution of an alkyl group adjacent to the dihydroxyl group is required for maximal activity. Among these, 3-isopropylcatechol is the most reactive artificial substrate. The enzyme has been purified to a state of apparent homogeneity from extracts of Nocardia restrictus induced with progesterone. Like other extradiol dioxygenases, this enzyme is also stabilized by acetone. The molecular weight of the enzyme is about 280,000. The enzyme has an ultraviolet absorption maximum at 280 nm and contains 1.13 g-atoms of ferrous ion per mole of enzyme (100). Metal chelating agents such as o-phenanthroline, 8-hydroxyquinoline1 and a,d-dipyridyl inhibit the enzyme noncompetitively with respect to both organic substrate and molecular oxygen. Sulfhydryl inhibitors inhibit the enzyme a t a concentration of 1 mM. I n 0.05 N NaOH, the enzyme is dissociated into identical subunits with sedimentation coefficient of 2.6 S (100). The steroid dioxygenase gives intersecting initial velocity plots that conform to a sequential mechanism. The product shows linear noncompetitive inhibition with respect to either organic substrate or oxygen, indicating the formation of a dead-end complex. Experiments with 4-isopropylcatechol, a structural analog of the organic substrate, show that the enzyme is inhibited competitively with respect t o the organic substrate and uncompetitively with respect to molecular oxygen, indicating an ordered bi-uni mechanism in which molecular oxygen is added first and then the organic substrate followed by the release of product (101). This mechanism is different from those of protocatechuate 3,4-dioxygenase and metapyrocatechase mentioned above. 3. Other Phenolic Dioxygenases

a. 3-Hydroxyanthranilate Oxygenase. 3-Hydroxyanthranilate oxygenase [ 3-hydroxyanthranilate :oxygen 3,4-oxidoreductase (decyclizing) , 99. D. T. Gibson, K. C. Wang, C. J. Sih, and H. Whitlock, Jr., JBC 241, 551 (1966). 100. H. H. Tai and C. J. Sih, JBC 245, 5062 (1970). 101. H. H. Tai and C. J. Sih, JBC 245, 5072 (1970).

3.

OXYGENASES : DIOXYGENASES

145

E C 1.13.11.61 catalyzes the cleavage of the benzene ring of 3-hydroxyanthranilic acid to yield a-amino-P-carboxymuconic semialdehyde as shown in Eq. (21) (10%).This enzyme is found in the liver and kidney

OH

of various species (10.3).The properties of the enzyme are very similar to those of the extradiol dioxygenases. The enzyme contains ferrous ion. It is also rather unstable in air and is inactivated during catalysis (104). This inactivation appears to result from either oxidation or removal of the bound iron (106). The inactivated enzyme is activated by ferrous ion on acidification to about pH 4 or exposure t o 6.8 M urea. Metal binding agents such as o-phenanthroline remove the bound iron and inactivate the enzyme. The substrate 3-hydroxyanthranilate prevents the removal of iron from the active enzyme by o-phenanthroline and also blocks complete reactivation of the enzyme from iron-free enzyme (106). The Michaelis constants for oxygen and 3-hydroxyanthranilate are 3.1 X lo-’ M and 4 x M , respectively. Lineweaver-Burk plots a t various fixed levels of the second substrate yield a set of parallel lines, suggesting that the binding of the first substrate is separated from the binding of the second substrate by a very slow irreversible reaction. Picolinic and quinolinic acids inhibit the reaction competitively with respect to 3-hydroxyanthranilate and uncompetively with respect to oxygen (106). b. Homogentisate Oxygenase. Homogentisate oxygenase [homogentisate: oxygen 1,2-oxidoreductase (decyclizing), EC 1.13.11.51 catalyzes the conversion of homogentisate to maleylacetoacetic acid with incorporation of two atoms of molecular oxygen [Eq. (22)l. The enzyme is distributed OH I

OH

102. A. H. Mehler, in “Oxygenases” (0. Hayaishi, ed.), p. 87. Academic Press, New York, 1962. 103. R. E.. Priest, A. H. Bokman, and B. S. Schweigert, Proc. SOC.E x p . Biol. M e d . 78, 477 (1951). 104. C. 0. Stevens and L. M. Henderson, JBC 234, 1188 (1959). 105. R. A. Mitchell, H. H. Kang, and L. M. Henderson, JBC 238, 1151 (1963). 106. N. Ogasawara, J. E. Gander, and L. M. Henderson, JBC 241, 613 (1966).

146

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

in both mammalian tissues and microorganisms. A requirement of ferrous ion for enzymic activity has been shown by Suda and Takeda (107,108) and others (109-111). Oxygen isotope experiments have shown the enzyme to be a dioxygenase (119). The enzyme has been prepared in crystalline form from extracts of Pseudmonus ftuoTescens adapted to tyrosine (113). The crystallized enzyme is homogeneous in the ultracentrifuge and has a molecular weight of about 380,000. Like the mammalian liver enzyme, bacterial homogentisate oxygenase requires ferrous ion as a cofactor. For maximal activity a t pH 6.0, the enzyme requires a t least 10 min of preincubation with ferrous ion, glutathione, and ascorbate, but a t pH 5.4 only ferrous ion is necessary. The K, values for homogentisate and ferrous ion a t pH 6.0 are 6 X and 1 X M , respectively. The homogentisate oxygenase in the soluble phase of calf liver extract has been partially purified (114,116). Like the extradiol dioxygenases, the enzyme is stabilized by the presence of 10% acetone (116). For activation a t pH 7.0 in phosphate buffer, the enzyme requires a combination of ascorbate, reduced glutathione, and ferrous ion. However, a t pH 5.3, in acetate buffer, ferrous ion alone fully activates the enzyme. These results suggest that activation is related to the soluble level of ferrous ion (114). The activation of the enzyme is also appreciably affected by the pH of the medium, with the rate of activation of the apoenzyme being greatly increased a t lower pH values. A pH 5.6, activation approaches a maximum within 2 hr, whereas a t pH 6.8, more than 24 hr are required for full activation ( 1 1 5 ) . c. 9,6-Dih ydroxyp yridine Oxygenase. 2,5-Dihydroxypyridine oxygenase [2,5-dihydroxypyridine :oxygen 4,5-oxidoreductase (decyclizing) , EC 1.13.11.91 was first described by Behrman and Stanier (116) with cell-free extracts of Pseudomonus putida. The enzyme catalyzes the oxidation of 1 mole of 2,5-dihydroxypyridine to equimolar amounts of maleamic and formic acids with the consumption of 1 mole of oxygen [Eq. 107. M. Suda and Y. Takeda, J . Biochem. ( T o k y o ) 37, 375 (1950). 108. M. Suda and Y. Takeda, J . Biochem. (Tokyo) 37,381 (1950). 109. D. I. Crandall, Fed. Proc., Fed. Amer. SOC. Exp. BioZ. 12, 192 (1953). 110. W. E. Knox and S. W. Edwards, JBC 216,479 (1955). 111. B. Schepartz, JBC 205, 185 (1953). 112. D. I. Crandall, R. C. Krueger, F. Anan, K. Yasunobu, and H. S. Mason, JBC 235, 3011 (1960). 113. K. Adachi, Y. Iwayama, H. Tanioka, and Y. Takeda, BBA 118, 88 (1966). 114. W. G. Flamm and D. I. Crandall, JBC 238, 389 (1963). 115. S. Takemori, E. Furuya, K. Mihara, and M. Katagiri, Eur. J . Bwchem. 6, 411 ( 1968). 116. E. J. Behrman and R. Y. Stanier, JBC 228, 923 (1957).

3.

OXYGENASES : DIOXYGENASES

HOD

O N

H

147

r

(23)]. Crystallization of the enzyme has been recently achieved by Gauthier and Rittenberg (117) from P. putida grown with sodium nicotinate as the major carbon source. The stoichiometry of the reaction has been established with crystalline enzyme. Studies with lSOz and H,lsO on the incorporation of oxygen-18 into products have also shown the enzyme to be a dioxygenase. Since N-formylmaleamic acid cannot be detected as an intermediate in the reaction and synthetic N-formylmaleamate is not a substrate for the enzyme, addition of molecular oxygen and hydrolytic cleavage of the N-formyl group appear to be catalyzed by a single enzyme in a concerted mechanism (118). The purified enzyme is labile, but it is stabilized by dithiothreitol. Activity lost on dialysis or purification is restored by incubation with dithiothreitol and ferrous sulfate. The enzyme is inhibited by sulfhydryl reagents and iron chelating agents. The molecular weight is estimated to be 242,000 by sucrose gradients containing dithiothreitol and the minimum molecular weight has been estimated to be 39,500 by sodium dodecyl sulfate (SDS) acrylamide gel electrophoresis. The enzyme is specific for 2,5-hydroxypyridine1 whereas the enzyme does not catalyze the oxidation of the following compounds: 2,3-dihydroxypyridine1 2,4-dihydroxypyridine1 2,6-dihydroxypyridine, 2-hydroxypyridine1 3-hydroxypyridine, 4-hydroxypyridine, dipicolinic acid, picolinic acid, nicotinic acid, 6-hydroxynicotinic acid, pyridoxal, pyridoxamine hydrochloride, catechol, and p-hydroxybenzoic acid (117). d. 2,s-Dihydroxyindole 2,s-Dioxygenase. 2,3-Dihydroxyindole 2,3-dioxygenase [ 2,3-dihydroxyindole :oxygen 2,3-oxidoreductase (decyclizing), EC 1.13.11.231 has been partially purified from an extract of a soil bacterium grown with indole as the sole source of carbon (119). The enzyme catalyzes the conversion of dihydroxyindole to anthranilic acid in the presence of oxygen with concomitant evolution of CO, [Eq. (2411. 117. J. J. Gauthier and S. C. Rittenberg, JBC 246, 3737 (1971). 118. J. J. Gauthier and S. C. Rittenberg, JBC 246, 3743 (1971). 119. M.Fujioka and H. Wada, BBA 158, 70 (1968).

148

OSAMU HAYAISHI,

MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

The enzyme does not require additional cofactor or metal ion for activity, but is moderately inhibited by o-phenanthroline. The purified enzyme shows no appreciable absorption spectrum except a t 280 nm, indicating that a heme cofactor is not involved in contrast with the other dioxygenases that cleave the indole ring (see Section 11).

B. CYSTEAMINE OXYGENASE Cydeamine oxygenase [ cysteamine :oxygen oxidoreductase, EC 1.13.11.19) catalyzes the conversion of cysteamine to hypotaurine with the incorporation of two atoms of molecular oxygen [Eq. (25)]. The enC%-SH I

C%-m?

+

02-

CH,- SO,H I Ch- NH,

zyme has been extensively studied by Cavallini and his colleagues (14,120-123). The enzymic activity is almost completely dependent on the presence of a catalytic amount of sulfide. Sulfide can be replaced by other compounds including elemental sulfur, elemental selenium, hydroxylamine, and redox dyes with E,,' higher than + O . O l l V (122,123). The enzyme is present in various tissues of different animals. It has been prepared in an almost homogeneous state from horse kidney ( ~ $ , l 2 4 ) . The enzyme contains 1 g-atom of iron per mole of enzyme, based on a molecular weight of 83,000 (123).The enzyme appears to consist of two very similar, possibly identical, subunits (125).The ESR spectrum 120. D. Cavallini, C. De Marco, and B. Mondovi, Nature (London) 192, 557 (1961). 121. D. Cavallini, C. De Marco, and R. Scandurra, Ztal. J. Biochem. 11, 196 (1962). 122. D. Cavallini, R. Scandurra, and C. De Marco, BJ 96, 781 (1965). 123. D. Cavallini, C. De Marco, R. Scandurra, S. DuprB, and M. T. Graziani, JBC 241, 3189 (1966). 124. D. Cavallini, C. Cannella, G. Federici, 9. Dupr6, A. Fiori, and E. D. Grosso, Eur. J . Biochem. 16,537 (1970). 125. G. Federici, D. Barra, A. Fiori, and M. Costa, Physiol. Chem. Phys. 3, 448 (1971).

3.

149

OXYGENASES : DIOXYGENASES

of the native enzyme shows a signal a t g = 4.3, typical of high-spin Fe(II1) in a ligand field of rhombic symmetry. The intensity of this signal indicates that nearly all of the iron in the enzyme is in the ferric state. The g = 4.3 signal is the same both in the presence or absence of oxygen, but is modified when the substrate, cysteamine, is added in the absence of oxygen. The modified signal shifts back to that of the original signal following the introduction of air. On adding sulfide ion, the signal a t g = 4.3 decreases and a new signal appears a t g = 7.25. The latter is abolished by the addition of substrate in the presence of air. These results indicate that changes in the environment of the iron occur following the addition of either activator or substrate. These changes are specific and fully reversible. It appears, therefore, that the ferric form of nonheme iron is involved in the catalytic action of cysteamine oxygenase (16). It has been suggested that superoxide anion may also be involved in the enzyme reaction (126).

C. CYSTEINE OXYGENASE In 1965, Sorbs and Eureta reported that rat liver cytoplasm catalyzes the net oxidation of L-cysteine to L-cysteinesulfinic acid in the presence of O,, NADPH, F e ( I I ) , and hydroxylamine [Eq. (26)] (17). I n view C&I H-C-NH, I

SH

COOH

C&-

+

I

0

2

hH-C-NH,

SO,H

I COOH

(26)

of the cofactor requirements, they proposed that the enzyme be classified as a monooxygenase (17). Wainer independently demonstrated the oxydation of cysteine to cysteine sulfinate by rat liver extracts (127). The enzyme appears to be specific for L-cysteine since very little sulfinate formation is detected when D-cysteine, L-cystine, glutathione, or cysteamine is used as substrate. The enzyme is inhibited by heavy metal reagents such as EDTA, cyanide, o-phenanthroline, and sulfhydryl reagents (p-hydroxymercuribenzoate and iodoacetate) , but not by arsenite. Heatlabile particulate fractions also stimulate the reaction (168). However, Lombardini et al. (18) have failed to demonstrate stimulation of cysteine sulfinate synthesis by microsomal or mitochondria1 factor. Instead, they have found an additional heat-stable cofactor which has a molecular weight of approximately 750 (18). Experiments with lSO indicate that 126. G. Federici, A. Fiori, M. Costa, E. Barboni, and Biochem., gth, 1973 7a4, p. 331 (1973). 127. A. Wainer, BBA 128, 296 (1966). 128. L.Ewetz and B. Sorbo, BBA 128, 296 (1966).

S.DuprQ, Proc. Znt. Congr.

150

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

the reaction is catalyzed by a dioxygenase rather than by two monooxygenases acting sequentially (19).

D. LIPOXYGENASE Lipoxygenase (linoleate: oxygen oxidoreductase, E C 1.13.11.12) catalyzes the hydroperoxidation of polyunsaturated fatty acids and esters containing a &,cis-l,4-pentadiene system [Eq. (27) 1. Theorell et al. CH,-

(C€i,),-

CH=CH-C€i,-CH=CH-(CH*),-COOH

C&-

(C€LJ,-

CH=CH-CH=CH-CH-(C~),I OOH

+

0,

f COOH

(129) first obtained the enzyme in a crystalline form from soybean and reported that the enzyme did not require a metal or prosthetic group. This conclusion was based on the absorption spectrum and metal analysis of the enzyme. Recently, however, three groups of investigators have demonstrated that the enzyme is an iron-containing dioxygenase (130-139). Metal analyses of highly purified lipoxygenase by atomic absorption, flameless atomic absorption, and spectrophotometric determination of Fe?+ (o-phenanthroline), have demonstrated the presence of 1 g-atom of iron per mole of enzyme, based on a molecular weight of 102,000 (130,131). The iron is tightly bound to the enzyme and is not released from the protein by well-known complexing agents (130). However, preincubation of the enzyme at pH 9 with chelating agents results in substantial inhibition (131). The removal of iron from the protein is achieved by using a strong Fez+chelator such as o-phenanthroline, but only after the enzyme has been treated with a reducing agent. The formation of the o-phenanthroline Fez+ complex is accompanied by a loss of enzymic activity (132).Although the ESR experiments have failed to show a signal a t g = 4.3, as has been observed with all dioxygenases containing ferric ion, the iron in lipoxygenase is believed to be in the ferric state (132).

E. CONCLUDING REMARKS Nonheme iron appears to be the most common cofactor of dioxygenases. The valence state of the iron involved may be either in the ferric or fer129. H. Theorell, R. T.Holman, and A. Akeson, Actu Chem. Scund. 1, 671 (1947). 130. M. Roza and A. Francke, BBA 327,24 (1973). 131. H. W. 5. Chan, BBA 327, 32 (1973). 132. E. K. Pistorius and B. Axelrod, JBC 249, 3183 (1974).

3.

151

OXYGENASES : DIOXYGENASES

row form. I n either case, the iron in the oxygenase has been assumed to be the site of oxygen activation. The iron also appears to be closely associated with the substrate binding site, although the substrate may bind primarily to the protein moiety (72,78). In some dioxygenases, iron may also be involved in the association of subunits (95,100). It is of interest to note that the intradiol dioxygenases contain the ferric form of iron, whereas the extradiol enzymes contain the ferrous form (72). The reaction mechanisms for both types of enzymes, however, appear to be the same; namely, a ternary complex of enzyme, substrate, and oxygen is formed in all reactions catalyzed by dioxygenases. Both substrate and oxygen are presumed to be activated in the ternary complex and react concertedly to form the oxygenated end product. Questions as to why some enzymes contain the ferric ion and others the ferrous ion, whether or not valency change of the iron is involved during the catalysis, and what is the active form of oxygen, require further investigations.

IV. a-Ketoglutarote Dioxygenarer

A. INTRODUCTION The a-ketoglutarate dioxygenases catalyze a variety of oxidations which range from the hydroxylation of a saturated carbon atom to the conversion of an aldehydic carbon to a carboxyl carbon. These intermolecular dioxygenase reactions are described by the following general equation :

ymH o z + s + cH' I

FH'

reduciine agent

-so

Fez+

YOOH

+

CHZ I

7%

+ co,

COCOOH

COOH

a-Ketoglutarate

Succinate

(28)

The requirement for Fez+is highly specific, whereas the requirement for a reducing agent can be fulfilled by several types of reductants including ascorbate and some sulfhydryl containing compounds. The discovery by Hutton et al. (155) of the dependency of the prolyl hydroxylase reaction on a-ketoglutarate has not only led to much insight into the area encompassed by the prolyl and lysyl hydroxylases but also into that of 7-butyrobetaine hydroxylase, pyrimidine deoxyribonucleoside 2'-hydroxylase, the three sequential reactions of thymine 7-hydroxylase, 133. J. J. Hutton, A. L. Tappel, and S. Udenfriend, ABB 118, 231 (1967).

152

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

and the related a-keto acid dioxygenase, p-hydroxyphenylpyruvate hydroxylase. Several articles have appeared which present the events leading up to our current knowledge of these enzymes (134-145), and the very recent review of the prolyl and lysyl hydroxylases by Cardinale and Udenfriend (144) is most penetrating and thorough. I n the article a t hand, the known a-keto acid dioxygenases, which oxidize substrates as diverse as proteins and pyrimidines and which are found in animals, plants, and microorganisms, are discussed individually.

B. PROLYL HYDROXYLASE H +

O2

+

Q o=cI c I

R’

NHR 0

0-ketoglutarate

reducing HO agent

Fe2+

+ succinate Q

c

o=cI

z

0

NHR

+

coz

(29)

I

R’

1. Introduction Prolyl hydroxylase (EC 1.14.11.2) converts proline in peptidyl linkage to trans-4-hydroxyproline which in vertebrate tissues is found primarily in collagen. I n contrast to the hydroxyl groups of hydroxylysine in collagen, those of hydroxyproline are not substituted, and only recently has come some comprehension of the function of this imino acid through the demonstration that it significantly contributes to the thermal stability 134. S. Udenfriend, Science 152, 1335 (1966). 135. S. Udenfriend, in “The Chemistry and Molecular Biology of the Intercellular Matrix” (A. E. Balazs, ed.), Vol. 1, p. 371. Academic Press, New York, 1970. 136. D. J. Prockop, in “The Chemistry and Molecular Biology of the Intercellular Matrix” (A. E. Balazs, ed.), Vol. 1, p. 335. Academic Press, New York, 1970. 137. M. E. Grant and D. J. Prockop, N. Engl. J. Med. 286, 194, 242, and 291 (1972). 138. M. J. Barnes and E. Kodicek, Vitam. Harm. (New York) 30, 1 (1972). 139. P. M. Gallop, 0. 0. Blumenfeld, and S. Seifter, Annu. Rev. Biochem. 41, 617 (1972). 140. L. E. Bennett, Progr. Inorg. Chem. 18, 1 (1973). 141. G. J. Cardinale and S. Udenfriend, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 195. Univ. Park Press, Baltimore, Maryland, 1973. 142. I. Kuttan and A. N. Radhakrishnan, Advan. Enzymol. 37, 273 (1973). 143. P. Bornstein, Annu. Rev. Biochem. 43, 667 (1974). 144. G. J. Cardinale and S. Udenfriend, Advan. Enzymol. (in press), 145. M. T. Abbott and S. Udenfriend, in “Molecular Mechanisms of Oxygen Activation’’ (0. Hayaishi, ed.), p. 167. Academic Press, New York, 1974.

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OXYGENASES: DIOXYGENASES

153

of the triple helical conformation of collagen (146-149). Thus, hydroxyproline stabilizes collagen in a form which is less susceptible to protease action a t physiological temperature and which may be required for secretion of collagen a t an optimal rate (150-158). The tissues in which prolyl hydroxylase activity has been demonstrated are many (169-174), but in general this activity appears to be higher in tissues engaged in the synthesis of collagen. Therefore, embryonic and fetal tissues had higher activities than did the homologous tissues from adult animals (159).Similarly, prolyl hydroxylase activity was elevated in response to injury (159,160,175-179), in neoplasms (161,180), and in 146. S. Sakakibara, K. Inouye, K. Shudo, Y. Kishida, Y. Kobayashi, and D. J. Prockop, BBA 303, 198 (1973). 147. R. A. Berg and D. J. Prockop, BBRC 52, 115 (1973). 148. S. Jimenes, M. Harsch, and J. Rosenbloom, BBRC 52, 106 (1973). 149. J. Darnell, S. Jimenez, L. Murphy, M. Harsch, and J. Rosenbloom, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 33, 1596 (1974). 150. K . Juva, D. J. Prockop, G. W. Cooper, and S. W. Lash, Science 152,92 (1966). 151. P. B. Ramaley and J. Rosenbloom, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 15, 59 (1971). 152. P. Dehm and D. J. Prockop, BBA 240, 358 (1971). 153. R. L. Margolis and L. N. Lukens, A B B 147, 612 (1971). 154. J. Rosenbloom and D. J. Prockop, JBC 246, 1549 (1971). 155. S. A. Jimenez, P. Dehm, B. R. Olsen, and D. J. Prockop, JBC 248,720 (1973). 156. C. J. Bates, C. J. Prynne, and C. I. Levene, BBA 263,397 (1972). 157. B. Peterkofsky, A B B 152, 318 (1972). 158. B. Peterkofsky, BBRC 49, 1343 (1972). 159. E. Mussini, J. J. Hutton, and S. Udenfriend, Science 157, 927 (1967). 160. T. Takeuchi, K. I. Kivirikko, and D. J. Prockop, BBRC 28, 940 (1967). 161. N. Roberts and S. Udenfriend, J. Nut. Cancer Znst. 45, 277 (1970). 162. J. Halme and M. Jaaskelainen, BJ 116,367 (1970). 163. B. Peterkofsky and S. Udenfriend, JBC 238, 3966 (1963). 164. D. J. Prockop and K. Juva, Proc. N u t . Acud. Sci. U . S. 53, 661 (1965). 165. B. Peterkofsky and S. Udenfriend, Proc. Nut. Acud. Sci. U . S. 53, 335 (1965). 166. J. J. Hutton and S. Udenfriend, Proc. N u t . Acad. Sci. U . S. 56, 198 (1966). 167. B. Goldberg and H. Green, Nature (London) 221, 267 (1969). 168. A . Nordwig and F. K . Pfab, BBA 181, 52 (1969). 169. K.-Y. T. Kao, C. R. Treadwell, J. M. Previll, and T. H. McGavack, BBA 151, 568 (1968). 170. H. 0 . Stein, H. R. Keiser, and A. Sjoerdsma, Lancet 1, 106 (1970). 171. R. 0. Langner and G. C. Fuller, BBRC 36,559 (1969). 172. H. Green and B. Goldberg, Proc. N u t . Acad. Sci. U . S. 53, 1360 (1965). 173. U. Langness and S. Udenfriend, Proc. N u t . Acad. Sci. U . S. 71, 50 (1974). 174. A. Ooshima, G. C. Fuller, G. J. Cardinale, S. Spector, and S. Udenfriend, Proc.

Nut. Acad. Sci. U. S. (in press). 175. J. Halme, J. Uitto, K. Khanpaa, P. Karkunen, and S. J. Lindy, J . Lab. Clin. M e d . 75, 535 (1970). 176. G . C. Fuller and R. 0. Langner, Science 168, 987 (1970).

154

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

various diseased and abnormal states (162,174,181-184) in which rapid collagen synthesis occurs. Prolyl hydroxylase activity has also been found in the muscle layers of the enteric roundworm Ascaris lumbricoides (1861, the earthworm body wall (168),plant tissues (186),and certain microorganisms which synthesize hydroxyproline containing actinomycins (187). The earlier names for prolyl hydroxylase, i.e., “collagen proline hydroxylase” (lSS), L‘protocollagen hydroxylase” (188), “protocollagen proline hydroxylase” (189), and “peptidyl proline hydroxylase” (190) reflect the first studies of this enzyme having been carried out with tissue associated with collagen synthesis. I n view of the substrate specificity of the enzyme, the name “prolyl hydroxylase” was recently proposed (146). 2. Purification and Molecular Properties

Rhoads and Udenfriend (191) purified prolyl hydroxylase 500-fold from the skin of newborn rats. Their purification procedure yielded an enzyme preparation that sedimented as a homogeneous substance in the ultracentrifuge and was estimated to be 90% pure by polyacrylamide gel electrophoresis. This preparation had a specific activity of 86-100 units/mg (192), with reduced Ascaris cuticle collagen as substrate, and possessed an absorption spectrum characteristic of a simple protein suggesting that cofactors such as flavins or pyridine nucleotides were not bound to the enzyme. The molecular weight was determined by several methods and estimated to be approximately 130,000. When the enzyme 177. H. L. Crossley, A. R. Johnson, K. K. Mauger, N. L. Wood, and G. C. Fuller, Life Sci. 11, 869 (1972). 178. G. C. Fuller, E. Miller, T. Farber, and E. Vanloon, Connect. Tissue Res. 1, 217 (1972). 179. R. 0. Langner and G. C. Fuller, Atherosclerosis 17, 463 (1973). 180. K. R. Cutroneo, N. A. Guzman, and A. G. Liebelt, Cancer Res. 32,2828 (1972). 181. J. Uitto, S. Lindy, P. Rokkanen, and K. Vainio, Clin. Chim. Acta 30, 741 (1970). 182. J. Uitto, J. Halme, M. Hannuksela, P. Peltokallio, and K. I. Kivirikko, Scand. J . Clin. Lab. Invest. 23, 241 (1969). 183. R. A. Salvador and I. Tsai, Biochem. Pharmacol. 22, 37 (1973). 184. R. A. Salvador and I. Tsai, ABB 154,583 (1973). 185. D. Fujimoto and D. J. Prockop, JBC 244,205 (1969). 186. D. Sadava and M. J. Chrispeels, BBA 227,278 (1971). 187. E. Katz, D. J. Prockop, and S. Udenfriend, JBC 237,1585 (1962). 188. K. I. Kivirikko and D. J. Prockop, Proc. N u t . Acad. Sn‘. U.S. 57, 782 (1967). 189. K. I. Kivirikko, H. J. Bright, and D. J. Prockop, BBA 151, 558 (1968). 190. J. O’D. McGee and 8. Udenfriend, ABB 152,216 (1972). 191. R. E. Rhoads and S. Udenfriend, ABB 139, 329 (1970). 192. A unit of activity with respect to the dioxygenases considered in Section I11 is defined as the amount of enzyme which catalyzes the oxidation of 1 nmole

of substrate per minute under the conditions of the standard assay.

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155

was treated with SDS and 2-mercaptoethanol and then subjected to polyacrylamide electrophoresis under denaturing conditions, two bands of equal intensity and similar mobility were obtained. The molecular weights of the proteins in these bands were determined to be about 65,000, and thus the enzyme of molecular weight 130,000 appeared to consist of two dissimilar subunits (191). Recent gel filtration studies with prolyl hydroxylase from mouse and rat skin showed that the enzyme may also exist in an active form which has a molecular weight in the range of 300,000400,000 (144). A subunit structure was also indicated for the prolyl hydroxylase from chick embryos (193,194) since its molecular weight was reduced from about 248,000 to 110,000 upon either ultracentrifugation in 5.8 M guanidine or subjection of dodecyl sulfate polyacrylamide gel electrophoresis. The isoelectric point of the enzyme was determined to be 4.4, which is consistent with the high proportion of acidic amino acids contained in the enzyme (193) as well as in the one from rat skin (191). Recently, the prolyl hydroxylase from chick embryo has been purified to homogeneity (specific activity, 540 units/mg) with an affinity column procedure devised by Berg and Prockop (195). Their procedure involves linking agarose to a peptide substrate with a high affinity for the enzyme, i.e., reduced and carboxymethylated collagen from the cuticle of Ascaris lumbricoides, and specifically eluting the enzyme from the affinity column with a high concentration of a second peptide substrate of lower affinity, i.e., poly (L-Pro-Gly-L-Pro) with an average molecular weight of 2400. In this manner large yields of the enzyme were obtained in a short time. The purity of the enzyme was established by gel filtration] ultracentrifugation, and polyacrylamide gel electrophoresis of the enzyme in the native and dissociated states and by production of specific antibodies in rabbits immunized with the enzyme (196,196). The amino acid analysis of this enzyme preparation gave results which differed from those previously reported for the less pure enzyme prepared by the method of Halme e t a l . (193). Studies (195) with the homogeneous enzyme from chick embryos indicated that prolyl hydroxylase is a tetramer (MW, 230,000) which consists of identical dimers. Berg and Prockop (195) suggested that the structural integrity of the enzyme is maintained by either intrachain or interchain disulfide bonds since the enzyme was dissociated into a mixture of monomers and dimers by exposure to either dithiothreitol or mercaptoethanol. 193. J. Halme, K. I. Kivirikko, and K. Simons, BBA 198,460 (1970). 194. M. Pankalainen, H. Aro, K. Simons, and K. I. Kivirikko, BBA 221,559 (1970). 195. R. A. Berg and D. J . Prockop, JBC 248, 1175 (1973). 196. R. A. Berg, B. R. Olsen, and D. J. Proekop, BBA 285, 167 (1972).

156

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

The monomers were shown to be enzymically inactive and to be of two types with molecular weights of 60,000 and 64,000 by polyacrylamide gel electrophoresis in SDS. The dimer form appeared to be active but had less than one-seventh of the specific activity of the tetramer form, and the data did not rule out the possible association of the dimers into tetramers under the conditions of the enzymic assay. Likewise, the apparent differences between the enzymes from chick embryo and rat skin may be a function of a more facile interconversion of dimer and tetramer rather than of a more active dimer. Electron microscopy of the pure enzyme from chick embryos has further detailed its subunit st.ructure (197’).The monomers appeared to be rod-shaped with a diameter of 3.3 nm and a length of 7.0 nm. Electron micrographs of preparations in which the protein was partially dissociated indicated that the dimers consist of two monomers joined a t one end so that they form a V. The largest regular protein structures seen in preparations of the enzyme were about 8 X 8 nm. These appeared to be the tetramers since in some of the structures four substructures were identified. Olsen et al. (197) proposed a model of the tetramer in which two V-shaped dimers are interlocked.

3. Catalytic Properties Prior to Peterkofsky’s and Udenfriend’s development (163) of the first cell-free system which could catalyze the prolyl hydroxylase reaction, molecular oxygen (198-201) and ascorbate (202-204) were implicated in the reaction, and proline appeared to be converted to hydroxyproline during or after the process of peptide bond formation (i?05-i?08). With the cell-free system, advantage was taken of the requirement for oxygen and of inhibitors of protein synthesis so that the period in which proline was incorporated into protein could be distinguished from the subsequent hydroxylation step. This allowed the authors to show conclusively that hydroxylation must occur after proline is incorporated into peptide link197. B. R. Olsen, R. A. Berg, K. I. Kivirikko, and D. J. Prockop, Eur. J . Biochem. 35, 135 (1973). 198. D. Fujimoto and N. Tamiya, BJ 84, 333 (1962). 199. D. J. Prockop, A. Kaplan, and S. Udenfriend, BBRC 9, 162 (1962). 200. D. Fujimoto and N. Tamiya, BBA 69, 559 (1963). 201. D. J. Prockop, A. Kaplan, and S. Udenfriend, ABB 101, 499 (1963). 202. W. van B. Robertson and B. Sehwartz, JBC 201,689 (1953). 203. W. van B. Robertson and J. Hewitt, BBA 49, 404 (1961). 204. N. Stone and A. Meister, Nature (London) 194,555 (1962). 205. M. R. Stetten and R. Schoenheimer, JBC 153, 113 (1944). 206. M. R. Stetten, JBC 181,31 (1949). 207. E. Hausmann and W. F. Newman, JBC 236, 149 (1949). 208. N. M. Green and P. A. Lowther, BJ 71, 55 (1959).

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157

age (163). The development of a cell-free system also allowed the more definitive work to be done in establishing the requirements for Fez+ (163,209-211) and ascorbate (165,211),and it to be discovered that another dialyzable substance was necessary for prolyl hydroxylase activity (133,212).T ha t the dialyzable substance was a-ketoglutarate was indicated by the loss of activity of crude undialyzed preparations upon their incubation with glutamic dehydrogenase, NADPH and NH,Cl, by the restoration of the activity upon addition of a-ketoglutarate (133) and by the high specificity of this requirement (133,212). A similar stringency was seen in the Fe?+requirement, and studies with various chelators indicated that the iron is bound to a hydrophobic region of the enzyme (133). However, there is disagreement over how tightly the metal ion is bound. Kivirikko et al. (188,189) suggested that ferrous iron is bound loosely to the enzyme, but studies (ZlS,Zl4) with radioactive iron indicate that it cannot be removed from the enzyme even after denaturation and subsequent treatment with strong chelating agents. I n addition, Hurych et al. (215) have recently reported spectrophotometric evidence not only for the firm binding of ferrous iron to the enzyme but also for oxidation to the ferric form on interaction with molecular oxygen. On the other hand, this work of Hurych and co-workers did not appear to exclude the possibility that the iron they detected was bound to protein other than prolyl hydroxylase. Moreover, Pankalainen and Kivirikko (616) have shown in spectrophotometric studies with highly purified preparations from chick embryos that far less than one mole of iron is bound per mole of eneyme. The requirements for a reducing agent was shown to be absolute but relatively nonspecific. Although ascorbate appeared most effective in meeting this requirement, other reductants such as dithiothreitol, tetrahydropteridines, reductones and analogs of ascorbate were also effective (133,165,191).The apparent K , values for each of the components of the prolyl hydroxylase system are shown in Table VI. As purer enzyme 209. M. Chvapil, M. Ehrlichova, and J. Hurych, Ezperientiu 22, 584 (1966). 210. D. J. Prockop and K. Juva, Proc. N u t . Acnd. Sci. U . S. 53, 661 (1965). 211. J. J. Hutton and S. Udenfriend, Proc. N u t . Acud. Sci. U . S. 56, 198 (1966). 212. J. J. Hutton, A. L. Tappel, and S. Udenfriend, BBRC 24, 179 (1966). 213. J. Rencovh, J. Hurych, J. Rosmus, and M. Chvagil, Abstr. 6th M e e t . , Fed. Eur. Biochem. SOC.,p. 82 (1968). 214. J. Rencovb, J. Hurych, J. Rosmus, and M. Chvapil, Abstr., Congr. Rheumutol. Znt., 12th, Ahstr. No. 708 (1969). 215. J. Hurych, P. Hobza, J. Rencovb, and R. Zahrradnik, in “The Biology of Fibroblasts” (E. Kulmen and J. Pikkaramen, eds.), p. 365. Academic Press, New York, 1973. 216. M. Pankalainen and K. I. Kivirikko, BBA 229, 504 (1971).

158

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

TABLE V I APPARENT K , VALUES FOR SUBSTRATES AND COPACTORS OF PROLYL HYDROXYLASE’ ~

Reactant Fe¶+ AsCorbate a-Ketoghtar ate Prolyl Substrate* Oxygen

K , for rat skin enzyme 0.001 m M 0 . 4 mM 0.01 mM 0 . 1 mM

-

K , for chick embryo enzyme

Ref.

191 141 191

0.005 m M

822

0.3 m M 0.01 mM

189

230

0.06 m M 6 vol % 2 . 6 vol %

189

Ref.

868

189

13s

~

From Abbott and Udenfriend (146). (Pro-Gly-Pro), substrates in the range 2000-8000 MW.

* Utilizing

preparations were obtained, additional factors were discovered which had stabilizing and stimulatory effects. One such factor is catalase (191,217) which appeared to have, in part, a “protein” effect (bovine serum albumin and heat denatured catalase were also stimulatory) and a ‘‘H202’’effect similar to that originally proposed by Kaufman and co-workers (218) for dopamine-p-hydroxylase. The effect of albumin did not appear to be the well-known one of stabilizing the enzyme during storage but one of stimulating the rate of the prolyl hydroxylase reaction (217). This stimulatory effect of albumin was somewhat mimicked by low concentrations of dithiothreitol, but the two compounds could not entirely replace one another (217 ) . Popenoe et al. (219) further distinguished between the effects of the two compounds and, in addition, showed that the enzyme from chick embryos was inactivated by sulfhydryl reagenb such as p-mercuribenzoate. Large amounts of glycine have been reported to stabilize purified preparations of the chick embryo enzyme (220), but the effect is less pronounced with the rat skin enzyme (144). The prolyl hydroxylases from both sources have pH optima near neutrality (191,195). The more decisive data on substrate specificity have come from experiments carried out with synthetic polypeptides (189,221-%29) and with 217. 218. 219. 220.

R. E. Rhoads, J. J. Hutton and S. Udenfriend, ABB 122, 805 (1967). E. Y. Levin, B. Levenberg, and S. Kaufman, JBC 235, 2080 (1960). E. A. Popenoe, R. B. Aronson, and D. D. Van Slyke, ABB 133, 286 (1969). J. Halme and K. I. Kivirikko, FEBS (Fed. Eur. Biochem. SOC.)Lett. 1, 223

(1968). 221. K. I. Kivirikko and D. J. Prockop, ABB 118, 611 (1967). 222. K. 1. Kivirikko and D. J. Prockop, JBC 242,4007 (1967). 223. J. J. Hutton, A. Marglin, B. Witkop, J. Kurtz, A. Burger, and S. Udenfriend, ABB 125, 779 (1968). 224. Y. Kikuchi, D. Fujimoto, and N. Tamiya, BJ 115,669 (1969).

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OXYGENASES : DIOXYGENASES

159

analogs of the peptide hormone bradykinin (22~7,230,231)as substrates for purified preparations of prolyl hydroxylase. The results indicated that the enzyme cannot hydroxylate proline or the polymer poly (L-proline) and that the minimum sequence requirement is an intact -X-Pro-Glytriplet in which there is considerable lattitude for variation in amino acid X. Moreover, the affinity of the enzyme for individual sequences which contain proline and the rate at which they are hydroxylated are dependent on adjacent amino acid residues and the size of the polymer. A controversy has existed ( 1gs,189,224,228,2g2-234) over whether the triple helical conformation of collagen prevents hydroxylation. However, the work of Berg and Prockop (235)strongly suggests that only prolyl residues in random coil chains can be hydroxylated. Rhoades et al. (232) found that although denaturation of the substrate did increase the degree to which susceptible prolyl residues could be hydroxylated, only 85% of the theoretical limit was reached. Pertinent in this regard appear to be studies on the formation of enzyme-substrate complexes (236) which showed that the affinity of the enzyme for substrate decreased upon partial hydroxylation of the substrate as well as the previously mentioned work ( 147-149) which indicated that hydroxylation stabilizes collagen in its triple helical conformation. The use of (Pro-Pro-Gly) as substrate for prolyl hydroxylase allowed Kivirikko et al. (237) to detect an aspect of asymmetry in the substrate specificity requirements in that the fourth triplet from the NHa-terminal end of this peptide was hydroxylated to a greater extent than any other triplet. Interestingly, even prolyl residues at the same site in identical peptides, obtained by CNBr cleavage of col225. R. E. Rhoads and S. Udenfriend, ABB 133, 108 (1969). 226. K. I. Kivirikko, D. J. Prockop, G. P. Lorenzi, and E. R. Blout, JBC 244, 2755 (1969). 227. K. Okada, Y. Kikuchi, Y. Kawashiri, and M. Hiramoto, FEBS (Fed. Eur. Biochem. Soc.) Lett. 28, 226 (1972). 228. K. I. Kivirikko, Y. Kishida, S. Sakakibara, and D. J. Prockop, BBA 271, 347 (1972). 229. R. S. Bhatnagar and R. S. Rapaka, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 33, 1596 (1974). 230. J. O’D. McGee, R. E. Rhoads, and S. Udenfriend, ABB 144,343 (1971). 231. J. O’D. McGee, M. H. Jimenez, A. M. Felix, G. J. Cardinale, and S. Udenfriend, ABB 154, 483 (1973). 232. R. E. Rhoads, S. Udenfriend, and P. Bornstein, JBC 246, 4138 (1971). 233. L. N. Lukens, Proc. Nut. Acad. Sci. U . S. 55, 1235 (1966). 234. A. Nordwig and F. K. Pfab, BBA 154, GO3 (1968). 235. R. A. Berg and D. J. Prockop, Biochemistry 12,3395 (1973). 236. K. Juva and D. J. Prockop, JBC 244, 8486 (1969). 237. K. I. Kivirikko, K. Suga, Y. Kishida, S. Sakakibara, and D. J. Prockop, BBRC 45, 1591 (1971).

160

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT 40r

__,-

- - - - - &

0

40

-

1 , _I-80 120 Time (minutes)

-a

,

160

F I ~2.. Plot against time of hydroxyproline formed ( A ) , a-ketoglutarate consumed

(01, and a-ketoglutarate consumed in the absence of H(Pro-Gly-Pro),OH

(0).

The amount of a-ketoglutarate added was 378 mpmoles. From Rhoads and Udenfriend (241).

lagen isolated from different sources, were shown to have been hydroxylated in vivo to different extents (232,238).A number of polypeptides can inhibit the prolyl hydroxylase reaction. Polyproline I1 (with peptide bonds in the tram position) (223,239),analogs of bradykinin ( 2 3 1 ) ,and polytripeptides (223,226) such as (Gly-Pro-Gly) were shown to inhibit competitively. The degree of inhibition was found to increase with the size of the polymer (223,239). All of the hydroxyproline synthesized by essentially homogeneous prolyl hydroxylase was shown to be 4-hydroxyproline. No 3-hydroxyprolyl residues were detected upon amino acid analysis of the product (195). Fujita et al. (240) demonstrated that the hydroxylation process displaces the 4-trans hydrogen of the prolyl residue. Succinate was identified as a product of the prolyl hydroxylase reaction and the amount of radioactive CO, formed from a- [ 1-14C]ketoglutarate was shown to be equal, on a molar basis, to the amount of hydroxyproline formed (Fig. 2 ) (241).Mass spectral analysis of the products produced when the reaction was carried out in an atmosphere of ISO,, with (Pro-Gly-Pro)-,, as substrate, revealed that one atom of molecular oxygen was incorporated into hydroxyproline and another into succinate (242). 238. P. Bornstein, Biochemistry 6, 3081 (1967). 239. D. J. Prockop and K. I. Kivirikko, JBC 244, 4838 (1969). 240. Y. Fujita, A. Gottlieb, B. Peterkofsky, S. Udenfriend, and B. Witkop, JACS 86, 4709 (1964). 241. R. E. Rhoads and S. Udenfriend, Proc. Nut. Acad. Sci. U.S. 60, 1473 (1968). 242. G. J. Cardinale, R. E. Rhoads, and S. Udenfriend, BBRC 43, 637 (1971).

3.

OXYGENASES : DIOXYGENASES

161

4. Assays

The earliest assays involved incubation of the enzyme and cofactors with a n underhydroxylated collagen which contained radioactive proline, hydrolysis of the hydroxyIated protein, isolation of hydroxyproline, and measurement of its radioactivity. The underhydroxylated collagen was prepared by incubation of labeled proline with slices or minces of collagen-forming tissues in the presence of an inhibitor, usually the Fez+ chelator a,&’-dipyridyl (164,165,233,243,244). Although this method is accurate, i t is time consuming and not well suited for enzyme purification. A variation of this method makes use of an underhydroxylated protein substrate which contains [3,4-3H]proline. On incubation of this substrate with enzyme, the tritium in the 4-trans position of certain prolyl residues is displaced by the entering hydroxyl group and is therefore released in proportion to the hydroxyproline formed (245). The tritium equilibrates with water which is then distilled and assayed for radioactivity. Since hydrolysis of the protein is not necessary and the tritiated water is easily separated from the other radioactive materials, this assay is both rapid and sensitive. The limitations of this procedure are the necessity of preparing labeled substrate and its nonuniformity which precludes comparison of enzymic activities from laboratory to laboratory. The data, by themselves, cannot be used for stoichiometric studies and yield enzymic activity in arbitrary units. An assay (241) which does not have these shortcomings is based on the conversion of a-[ 1J4CJ ketoglutarate to radioactive CO,. As discussed in Section IV,B,3 the production of CO, is stoichiometric with that of hydroxyproline. This assay is the simplest and most rapid and yields data in molar units. Although its sensitivity is reduced when other a-ketoglutarate decarboxylating enzymes are present, this usually is only a problem with crude preparations. The assay is inherently less sensitive than the assays which use tritiated substrates. Therefore, the a- [ 1-14C]ketoglutarate assay has been used to calibrate the assay for tritium release so that some stoichiometry was obtained (191).

5. Regulation

Important to an understanding of the factors which regulate prolyl hydroxylase activity is the determination of the intracellular site and the step in the biosynthesis of collagen a t which the enzyme functions. Early work indicated that a significant amount of prolyl hydroxylase 243. K. I. Kivirikko and D. J. Prockop, BJ 102, 432 (1967). 244. L. N. Lukens, JBC 245, 453 (1970). 245. J. J. Hutton, A, L. Tappel, and S. Udenfriend, Anal. Biochem. 16, 384 (1966).

162

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

was localized in the microsomal fraction (165). Recent reports suggest a vesicular localization (243,,!246,,!247) , and data based on ultracentrifugation studies of collagen synthesis in a membrane-bound polysome system from chick embryos show that prolyl hydroxylase is attached to the membranes of the rough endoplasmic reticulum (248). With ferritin-labeled antibodies in an electron microscopic study, Olsen et al. (249) have obtained results which indicate that the enzyme is localized within the cisternae rather than on the ribosomal side of the rough endoplasmic reticulum. However, Al-Adnani et al. (250) claimed that prolyl hydroxylase is attached to the membrane of the rough endoplasmic reticulum in rat fibroblasts and chondrocytes and suggested that the preparative procedures utilized by Olsen et al. (24.99)might have displaced the enzyme from the membranes into the cisternae. The electron microscopic study of AlAdnani et al. (250) involved treating fixed fibroblasts with specific goat antiserium to rat prolyl hydroxylase followed by rabbit anti-goat IgG conjugated with horseradish peroxidase and a histochemical reaction for peroxidase. To determine which of the intermediate peptidyl precursors of collagen is in vivo the substrate for prolyl hydroxylase, Miller and Udenfriend (251) carried out experiments which involved prereleasing nascent collagen chains with puromycin from ribosomes purified from guinea pig granuloma minces. These chains were shown to be covalently linked to puromycin and to contain hydroxyproline. This work, the studies with the membrane-bound polysome system previously referred to (248), and the experiments of Lazarides et al. (252), which showed that most of the proline in the polysomes isolated from 3T6 fibroblasts was not susceptible to hydroxylation by purified prolyl hydroxylase, argue for the nascent collagen peptides being the major sites of hydroxylation. Ascorbate not only can affect the rate of hydroxyproline formation by serving as a cofactor in the prolyl hydroxylase reaction but also can affect this rate in an entirely different way (253,,!2&).More insight into the latter role of ascorbate has come recently from studies with fibro246. N. A. Guzman and X. R. Cutroneo, BBRC 52, 1263 (1973). 247. N. A. Guzman, P. M. Prichard, M. M. Sharaway, and K. R. Cutroneo, Fed. Proc., Fed. Amer. Sac. E q . Biol. 33, 1535 (1974). 248. R. F. Diegelman, L. Bernstein, and B. Peterkofsky, JBC 248, 6514 (1973). 249. B. R. Olsen, R. A. Berg,Y. Kishida, and D. J. Prockop, Science 182, 826 (1973). 250. M. S. Al-Adnani, R. S. Patrick, and J. O'D. McGee, J . Cell Sci. (in press). 251. R. L. Miller and S. Udenfriend, ABB 139, 104 (1970). 252. E. L. Lazarides, L. N. Lukens, and A. A. Infante, J M B 58, 831 (1971). 253. F. L. H. Stassen, G. J. Cardinale, and S. Udenfriend, Proc. Nut. Acud. Sci. U . S. 70, 1090 (1973). 254. C. 1. Levine, J. J. Aleo, C. J. Prynne, and C. J. Bates, BBA 338, 29 (1974).

3.

163

OXYGENASES: DIOXYGENASES

blasts which have revealed an inactive form of prolyl hydroxylase (655). This inactive form of the enzyme specifically cross-reacts with antibody which was prepared by using pure prolyl hydroxylase as antigen. The cross-reacting protein is converted intracellularly to active enzyme in the presence of lactate (656) or small amounts of ascorbate (253). The reverse conversion, that of active enzyme to cross-reacting protein, is effected by incubation of the cells with dithiothreitol (255).The crossreacting protein has been purified and shown to have a molecular weight of about 85,000-100,000in comparison to the active enzyme which has a molecular weight of 260,000-300,000 (190). On the basis of the molecular weight relationships and their studies with dithiothreitol, Udenfriend and co-workers (190,653)have suggested that the cross-reacting protein may be a subunit precursor of the active enzyme. In accord with this suggestion, the active tetrameric chick enzyme was dissociated into inactive monome'rs by treatment with high concentrations of dithiothreitol (195). Whether there is any regulatory significance to this putative precursor and the mechanism for its conversion to enzyme are still to be dehermined. 6. Nonvertebrate Prolyl Hydroxylase

Although less is known about prolyl hydroxylase in plank, the chronological development of the subject has paralleled that in animals. Proline was shown t o be the precursor of peptidyl trans-4-hydroxyproline (657-262). Its hydroxyl group was shown to be derived from molecular oxygen (263) by a process which displaced the 4-trans hydrogen atom of proline (664).Peptidyl proline hydroxylation was found to be inhibited in carrot disks by a-a'-dipyridyl, and the effect was reversed with exogenous Fez+ (665). Sadava and Chrispeels (186) have obtained a cellfree system from carrot disks and purified prolyl hydroxylase about 24fold. This degree of purification permitted the requirements for Fez+,as-

s.

255. J. O'D. McGee, U. Langness, and Udenfriend, Proc. Nut. Acad. Sci. U. S. 68, 1585 (1971). 256. J. P. Comstock and S: Udenfriend, Proc. Nut. Acad. Sci. U . 8.68, 552 (1970). 257. F. C. Steward, J. F. Thompson, F. K . Miller, M. D. Thomas, and R. H. Hendricks, Plant Physiol. 26, 123 (1951). 258. J. K . Pollard and F. C. Steward, J . Exp. Bot. 10, 17 (1959). 259. F. C. Steward and L. 0.Chang, J. Exp. Bot. 14,379 (1963). 260. A. C. Olson, Plant Physiot. 39, 543 (1964). 261. R. Cleland, Plant Physiol. 43, 865 (1968). 262. M. J. Chrispeels, Plant Physiol. 44, 1187 (1969). 263. D. T. A. Lamport, JBC 238, 1438 (1963). 264. D. T. A. Lamport, Nature (London) 202, 293 (1964). 265. M. J. Chrispeels, Plant Physiol. 45,223 (1970).

164

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

corbate, and a-ketoglutarate to be demonstrated, but the latter could also be met by other a-keto acids and the stoichiometry of the reaction has not been shown. Perhaps of evolutionary significance is the finding by Sadava and Chrispeels (186) that the plant prolyl hydroxylase can hydroxylate substrate prepared from chick embryos and that the animal enzyme can hydroxylate substrate prepared from plants. The leaves of the sandal, Santalum album L., are of interest in that although they apparently make trans-4-hydroxy-~-prolinein the usual manner, they also seem to be able to hydroxylate free proline to yield cis-4-hydroxy-~-proline (266,967).The mechanism of the latter hydroxylation has not been investigated. A prolyl hydroxylase from earthworm subcuticular cells has the same cofactor requirements as the vertebrate enzymes (168,688,269). The assays based on tritium release and the production of radioactive CO, has permitted its purification 150-fold (670). Preliminary experiments indicate the molecular weight of the enzyme is about 300,000 and that its isoelectric point is below p H 5 (270). The substrate specificity of this hydroxylase differs markedly from that of the vertebrate enzymes; for example, although the latter enzymes cannot hydroxylate (Gly-L-Pro+Ala),, the earthworm enzyme can hydroxylate it at a rate 8-10 times faster than it can (Gly-L-Ala-L-Pro), (270).The earthworm hydroxylase has been reported ( 1 68) to produce 3-hydroxyproline ; however, 4-hydroxyproline is the only product detected by Rao and Adams (270). The muscle layers of Ascaris lubricoides have also yielded a prolyl hydroxylase which requires Fez+,ascorbate, a-ketoglutarate, and molecular oxygen to synthesize peptidyl hydroxyproline (186). When the hydroxylase was compared with the chick embryo enzyme in regard to its substrate specificity, the two enzymes appeared similar, but the Ascaris enzyme was not inhibited by poly-L-proline. Also, in contrast to prolyl hydroxylase in higher animals, the Ascaris enzyme is inhibited by oxygen a t normal atmospheric concentrations (186).I n addition, A. lumbricoides may contain an isozyme of prolyl hydroxylase which is not inhibited by oxygen (271-873). Streptomyces antibioticus also appears to contain a prolyl hydroxylase. Studies with intact cells have shown that the synthesis of trans-4-hydroxy266. 267. 268. 269. 270. 271. 272. 273.

R. Kuttan and A. N. Radhakrishnan, BJ 117, 1015 (1970). R. Kuttan and A. N. Radhakrishnan, BJ 119,651 (1970). A. Nordwig, V. Kobrle, and F. K . Pfab, BBA 147,487 (1967). A. Goldstein and E. Adams, JBC 245, 5478 (1970). N. V. Rao and E. Adams, Fed. Proc., Fed. Amer. Sac. Exp. Biol. 33, 1535 (1974). M. Chvapil, M. Boucek, and E. Ehrlich, ABB 140, 11 (1970). M. Chvapil and E. Ehrlich, BBA 208, 467 (1970). G. D. Cain and D. Fairbain, Comp. Biochem. Physiol. B 40, 165 (1971).

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165

prolined of peptide antibiotics is very similar to the synthesis of peptidy1 hydroxyproline in animals (187,274,275).Further description of the prolyl hydroxylase awaits the development of a cell-free system.

C. LYSYLHYDROXYLASE CH,NH, I

0, +

7%

(FH,), R-NH-CHCOR'

a-keto+ glutarate

CH,NH,

reducing agent

I

CHOH

Fez+

R-NH-

+ succinate (30)

+ COz (FH,), kH- COR'

1. Introduction 5-Hydroxylysine, like hydroxyproline, is almost unique to collagen. The hydroxyl group of hydroxylysine is the site of carbohydrate attachment and may be necessary for the secretion of collagen from the cell (see Section IV,B,2). Hydroxylysine also participates in various crosslinking reactions including Schiff base formation (139,676-d78). The hydroxylysine content of collagen is much less than that of hydroxyproline and, accordingly, lysyl hydroxylase activity of most tissues is lower than prolyl hydroxylase activity but may also correlate with changes in collagen biosynthesis in several situations (279).

2. Purification and Molecular Properties Miller (280) used DEAE-Sephadex chromatography to separate lysyl hydroxylase (EC 1.14.11.4) from prolyl hydroxylase and hence proved that they are distinct enzymes. Popenoe and Aronson (281) also separated the two activities as did Kivirikko and Prockop (282) who in the process purified lysyl hydroxylase 250-fold from chick embryos. I n the purification scheme of Kivirikko and Prockop (282),gel filtration chromatography appeared to separate two forms of the enzyme. They had apparent molecular weights of 550,000 and 200,000. The most purified preparations (282) had a specific activity of 60 units/mg when assayed with a synthetic substrate. 274. L. A . Salzman, H. Weisbach, and E. Katz, Proc. N u t . Acad. SCi. U . 8. 54, 542 (1965). 275. R. F. Diegelman, 0. Ondrejickova, and E. Katz, ABB 131, 276 (1969). 276. W. T. Butler and L. W. Cunningham, JBC 241, 3882 (1966). 277. P. H. Morgan, H. G. Jacobs, J. P. Segrest, and L. W. Cunningham, JBC 245, 5042 (1970). 278. R. G. Spiro, JBC 244, 602 (1969). 279. L. Ryhanen and K . I. Kivirikko, BBA 343, 121 (1974). 280. R. L. Miller, ABB 147, 339 (1971). 281. E. A. Popenoe and R. B. Aronson, BBA 258, 380 (1972). 282. K. I. Kivirikko and D. J. Prockop, BBA 258, 366 (1972).

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3. Catalytic Properties Van Slyke and co-workers (283,284) established that lysine serves as the precursor of hydroxylysine in a process analogous to that involved in hydroxyproline formation. This analogy was extended by findings (285,286) that suggested that hydroxyl group of hydroxylysine is derived from molecular oxygen. Prockop et al. (286) obtained a cell-free system which could hydroxylate an underhydroxylated lysine-containing peptide, and Hausmann (287) showed that this reaction required Fez+,ascorbate, and a-ketoglutarate. Kivirikko and Prockop (282) obtained apparent K , values of 0.05 mM for a-ketoglutarate, 0.001 mM for Fez+, and 0.05 mM for ascorbate and showed that maximal activity was not observed unless serum albumin, catalase, and dithiothreitol were included in the incubation mixture. Also similar to that found with prolyl hydroxylase is the increased stability of lysyl hydroxylase in the presence of glycine (281). Kivirikko et al. (288) have shown that almost any peptide containing an -X-Lys-Gly- sequence, including lysine-vasopressin, can serve as substrate. As with prolyl hydroxylase the length of the chain and residues adjacent to this sequence influence the K , and Vmnxof the substrates (288), and Stoltz et al. (289) have shown that the hydroxylysine is not always followed by a glycine residue in collagen. Furthermore, just as not all susceptible prolyl residues in collagen are hydroxylated neither are all susceptible lysyl residues (290-292),and the triple helical structure appears to be one of the critical factors limiting this hydroxylation (293).

Succinate was identified as a product of a-ketoglutarate in the lysyl hydroxylase reaction, and the release of CO, was shown to be equimolar with the formation of hydroxylysine (288). Thus, one of the assays for lysyl hydroxylase involves the measurement of released COz (288). Other 283. F. M. Sinex, D. D. Van Slyke, and D. R. Christman, JBC 234, 918 (1959). 284. E.A. Popenoe and D. D. Van Slyke, JBC 237, 3491 (1962). 285. E. A. Popenoe, R. B. Aronson and D. D. Van Slyke, Proc. N u t . Acad. Sci. U.S.55, 393 (1966). 286. D. J. Prockop, E. Weinstein, and T. Mulveny, BBRC 22, 124 (1966). 287. E. Hausmann, BBA 133,593 (1967). 288. K. I. Kivirikko, K. Shudo, S. Sakakibara, and D. J. Prockop, Biochemistry 11, 122 (1972). 289. M. Stolte, H.Furthmayr, and R. Timpl, BBA 310,461 (1973). 290. W. T. Butler, Science 161, 796 (1968). 291. W. T.Butler, BBRC 48, 1540 (1972). 292. K. I. Kivirikko, L. Ryhanen, H. Anttinen, P. Bornstein, and D. J. Prockop, Biochemk-try 12, 4966 (1973). 293. L.Ryhhen and K. I. Kivirikko, BRA 343, 129 (1974).

3.

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OXYGENASES: DIOXYGENASES

assays are also used which are comparable to those developed for prolyl hydroxylase, such as those based on the measurement of radioactive hydroxylysine (2881) and on tritium release ( 2 9 4 ) . Blumenkrantz and Prockop (696) have described an assay which involves the periodate oxidation of [ "C] hydroxylysine to give [ '*C] formaldehyde.

D. 7-BUTYROBETAINE HYDROXYLASE 02

+ (CH3)3NfCHzCH2CH~COO-+ a-ketoglutarate

-

7-Butyrobetaine reducing agent Fez+

+

(CHa)sN+CH2CHOHCH2COO- succinate Carnitine

+ COZ

(31)

1. Introduction

Carnitine, which functions in fatty acid oxidation and synthesis (d96), appears to be synthesized via the following pathway: lysine + e-N-trimethyllysine + y-butyrobetaine + carnitine (297-303). Lindstedt and co-workers prepared cell-free systems, which could catalyze the latter reaction, from rat liver (304) and from a strain of Pseudomonas obtained by enrichment culture on media which contained 7-butyrobetaine (305,SOS). 2. Purification

The y-butyrobetaine hydroxylase (EC 1.14.11.1) from rat liver was purified by ammonium sulfate fractionation and hydroxyapatite chromatography to the extent that a specific activity of 3 units/mg was obtained (307).The enzyme from Pseudomonas was purified 18-fold in 6% yield and had a specific activity of 220 units/mg (306).Since the hydroxylase from the two sources are similar in many of their properties, the discussion which follows pertains to both enzymes unless otherwise indicated. 294. R. Miller, Anal. Biochem. 45, 202 (1972). 295. N. Blumenkrrtntz and D. J. Prockop, Anal. Biochem. 30, 377 (1969). 296. I. B. Fritz, Advan. Lipid Res. 1, 285 (1963). 297. W.Linneweh, Hoppe-Seyler's Z . Physiol. Chem. 181, 42 (1929). 298. J. Bremer, BBA 48, 622 (1961). 299. J. Bremer, BBA 57, 327 (1962). 300. G.Lindstedt and S. Lindstedt, BBRC 6, 319 (1961). 301. G. Lindstedt and S. Lindstedt, JBC 240, 316 (1965). 302. V. Tanphaichitr and H. P. Broquist, JBC 248,2176 (1973). 303. T. Bplhmer, BBA 343, 551 (1974). 3 M . G. Lindstedt, BiochemZstry 6, 1271 (1967). 305. G. Lindstedt, S. Lindstedt, T. Midvedt, and M. Tofft, Biochemistry 6, 1262 (1967). 306. G. Lindstedt, S. Lindstedt, and M. Tofft, Biochemistry 9, 4336 (1970). 307. G.Lindstedt and S. Lindstedt, JBC 245, 4178 (1970).

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OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

3. Catalytic Properties As with other a-ketoglutarate dioxygenases, the requirements for Fez+ and ascorbate were discovered first (308).An NADPH generating system was also found to be stimulatory, but, in retrospect, this functioned as an a-ketoglutarate generating system (304,305,309).With partially purified enzyme preparations (306,307) the specificity of the requirement for a-ketoglutarate wm found to be high. It could not be replaced by such compounds as glutarate, oxaloacetate, 2-ketoadipate, glutamate, 2-hydroxyglutarate, and succinic semialdehyde. However, several of the analogs tested were inhibitory to the hydroxylase. Most likely a-ketoglutarate per se is in vivo the substrate for the hydroxylase since the addition of either glutamate-oxaloacetate transaminase and asparate or of glutamate dehydrogenase, ammonium chloride, and NADPH inhibited the low level of hydroxylase activity observed in the absence of exogenous a-ketoglutarate. In those studies a-ketoglutarate had an apparent K , value of about 0.5 mM. The ferrous ion requirement was also found to be highly specific. 1 , l O Phenanthroline inhibited the hydroxylase activity of partially purified enzyme preparations in the absence of added iron, and only Fez+could reverse this inhibition. Several metal ions inhibited the hydroxylase (306,310). Apparent K , values of 0.1 and 0.06 mM were obtained for Fe2+with the rat liver and Pseudomonas hydroxylase systems. Although ascorbate did have a stimulatory effect, appreciable hydroxylase activity was observed in the absence of added ascorbate, and it could be a t least partially replaced by a number of reducing agents including tetrahydrofolate. Catalase stimulated the hydroxylase reaction, but the specificity of this effect was not investigated. Catalase was also shown to prevent the loss of activity which resulted from preincubation of the hydroxylase with a mixture of Fez+and ascorbate. Mercurials and other reagents which react with sulfhydryl groups were shown to inhibit the enzyme. The hydroxylase was found to have maximal activity a t pH values near neutrality (306,307). Several analogs of 7-butyrobetaine were tested as substrates for the partially purified enzyme preparation from rat liver. Hydroxylation of 3-trimethylaminopropionic acid and of 4-dimethylaminobutyric acid did occur but at slower rates. However, 5-trimethylaminovaleric acid and quaternary amines without a carboxyl group were not hydroxylated to any extent (304).A number of substrate analogs were found to be inhibi308. G. Lindstedt and S. Lindstedt, BBRC 7, 394 (1962). 309. G. Lindstedt, S. Lindstedt, T. Midvedt, and M. Tofft, BJ 103, 19P (1967). 310. G. Lindstedt, BBA 141, 492 (1967).

3.

169

OXYGENASES : DIOXYGENASES

tory. Of these, 3-trimethylaminopropylsulfonate was the most effective (307).Bacterial y-butyrobetaine hydroxylase preparations had a higher substrate specificity. A number of analogs of 7-butyrobetaine were found to be inhibitory, but none were hydroxylated themselves (306). Lindstedt and co-workers (307,311,312) made a contribution of farreaching significance when they showed that a-ketoglutarate is degraded to succinate during the formation of carnitine and that succinic semialdehyde is most likely not an intermediate in the reaction. Succinic semialdehyde was not able to substitute for a-ketoglutarate in the hydroxylase reaction and the formation of the aldehyde was not detected in incubation mixtures containing a-[5J4C]ketoglutarate and nonradioactive succinic semialdehyde which had been added as a trapping agent. I n the studies of stoichiometry, the disappearance of a-ketoglutarate was measured enzymically with a spectrophotometric assay using glutamate dehydrogenase, ammonium chloride, and NADH. A 1: 1 molar correlation between the utilization of a-ketoglutarate and the appearance of carnitine was demonstrated (312). A 1 : 1 molar correlation between the formation of CO, and carnitine was also established with the use of a-[ 1-I4C]ketoglutarate (306,307). The demonstration that the 7-butyrobetaine hydroxylase reaction was inhibited under anerobic conditions suggested that the hydroxyl group of carnitine is derived from molecular oxygen (304,305). That an intermediate peroxy compound is formed in the y-butyrobetaine hydroxylase reaction gained strong support. when Lindblad et al. (313) showed that one atom of molecular oxygen was incorporated into succinate. I n that study the hydroxylase incubation mixture was incubated in the presence of oxygen gas enriched with I8Q. The succinate which was formed was converted to its bis (trimethylsilyl)ester and sub,iected to mass spectrometry. As shown in Fig. 3 about 75% of the succinate molecules were found to contain one atom of 1 8 0 . The l8O content of carnitine was not determined in that study.

E.

DIOXYGENASE

REACTIONS OF

PYRIMIDINES AND NUCLEOSIDES

3. Introduction Fink and Fink (314,315)originally proposed that in Neurospora crassa the pyrimidine ring of thymidine might be salvaged by a pathway which 311. E. Holme, G.Lindstedt, S. Lindstedt, and M. Tofft, FEBS (Fed. Eur. Biochem. Sac.) Lett. 2, 29 (1968). 312. G. Lindstedt, S. Lindstedt, B. Olander, and M. Tofft, BBA 158, 503 (1968). 313. B.Lindblad, G.Lindstedt, M. Tofft, and S. Lindstedt, JAGS 91, 4604 (1969). 314. R.M.Fink and K. Fink, Fed. Proc., Fed. Amer. Soc. Ezp. BioE.21,377 (1962). 315. R. M.Fink and K. Fink, JBC 237, 2889 (1962).

170

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

m /e

FIG.3. Mass spectrum of the bis(trimethylsily1)ester of succinic acid formed during the incubation of y-butyrobetaine hydroxylase in 90% "0,(lower graph) and of bis(trimethylsi1y) ester of authentic succinic acid (upper graph). M is the molecular ion and M-15 the positive ion after loss of one methyl radical from trimethylsilyl group of the ester. The incorporation of one atom of "0 in about 75% of the formed succinic acid is evident from the presence of the corresponding ions plus two mass units in the lower graph. From Lindblad et al. (313).

involves oxidative demethylation. They demonstrated that thymidine was incorporated into the pyrimidines of RNA in a process that does not involve fragmentation of the pyrimidine ring and detected the formation of 5-hydroxymethyluraci1 and 5-formyluracil. The reactions depicted in Fig. 4 have been shown to be catalyzed by purified enzyme preparations obtained from Neurospora (316-3g2). Further evidence that these reactions occur in vivo have come from studies by Williams and Mitchell (323) who have developed pyrimidineless mutants which can either utilize or accumulate some of the intermediates and from additional studies 316. P. M. Shaffer, R. P. McCroskey, R. D. Palmatier, R. J. Midgett, and M. T. Abbott, BBRC 33, 806 (1968). 317. P. M. Shaffer, R. P. McCroskey, and M. T. Abbott, BBA 258, 387 (1972). 318. M. T. Abbott, R. J. Kadner, and R. M. Fink, JBC 239, 156 (1964). 319. M . T. Abbott, T. A. Dragila, and R. P. McCroskey, BBA 169, 1 (1968). 320. M. S. Watanabe, R. P. McCroskey, and M. T. Abbott, JBC 245, 2023 (1970). 321. R. D. Palmatier, R. P. McCroskey, and M. T. Abbott, JBC 245,6706 (1970). 322. P. M. Shaffer, and M. T. Abbott, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 30, 1222 (1971). 323. L. G. Williams and H. K. Mitchell, J. Bacteriol. 100, 383 (1969).

3.

171

OXYGENASES: DIOXYGENASW

a H

OH

OH

OH

Thymine ribonucleoside

Thymidine

DNA RNA

I ,II

H

N

F

oAN H

9 fb- ,’ 5- Hydroxymethyluracil

Deoxycytidine

h H

Uracil-5- carboxylic acid

OH Deoxy uridine

OH

OH

Uridine

FIG.4. Pathway proposed for the utiliiation of deoxyribonucleosidesby Neurospora crassa: a and h, pyrimidine deoxyribonucleoside 2’-hydroxylase (516,517); b, hydrolase reaction (317) ; c, d, and e, thymine 7-hydroxylase (518-320); f, uracil-5-carboxylic acid decarboxyhe (321); and g, deaminase reaction (317). From Shaffer et al. (524).

(394) with intact cells of these mutants. That thymine is demethylated a t the free base level, as shown in Fig. 4, is strongly suggested by the inability of thymidine and thymine ribonucleoside (317)to 8erve as substrates for thymine 7-hydroxylase since reactions c, d, and e (Fig. 4) all appear to be catalyzed by this enzyme (326).Although the discovery of the 2’-hydroxylase was totally unexpected and there is presumptive evidence for it not being peculiar to Neurospora (1.4~5)~ the presence of this enzyme in Neurospora can be rationalized from a teleological point of view. Since Neurospora lacks a thymidine phosphorylating enzyme 324. P. M. Shaffer, C. A. Hsu, and M. T. Abbott, J . Bacteriol. 121, 648 (1975). 325. C. K. Liu, C. A. Hsu, and M. T. Abbott, ABB 159, 180 (1973).

172

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

PURIFICATION

Step

Fraction

TABLE VII THYMINE 7-HYDROXYLASE’

OF

VolAcume Protein tivity (ml) (m@;/ml) (U)

Specific activity (U/mg)

Yield (%)

Purification (fold)

Extract 2200 9 . 1 15,400 0 . 7 7 100 Calcium phosphate I 2400 2 . 9 12,500 1.8 81 2.3 Calcium phosphate I1 1200 3 . 0 9,840 2.7 64 3.5 Ammonium sulfate 130 2 7 . 0 10,000 2.9 65 3.6 Sephadex G-150 800 1 . 2 7,280 7.6 47 9.9 DEAE-cellulose I 150 0.35 54 19 70 2,850 Sephadex G-100 25 0 . 7 1 90 10 116 1,580 DEAE-cellulose I1 33 0 . 1 3 1,260 290 8 . 2 380 Preparative disc 21 0.0048 2.0b 1370 303b 1060 electrophoresis From Liu et al. (3.86).

* Value adjusted to take into account that only one-third of the DEAE-cellulose I1 fraction waa subjected to step 9.

(316,326),the pathway shown in Fig. 4 provides a means of salvaging the pyrimidine ring of thymidine, but a source of ribose may still be required in order for the uracil end product to be converted to the nucleotide precursors of RNA and DNA. Thus, the 2’-hydroxylase reaction allows the otherwise discarded deoxyribose moiety also to be salvaged (324). Interestingly, the 2’-hydroxylase reaction is catalyzed by an aketoglutarate dioxygenase as are the other three oxidative steps of the pathway.

2. Purification and Molecular Properties

The enzymes involved in the oxidative demethylation of thymidine were purified and separated from each other when extracts from Neurospora were subjected to the scheme shown in Table VII. Uracil-5-carboxylic decarboxylase was removed from the rest of the enzymes of the pathway (32) by the calcium phosphate gel fractionation procedure (steps 2 and 3, Table VII). Separation of the other three enzymes of the pathway (327,328) was achieved with DEAE-cellulose chromatography (step 6, Table VII). Experiments in which the fractions with the 2’-hydroxylase and 7-hydroxylase activities were recombined in various propor326. A. R. Grivell and J. F. Jackson, J . Gen. Microbiol. 54, 307 (1968). 327. C. K. Liu, P. M. Shaffer, R. S.Slaughter, R. P. McCroskey, and M. T. Abbott, Fed. Proc., Fed. Amer. Soc Exp. Biol. 31,882 (1972). 328. C. K. Liu, P. M. Shaffer, R. S. Slaughter, R. P. McCroskey, and M. T. Abbott, Biochemistry 11,2172 (1972).

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OXYGENASES: DIOXYGENASES

173

tions (327), the difference in stability of the two activities (Sag), and the finding (324) that one of the pyrimidineless mutants developed by Williams and Mitchell (323) appears to lack only the 7-hydroxylase and that another of the mutants only the 2’-hydroxylase supported the conclusion that the 2’-hydroxylase and the 7-hydroxylase are distinct enzymes. The enzyme preparation with 2’-hydroxylase activity obtained from the DEAE-cellulose column (step 6, Table VII) had a specific activity of 50 units/mg with respect to the substrate thymidine and also catalyzed the 2’-hydroxylation of deoxyuridine. These two deoxyribonucleosides seem to be hydroxylated by a single enzyme since the mutant, mentioned above, which appeared to lack only the 2’-hydroxylase when assayed with thymidine as substrate, yielded comparable results when assayed with deoxyuridine (32.4). Furthermore, although the 2‘-hydroxylase has been only partially purified ( 3 2 4 ) ,the ratio of the specific activities, obtained with each substrate, remained constant throughout the purification, and preliminary kinetic experiments showed thymidine and deoxyuridine to be competitive inhibitors of each other (324). Further purification of the 7-hydroxylase yielded a preparation with a specific activity of 1200 units/mg (Table VI). This preparation also catalyzed the conversions of 5-hydroxymethyluracil to 5-formyluracil and of the latter to uracil-5-carboxylic acid and had specific activities, with respect to these reactions, of 600 and 250 units/mg (326‘).Several observations (325) are consistent with a single protein catalyzing reactions c, d, and e (Fig. 4 ) . These include the parallel purification of the three activities throughout the purification scheme, the inhibition of each reaction by the substrates of the other two, the inhibition of the three reactions by uracil, the parallel loss of the three activities upon heat denaturation, and considerations of mechanism which suggest that a single active site may be involved (see Section IV,G). The catalysis of the conversion of 5-formyluracil to uracil-5-carboxylic acid by thymine 7-hydroxylase appeared not to be in accord with the nutritional requirements of a pyrimidineless mutant of Neurospora which can utilize 5-formyluracil but not thymine or 5-hydroxymethyluracil (323). However, the apparent conflict was resolved by the discovery of another enzyme in Neurospora which can catalyze this conversion (324). This enzyme, as well as one found in rat liver (330),does not appear to have an a-ketoglutarate dependency (324) and may be a relatively nonspecific dehydrogenase. 329. R. P. McCroskey, W. R. Griswold, R. L. Sokoloff, E. Sevier, S. Lin, C. K. Liu, P. M. Shaffer, R. D. Palmatier, T. S. Parker, and M. T. Abbott, BBA 227, 264 (1971). 330. R. G. Finley and M. T. Abbott, unpublished.

174

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

Lindstedt and co-workers (331,332) have used DEAE-Sephadex chromatography to separate the 2’-hydroxylase from the 7-hydroxylase. The molecular weight of the 2’-hydroxylase was determined, by means of gel filtration, to be 47,000. The data of this determination also indicate that the molecular weight of the 7-hydroxylase is somewhat lower than that of the 2’-hydroxylase (332). Isoelectric focusing was used to determine that the isoelectric point of the 2‘-hydroxylase is 4.6 and that of the 7hydroxylase is 4.9 (332). 3. Catalytic Properties of the Thymine 7-Hydroxylase (EC 1.14.11.6)

a. Thymine Dioxygenase Reaction (c, Fig. 4).An oxygenase was proposed as catalyst for the conversion of thymine to 5-hydroxymethyluracil a t the time of development of the first cell-free system since it was stimulated by molecular oxygen (318,333). The requirements for Fez+, ascorbate, and a-ketoglutarate were demonstrated with crude extracts which had been subjected to Sephadex G-50 chromatography (334). The specificity of the a-ketoglutarate requirement appeared high even when studied (334) with the relatively impure calcium phosphate gel fraction (step 3, Table VII) . Little or no activity was observed when the following compounds were substituted for a-ketoglutarate: glutarate, levulinate, a-ketovalerate, pyruvate, glyoxylate, oxaloacetate, isocitrate, glutamate, citrate, succinate, fumarate, NADPH, and NADH (145). Holme et al. (335) determined that a-ketcglutarate had an apparent K, value of 0.2 mM. The Fez+requirement could be met by no other metal ion tested, and several metal ions were shown to be inhibitory (145). Appreciable hydroxylase activity was observed in the absence of added Fe2+,but this activity was eliminated by preincubation of the enzyme preparation with EDTA and was fully restored upon dialysis followed by incubation in the presence of Fe2+.The reducing agent requirement appeared to be best met by ascorbate or isoascorbate, but either reduced glutathione or 2-mercaptoethanol was also effective (145). In addition, catalase was stimulatory (336),but the specificity of the effect was not studied. The pH optimum of the hydroxylase reaction was determined to be about 7.0 (329). Thymine 7-hydroxylase does not appear to be an enzyme of low substrate specificity since no hydroxylation of the methyl groups of thymi331. L. Bankel, E. Holme, G. Lindstedt, and S.Lindstedt, FEBS (Fed. Eur. Biochem. Soc.) Lett. 21, 135 (1972). 332. L. Bankel, G. Lindstedt, and S. Lindstedt, JBC 247. GI28 (1972). 333. M. T. Abbott, in “Methods in Enzymology” Vol. 12, Part A, p. 47, 1967. 334. M. T. Abbott, E. K. Schandl, R. F. Lee, T. S. Parker, and R. J. Midgett, BBA 132, 525 (1967). 335. E. Holme, G. Lindstedt, S. Lindstedt, and M. Tofft, BBA 212, 50 (1970).

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OXYGENASES : DIOXYGENASES

175

dine, thymine ribonucleoside (317), or 5-methylcytosine (145) was detected when these compounds were tested as substrates. Each of the three reactions of thymine 7-hydroxylase is inhibited by the substrates of the other two reactions and by uracil (325). The conversion of thymine to 5-hydroxymethyluracil was shown to be ketoglutarate to 14C02 coupled both to the conversion of a-[ l-*"C] (329,335) and of a[5-14C] ketoglutarate to ['*C]succinate (329). Thus the time-consuming assays based on chromatographic techniques could now often be circumvented. The crucial 1 8 0 2 studies were carried out by Holme et al. (336). The lSO content of the products was measured by means of mass spectrometry after gas chromatographic separation of the trimethylsilyl derivatives of 5-hydroxymethyluracil and succinate. One atom of molecular oxygen was shown to be incorporated into 5-hydroxymethyluracil and another into succinate. b. 6-HydroxymethyZuraci1 and 5-Formyluracil Dioxygenase Reactions (d and el Fig. 4). When enzyme preparations were obtained which were more effective in catalyzing the conversion of thymine to 5-hydroxymethylracil, the latter's conversion 5-formyluracil was detected (319). Unexpectedly, cofactors which are usually required by dehydrogenases did not stimulate the conversion of 5-hydroxymethyluracil to 5-formyluracil, but the combination of Fez+,ascorbate, and a-ketoglutarate did (319).Similarly, the conversion of 5-formyluracil to uracil-5-carboxylic acid was stimulated by the same combination (320), and the description in the balance of this paragraph pertains to both of these dioxygenase reactions. Even though no activity was detected in incubations carried out with the calcium phosphate gel fraction (step 3, Table VII) in the absence of Fez+,ascorbate, and a-ketoglutarate, some activity was obtained when one of the three was omitted. However, the requirement for the omitted component could not be satisfied by an increase in the concentrations of the other two (319,390).As with the thymine dioxygenase reaction, the specificity of the requirements for a-ketoglutarate (319,520,336) and Fez+ (319,320) appeared very high. The ascorbate requirement could be a t least partially met by such compounds as glutathione and dithiothreito1 (319,320). I n the studies of the stoichiometry of the 5-hydroxymethyluracil dioxygenase reaction, the further oxidation of the 5-formyluracil product by the 5-formyluracil dioxygenase reaction did riot have to be contended with since little or no uracild-carboxylic acid was formed when the incubation period was not prolonged. It is pertinent in this regard that the 5-formyluracil dioxygenase reaction is inhibited by 5-hydroxymethyluracil (323) and has a lower specific activity (see Section IV,E,P). Liu 336. E. Holme, G. Lindstedt, S. Lindstedt, and M. Tofft, JBC 246, 3314 (1971).

176

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

et al. (328) showed that when 5- [7-"C] hydroxymethyluracil and a-[ 1J4C]ketoglutarate were used as substrates and the time of incubation was varied, CO, and 5-formyluracil were produced in a 1: 1 molar ratio. When unlabeled 5-hydroxymethyluracil and a- [ 1,5-14C]ketoglutarate were used as substrates, CO, and succinate were produced in a 1: 1 molar ratio. Similarly, when 5-formyluracil was used as substrate, uracil-5-carboxylic acid, succinate, and CO, were produced in a 1:1: 1 molar ratio (328). That molecular oxygen is incorporated into the formyl group of 5-formyluracil in the 5-hydroxymethyluracil dioxygenase reaction and into the carboxyi group of uracil-5-carboxylic acid in the 5-formyluracil dioxygenase reaction was suggested by studies which showed that these reactions were inhibited by exclusion of oxygen from the reaction mixtures (319,320).I n experiments with ISO,, one atom of molecular oxygen was shown to be incorporated into uracil-5-carboxylic acid in its formation from 5-formyluracil (336).A similar attempt to show the incorporation of molecular oxygen into 5-formyluracil in its formation from 5-hydroxymethyluracil has not been made because of the anticipated rapid exchange of the oxygen of the formyl group with that of water. However, one atom of molecular oxygen has been shown to be incorporated into succinate in each of these reactions (336). 4. Catalytic Properties of Pyrimidine Deoxyribonucleoside 9'-Hydroxylase (EC l .l.4.ll.S)

In a study of substrate specificity, impure preparations of thymine 7-hydroxylase were incubated with thymidine, and although its methyl group was not hydroxylated thymine ribonucleoside was formed. It did not appear that this compound was formed by the ribosylation of thymine or the methylation of uracil a t the polynucleotide level since the ribonucleoside was still formed in enzyme preparations which had been subjected to some purification. Further evidence that the conversion of thymidine to thymine ribonucleoside occurred without detachment of the deoxyribose was obtained by incubation of the calcium phosphate gel fraction with thymidine which was uniformly labeled with 14C and enriched with 3H in position 6 of the pyrimidine ring. Determination of the specific radioactivities, with respect to the two isotopes, revealed that the ratios were the same in substrate and product and unchanged even when nonradioactive, potential intermediates, such as ribose 1-phosphate, were included in the incubation mixture (316).The conversion of deoxyuridine to uridine has also been shown to occur at the nucleoside level (317,322). Thus, pyrimidine deoxyribonucleoside 2'-hydroxylase catalyzes the hydroxylation of either thymidine or deoxyuridine in the only known direct transformation of deoxyribose to ribose. This reaction is in part

3.

177

OXYGENASES : DIOXYGENASES

the reverse of the ribonucleotide reductase reactions in which the 2’-hydroxyl group is reduced by dithiols. These reductases convert purine and pyrimidine ribonucleotide diphosphates and triphosphates to the corresponding deoxyribonucleotide phosphates in apparently irreversible reactions (337,%l8). The pyrimidine deoxyribonucleoside 2‘-hydroxylase reaction was shown to be dependent on the presence of a-ketoglutarate, Fez+,and ascorbate in the incubation mixture (316).A number of analogs of a-ketoglutarate were tested but none was able to replace it (332). Some of the analogs, e.g., 3-ketoadipate and 2-ketoadipate1 were moderately inhibitory to the reaction when they were included in the incubation vessel a t a concentration ten times that of a-ketoglutarate. An apparent K , value of about 0.2 mM was obtained for a-ketoglutarate. The metal ion requirement was shown to be highly specific for Fez+which had an apparent K , value of 0.45 mM (332). The specificity of the ascorbate requirement has not been studied. Catalase was found to stimulate the 2’-hydroxylation of thymidine and deoxyuridine although the specificity of the effect was not examined. The maximum stimulation, obtained by varying the concentration of catalase in the incubation mixture, was 3-fold (328). Shaffer e t al. (317,322) tested various radioactive compounds as substrates for the 2’-hydroxylase reaction. These were incubated in the presence and absence of the nonradioactive compounds which were the anticipated products and used as trapping agents. Since deoxycytidine is incorporated into RNA to about the same extent as is thymidine (315),it was suspected that deoxycytidine would also be hydroxylated. However, the calcium phosphate gel fraction did not catalyze the hydroxylation of deoxycytidine but instead its deamination to deoxyuridine, which then underwent 2‘-hydroxylation. Deoxyadenosine and deoxyguanosine were also not hydroxylated by these preparations. Therefore, thymidine and deoxyuridine are the only commonly occurring deoxyribonucleosides which are substrates for the 2’-hydroxylase. Since neither deoxyuridylate nor deoxyribose was hydroxylated, the enzymic 2’-hydroxylation reaction appears to be specific for substrates at the nucleoside level (317,322). Bankel et al. (332) confirmed these findings by testing the capacity of potential substrates to stimulate CO, production in the 2’-hydroxylation reaction carried out in the presence of [ 1-14C]ketoglutarate. Thymidine and deoxyuridine had apparent K , values of 0.09 and 0.19 mM and Vmax values of 14 and 35 units/g, respectively. Thymidylate appeared to be slightly active with a V,,, one-thirtieth that of thymidine. In addition, (Y-

337. P. Reichard, “The Biosynthesis of Deoxyribose,” Ciba Lect. Wiley, New York, 1967. 338. H. P. C. Hogenkamp, Annu. Rev. Biochern. 37, 233 (1968).

178

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

the deoxyribose derivatives of 5-hydroxymethyluracil, 5-bromouracil, and 6-azathymine were found to be active substrates. However, substrate specificity data that are solely based on the assay for CO, should be interpreted with some caution since by this criterion a compound which cannot serve as a substrate but which is contaminated with one, would appear to be active. Moreover, it may be possible to uncouple the decarboxylation of a-ketoglutarate from the hydroxylation step as has been done with the prolyl hydroxylase reaction (see Section IV,G). The production of CO, was not detected when deoxycytidine, deoxyadenosine, deoxyguanosine, deoxyinosine, and deoxyribose were used as substrates (332). Liu et al. (328) showed in studies with a-[1-14C]ketoglutarate that CO, and thymine ribonucleoside are produced in a 1:l molar ratio throughout the incubation period. I n similar experiments with a-[ 1,514C]ketoglutarate, CO, and succinate were shown to be produced in a 1:1 molar ratio. When deoxyuridine was used as substrate, uridine, succinate, and CO, were also shown to be produced in a 1:1:1 molar ratio (328).Moreover, in studies in which the concentration of either a-ketoglutarate or Fez+ was made rate limiting, stochiometric amounts of thymine ribonucleoside and CO, were produced (332). The formation of thymine ribonucleoside in the 2'-hydroxylase reaction was markedly inhibited when the air of the incubation mixture was replaced with nitrogen. When the nitrogen gas was subsequently replaced with air or pure oxygen, full activity was restored (3 1 7 ) .These findings have not been extended with studies using lSO,. 5. Regulation

The amount of thymine 7-hydroxylase contained by Neurospora is affected by several factors Which do not have a comparable effect on the 2'-hydroxylase. The 7-hydroxylase and 2'-hydroxylase being under different regulatory influences is consistent with the 2'-hydroxylase having the role of providing ribose for the utilization of purines and other intermediary metabolites in addition to the role, shared with the 7-hydroxylase1 of salvaging pyrimidines (339). The uc-1 mutation developed by Williams and Mitchell (323) appeared, primarily on the basis of nutritional studies, to affect a regulatory gene which controls the amount or activity of one or more enzymes involved in the oxidative demethylation of thymidine. Griswold et al. (339) have recently shown that this mutation effects a dramatic increase in the amount of thymine 7-hydroxylase contained in Neurospora. It has also been shown that the thymine 7-hydroxylase ac339. W. R. Griswold, V. 0. Madrid, P. M. Shaffer, D. C. Tappen, and M. T. Abbott, in preparation.

3.

179

OXYGENASES : DIOXYGENASES

tivity of mycelia can be markedly increased by growth of the mold in a media lacking ammonia. This might be expected for an enzyme that salvages nitrogen-containing compounds (339). Other factors that influence thymine 7-hydroxylase activity are the growth temperature and the amount of carbon dioxide in the atmosphere in which Neurospora is cultured (340). When grown under forced aeration, the mycelia usually contain a very low amount of the 7-hydroxylase (329). On the basis of these studies, nonaerated conditions of growth have been developed which permit Neurospora to be grown in large batches that are a good source of thymine 7-hydroxylase (325,329).

F. ~HYDROXYPHENYLPYRUVATE HYDROXYLASE (EC 1.13.11.27)

0 ;;' + OH

OH

reducing

0, f

y*

=

?& c=o

OH

f

co2

(32)

COO-

I

COO-

1. Introduction

Although evidence for the conversion of tyrosine to homogentisate was obtained a t the end of the last century (341) it was not until the middle of this one that the enzymic conversion of p-hydroxyphenylpyruvate to homogentisate was shown conclusively. The migration of the side chain to the ortho position was proved by determining the isotopic labeling pattern of a metabolite, acetoacetate, formed from radioactive phenylalanine and tyrosine in experiments carried out in vivo and with liver slices (342,343). The direct conversion of p-hydroxyphenylpyruvate to homogentisate was shown to be catalyzed by enzyme fractions from beef, pig, and dog liver (344-346). Other animals from which the oxygenase has been a t least partially purified include the guinea pig ( 3 4 7 ) ) rat 340. W. R . Griswold, V. 0. Madrid, P. M. Shaffer, D. C. Tappen, and M. T. Abbott, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 34, 568 (1975). 341. M. Wolkow and E. Baumann, Hopps-Seyler's Z. Physiol. Chem. 15, 228 (1891). 342. S. Weinhouse and R. H. Millington, JBC 175, 995 (1948). 343. B. Schepartz and S. Gurin, JBC 180, 663 (1949). 344. S.W. Edwards, D. Y. Y. Hsia, and W. E. Knox, Fed. PTOC.,Fed. Amer. SOC. E z p . Biol. 22, 232 (1963). 315. B. N. LaDu and V. G. Zannoni, JBC 217, 777 (1955). 346. S.E. Hager, R. I. Gregerman, and W. E. Knox, JBC 225, 935 (1957).

180

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

(347-349),rabbit (348), frog (350), man (352), chicken (359),monkey, opossum, and salmon (349). No enzymic activity has been observed in any tissue other than liver and kidney. 2. Purification and Molecular Properties

The hydroxylase from pig liver (348) was purified 65-fold. That from frog liver appears to be present in an inactive form which can be activated by treatment with trypsin or by autolysis. After activation with trypsin the enzyme was purified 80-fold, and its molecular weight was determined to be 85,000 by gel filtration (350). The hydroxylase from human liver has been purified extensively and has a molecular weight of 90,00&100,000 as determined by sedimentation equilibrium ultracentrifugation. The isoelectric point was found to be near pH 7 using isoelectric focusing in a sucrose density gradient (351). A preparation of the hydroxylase which has been obtained from chicken liver appears to be homogeneous on the basis of disc gel electrophoresis carried out a t several pH values and in the presence of sodium dodecyl sulfate (359). Recent results indicate the enzyme from rabbit liver, with a molecular weight of about 150,000, is composed of four subunits of 40,000 molecular weight which are inactive (365).The data suggest that Fez+is necessary for maintenance of the tetrameric form of the enzyme and its full activity.

3. Catalytic Properties The p-hydroxyphenylpyruvate hydroxylase reaction w'as shown to be inhibited by high concentrations of p-hydroxyphenylpyruvate and 0, (346,346,349).This inhibition by substrate can be reversed by a variety of reducing agents such as ascorbic acid, 2,6-dichlorophenolindophenol, tetrahydrofolic acid, and coenzyme Q,, (345,346,354-357). The oxygeM. N. D. Goswami, BBA 85, 390 (1964). K. Taniguchi, T. Kappe, and M. D. Armstrong, JBC 239, 3389 (1964). J. H. Fellman, T. S. Fujita, and E. S. Roth, BBA 284,90 (1972). T. Laskowska-Klita, Actu Biochim. Pol. 16, 35 (1969). B. Lindblad, S. Lindstedt, B. Olander, and M. Omfeldt, Aclu Chem. Scund. 25, 329 (1971). 352. J. H. Fellman, T. S. Fujita, and E. S. Roth, BBA 268,601 (1972). 353. T. Laskowska-Klita and I. Mochnacka, Acta Biochim. Pol. 20, 259 (1973). 354. W. E. Knox and M. Le May-Knox, BJ 49,686 (1951). 355. V. G. Zannoni, JBC 237, 1172 (1962). 356. V. G. Zannoni, G. A. Jacoby, and B. N. LaDu, BBRC 7, 220 (1962). 357. V. G. Zannoni, N. C. Brown, and B. N. LaDu, Fed. Proc., Fed. Amer. SOC. E x p . Biol. 22, 232 (1963). 347. 348. 349. 350. 351.

3.

OXYGENASES : DIOXYGENASES

181

nase was shown to be similarly inactivated and reactivated in vivo (356,358-361). Catalase also stimulated the hydroxylase in the presence of large amounts of substrate (349,362).The kinetics of the inhibition were compatible with an inhibitory product being formed during the course of the reaction (363).p-Hydroxyphenylpyruvate and oxygen have been shown to generate hydrogen peroxide which may account for the inhibition (349).Moreover, the enzyme was inactivated by storage, purification, or treatment with oxidizing agents ; this inactivation was also reversed by reducing agents (347').A metal ion was implicated in studies in which the hydroxylase was inhibited by 1,lO-phenanthroline and other chelators but not by the relatively Cu+-specific neocuproine. Of the large number of metal ions tested only Fez+restored activity to enzyme which had been inactivated with 1,lO-phenanthroline. This restoration of activity with Fez+required a reducing agent. When reactivation was carried out with Fez+and ascorbate, reagents which react w'ith sulfhydryl groups, such as N-ethylmaleimide, did not inhibit the process. Thus, the oxidation and reduction of iron was proposed as the cause of the reversible inactivation (347). This has recently gained support from the studies (referred t.0 in Section IV,F,P) which indicated Fez+ affects the subunit structure of the hydroxylase (353). With the frog liver enzyme, p-hydroxyphenylpyruvate has an apparent K , value of 0.5 m M (350), which is higher than the values obtained using enzyme preparations from dog liver (0.04 mM) (363) and r a t liver (0.02-0.05 mM) (349,364). Other compounds reported to serve as substrates for p-hydroxyphenylpyruvate hydroxylase include phenylpyruvate ($48,352),fluorophenylpyruvate (348), and 3,4-dihydroxyphenylpyruvate (352). Neither 2,5-dihydroxyphenylpyruvic acid nor any other potential intermediate was detected in the p-hydroxyphenylpyruvate hydroxylase reaction (344,365). The stochiometry of the p-hydroxyphenylpyruvate reaction was demonstrated to be t,hat indicated in Eq. (32) (344-346). Studies with lSO2 were carried out which suggested that one oxygen atom of homogentisate was derived from molecular oxygen and two atoms from 358. 359. 360. 361. 362.

W. E. Knox and M. N. D. Goswami, JBC 235, 2662 (1960). V. G. Zannoni and B. N. LaDu, JBC 235,165 (1960). V. G. Zannoni and B. N. LaDu, JBC 235,2667 (1960). M. N. D. Goswami and W. E. Knox, BBA 50, 35 (1961). V. G. Zannoni and B. N. LaDu, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 15,

391 (1956). 363. V. G. Zannoni and B. N. LaDu, JBC 234,2925 (1959). 364. E. C. C. Lin, B. M. Pitt, M. Civen, and W. E. Knox, JBC 233,668 (1958). 365. B. N. LaDu and V. G. Zannoni, JBC 219, 273 (1956).

182

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

water, and thus the reaction seemed to be catalyzed by a monooxygenase (366‘).It appeared (36‘6,367)that the atom which was derived from molecular oxygen was incorporated into the new hydroxyl group, whereas the atoms derived from w,ater were incorporated into the carboxyl group of homogentisate. Subsequently, however, Goodwin and Witkop (368,369) proposed that both atoms of molecular oxygen are incorporated in a mechanism which involves a peroxide bridge being formed between the benzene ring and the side chain. Direct evidence for the formation of such a cyclic peroxide came from the laboratory of Lindstedt and coworkers (370) in studies in which the p-hydroxyphenylpyruvate system was incubated with 1802 or H,180. Incubation of the hydroxylase system with lSOz was shown to result in the incorporation of one atom of l*O into the carboxyl group of homogentisate, but only 30% of the homogentisate molecules were found to contain lSO in their hydroxyl groups. Thus it appeared that the molecular oxygen which was incorporated into the new hydroxyl group subsequently exchanged with water. In support of such an exchange was the demonstration that upon incubation of H,180 with the hydroxylase system, lSO was incorporated into the hydroxyl groups of 70% of the homogentisate molecules. This exchange may occur in a quinoid structure since an exchange of quinone oxygens with water has been reported (371,372!).The experiment in which H,l8O was incubated with the hydroxylase system also showed that one atom of oxygen derived from water was incorporated into the carboxyl group of homogentisate. This incorporation can be explained by the exchange of the ketonic oxygen of p-hydroxyphenylpyruvate with water since after the hydroxylase system was incubated with H,lsO, 82% of the residual p-hydroxyphenylpyruvate molecules were found to contain one atom of ISO. Incubation of homogentisate with H,180 in the complete system (minus p-hydroxyphenylpyruvate) was shown to result in l80being incorporated into less than 0.5% of the homogentisate molecules. In most of the earlier studies with the hydroxylase, the enzymic assays involved determination of the loss of substrate. One of these, which has recently been modified (373), is a microassay and quite sensitive. It is 366. K. Yasunobu, T.Tanaka, W. E. Knox, and H. S. Mason, Fed. Proc., Fed. Amer. SOC.Ezp. B i d . 17, 340 (1958). 367. D. I. Crandall, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds), Vol. 1, p. 275. Wiley, New York, 1964. 368. S. Goodwin and B. Witkop, JACS 79, 179 (1956). 369. J. W. Daly and B. Witkop, Angew. Chem., Int. Ed. Engl. 2, 421 (1963). 370. B.Lindblad, G.Lindstedt, and S. Lindstedt, JACS 92, 7446 (1970). 371. H.Dahm and J.-D. Aubort, Helv. Chim. Acta 51, 1348 (1968). 372. H.Dahm and J.-D. Aubort, Helv. Chim. Acta 51, 1537 (1968). 373. D. T.Whelan and V. G. Zannoni, Biochem. Med. 9,19 (1974).

3.

183

OXYGENASES : DIOXYGENASES

based on the absorption of the enol-borate complex of p-hydroxyphenylpyruvate a t 308 nm (346). A more sensitive assay involved the use of substrate which is uniformly labeled with I4C and thin layer chromatography to separate the radioactive homogentisate (374). As with the a-ketoglutarate dioxygenases, a CO, assay has been developed for this hydroxylase, too, in which radioactive CO, formed from carboxyl-labeled p-hydroxyphenylpyruvate is determined (2749,351,374,375).This assay is, of course, more suitable for enzyme purification. A spectrophotometric assay for homogentisate has also been developed. I n this method, homogentisate is condensed with cysteine to yield a 1,4-thiazine with an absorption maximum a t 390 nm (349).

G. MECHANISM Lindstedt and co-workers (311) have proposed that a-ketoglutarate participates in the dioxygenase reaction as shown in Fig. 5. In this m.echanism the peroxide anion of the substrate makes a nucleophilic attack on the carbonyl carbon of a-ketoglutarate so that a peroxide bridge is formed between the two compounds. The proposal was made on the basis that the decarboxylation of a-ketoglutarate was stoichiometric with the hydroxylation of 7-butyrobetaine and that succinic semialdehyde did not appear to be a product in the y-butrobetaine hydroxylase reaction. This R3

\u+ u-

71

[Fe02I2+

H-CI R2

91

n-c-o-a-

&+

I

c-a caa-

/

I

R2

lI

~ _ _ _

CH3 I RI CH2-+N-CH3

I

R2 CHZ-COO-

~~

R3 = CHr-CH2-COO-

CH3

FIG. 5 . Mechanism proposed for y-butyrobetaine hydroxylase reaction. From Holme et al. (311). 374. B. Lindblad, Clin. Chim. Acta 34, 113 (1971). 376. M. C. Raheja, I. Dylewski, and C. J. Crawhall, Can. J . Biochem. 51,172 (1973).

184

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

coupling of decarboxylation to hydroxylation was independently shown by Rhoads and Udenfriend (241) in their studies with prolyl hydroxylase. Subsequently, it was shown that in the 7-butyrobetaine hydroxylase reaction (313) and then in the prolyl hydroxylase reaction (24.2)that lSOa was incorporated into the succinate formed. The stoichiometry of all the a-keto acid dioxygenase reactions has now been established, and molecular oxygen has been shown to be incorporated into succinate in all of the reactions except those catalyzed by pyrimidine deoxyribonucleoside 2‘-hydroxylase and lysyl hydroxylase. As further support for the mechanism depicted in Fig. 5 , Lindstedt and co-workers (313) claim to have developed a model system in which t-butyl hydroperoxide and a-ketoglutarate react to produce t-butyl alcohol and succinic acid. Nonenzymic reactions of hydroperoxides with carbonyl groups to form hydroxy peroxides (375-378) and analogous peroxide fragmentation reactions (379,380) are known. The nonenzymic oxidative decarboxylation of a-keto acids by peroxides occurs even under the mild conditions tolerable to enzymes (381).However, Hamilton favors an oxenoid mechanism (381,389) since it is easier to explain an initial oxidative attack on a-ketoglutarate than on the inactivated carbon atoms of such substrates as proline and y-butyrobetaine. The oxenoid reagent, persuccinic acid, would be formed as follows: Oz + COOHC&C€&COCOOH

-

COOHCH,CH,C, /p + OOH

co,

(33)

[For the review of evidence indicating peracids can convert alkanes to alcohols, see Hamilton (382) .] Persuccinic acid has been tested as a substitute for a-ketoglutarate in the prolyl hydroxylase reaction by Cardinale and Udenfriend (144,145) and, subsequently, in the pyrimidine deoxyribonucleoside 2’hydroxylase reaction by Liu and Abbott (146). I n neither case was any hydroxylation detected, but, of course, enzyme-bound persuccinate or other peroxy intermediates could not be ruled out by such experiments. In accord with the oxenoid mechanism is the demonstration by Cardinale 376. E. G. E. Hawkins, in “Organic Peroxides,” p. 274. Van Nostrand-Reinhold, Princeton, New Jersey, 1961. 377. C. A. Bunton, in “Peroxide Reaction Mechanisms” (J. 0. Edwards, ed.), p. 16. Wiley (Interscience), New York, 1962. 378. M. C. V. Sauer and J. 0. Edwards, J . Phys. Chem. 75, 3377 (1971). 379. W.H.Richardson and R. S. Smith, JACS 89,2230 (1967). 380. W. H. Richardson and T. C. Heesen, J. Org. Chem. 37, 3416 (1972). 381. G.A. Hamilton, Progr. Bioorg. Chem. 1, 83 (1971). 382. G.A. Hamilton, in “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 405. Academic Press, New York, 1974.

3.

OXYGENASES : DIOXYGENASES

185 OH

or 0

OH

OH

CH2 I

OH

'OH

0

CH2 -C@ 'OH

FIG.6.Oxenoid mechanism proposed for the p-hydroxyphenylpyruvate hydroxylase reaction. From Hamilton (381). and Udenfriend (144,l 4.5) of the decarboxylation of a-ketoglutarate by large amounts of prolyl hydroxylase in the absence of peptidyl substrate. This putative partial reaction yielded, in a 20-min incubation, about onefortieth as many moles of CO, per mole of enzyme as did the whole reaction (presence of peptidyl substrate) and was dependent on Fez+,ascorbate, oxygen, and active enzyme. I n preliminary studies (383), which included treatment of the enzyme with collagenase, no evidence was obtained for contamination of the prolyl hydroxylase preparation with prolyl substrate. Lindstedt and Lindstedt have reported ($07) that such an uncoupling of decarboxylation from hydroxylation does not occur in the y-butyrobetaine hydroxylase reaction. The oxenoid mechanism (Fig. 6) , as well as others, proposed for the p-hydroxyphenylpyruvate hydroxylase reaction has recently been reviewed (145). All of the a-keto acid dioxygenases studied have been shown to have requirements for a metal ion and a reducing agent. I n every instance the former requirement could only be satisfied with Fez+,whereas several substances fulfilled the latter. Ascorbate was frequently more effective than other reducing agents, but there is no direct evidence that ascorbate per se is in vivo the cofactor. In fact, the hydroxylation of prolyl residues has been demonstrated in cultured fibroblasts which have been reported not t o contain ascorbate (167,384).The function of the reductant may 383. G. J. Cardinale and S. Udenfriend, personal communication. 384. C. Bublitz and R. E. Priest, Lab. Invest. 17, 371 (1967).

186

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

be to maintain iron in the reduced form or to prevent oxidation of sulfhydryl groups (f219,306,534) of the enzyme. Ascorbate and Fez+ could be required for the generation of superoxide ion, Ot7, (385,386) since oxygen must be activated by some mechanism prior to hydroxylation. The studies of Hirata and Hayaishi (65,69) with tryptophan 2,3-dioxygenase made this a particularly attractive possibility. They have not only shown that this reaction is inhibited by superoxide dismutase but also that the ascorbate requirement can be met by superoxide generating systems. Bhatnagar and Liu (387,388) have reported that superoxide ion is involved in the prolyl hydroxylase reaction, but their evidence is unconvincing. Not only did they fail to report inhibition with superoxide dismutase but also this enzyme was used by Cardinale and Udenfriend (144) in unsuccessful attempts to inhibit the reaction. Moreover, the pyrimidine deoxyribonucleoside 2’-hydroxylase reaction was not inhibited with the dismutase in experiments in which the pH and ionic strength of the incubation mixture were varied (589). Of course, even if these types of experiments eventually implicate 0 2in~the a-keto acid dioxygenase reaction, they will not show that this ion reacts directly with the substrate. The requirements for Fez+and ascorbate also bring to mind the chemical model systems which have been used to study nonenzymic hydroxylation (for reviews, see references 382,390-394). The Udenfriend model system (395), which requires Fez+and ascorbate, as well as molecular oxygen, can effect such reactions as the hydroxylation of aromatic compounds and of saturated hydrocarbons and the oxidative demethylation of N-methyl groups. 385. S. Nakamura, BBRC 41, 177 (1970). 386. I. Fridovitch, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 453.Academic Press, New York, 1974. 387. R. S. Bhatnagar and T. Z. Liu, FEBS (Fed. Eur. Biochem. Soc.) Lett. 28, 226 (1972). 388. T.Z.Liu and R. S. Bhatnagar, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 32, 613 1973). 389. C. K. Liu and M. T.Abbott, unpublished. 390. R. 0. C. Norman and J. R..L. Smith, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 131. Wiley, New York, 1965. 391. V. Ullrich and Hj. Staudinger, in “Biological and Chemical Aspects of Oxygenases” (K. Block and 0. Hayaishi, eds.), p. 235. Maruzen, Tokyo, 1966. 392. G.A. Hamilton, Advan. Enzymol. 32, 55 (1969). 393. V. Ullrich, H.-H. Ruf, and H. Mimoun, in “Biological Hydroxylation Mechanisms” ( G . S. Boyd and R. M. S. Smellie, eds.), p. 11. Academic Press, New York, 1972. 394. R. K. Locke and V. W. Mayer, Biochem. Pharmacol. 23, 1979 (1974). 395. S. Udenfriend, C. T. Clark, J. Axelrod, and B. B. Brodie, JBC 208, 731 (1954).

3.

OXYGENASES : DIOXYGENASES

187 R-CHO

+ He0

% +

I

’\\

,. I\ \\I

’

/’

I’

Reducinq aqenl f.2’

t

,,

I

I CHe I CHL

+ HpO

,‘

I ‘\

COOH

R-COOH

.t

-

COOH

I

CHC

I

CHI

+

cop

I

cocow

COOH

a- Ketoglularate

Succinate

FIG.7. The conversion of a methyl group to a carboxyl group via reactions with a common mechanism.

The enzymic conversion of 5-hydroxymethyluracil to 5-formyluracil and of the latter to uracil-5-carboxylic acid may occur by essentially the same mechanism as does the conversion of thymine to 5-hydroxymethyluracil (Fig. 7). This proposal (3ZO,SZ5) is consonant with the stoichiometry of these reactions, the studies with and the catalysis of the three reactions by a single enzyme. The 5-hydroxymethyluracil and 5-formyluracil dioxygenase reactions are unique in that no other dioxygenase is known that catalyzes either the conversion of an alcohol to an aldehyde or of an aldehyde to an acid. Although dehydrogenases which catalyze these types of oxidations are more familiar, it appears that similar oxidations are catalyzed by microsomal monooxygenases in NADPH-dependent reactions (596-398). The mechanism proposed for the a-keto acid dioxygenase reactions may be a more general phenomenon of oxygenase reactions; for example, one might regard monooxygenase reactions as those requiring a reducing agent such as NADPH, rather than decarboxylation of an a-keto acid, for fragmentation of a peroxide intermediate (145). Another reaction in which decarboxylation appears to be involved in fragmentation of an intermediate hydroperoxide is that catalyzed by lactate oxygenase :

+

OZ+ lactate -+ acetate + CO, HzO (34) This is a monooxygenase reaction in which molecular oxygen is incorporated into the products, acetate and water. The oxygenase is a flavoprotein with F M N as the prosthetic group. In studies on the mechanism 396. P. J. Murphy and C. A. West, ABB 133,395 (1969). 397. R. Teschke, Y. Hasumura, J.-G. Joly, H. Ishii, and C. S. Lieber, BBRC 49, 1187 (1972). 398. J. L. Gaylor, S.-T.Hsu, C. V. Delwiche, K. Comai, and H. E. Seifried, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 2, p. 431.Univ. Park Press, Baltimore, Maryland, 1973.

188

OSAMU HAYAISHI, MITSUHIRO NOZAKI, AND MITCHEL T. ABBOTT

PH-

hv

FIO.8. Oxidative mechanism proposed for firefly bioluminescence. From McElroy and DeLuca (401). of this reaction, Massey and co-workers (399,400) have obtained evidence that an enzyme. FMNH, pyruvate complex reacts with oxygen to yield acetate, CO,, H,O, and enzyme-bound FMN. The reaction of this complex with oxygen appears to be analogous to the partial reaction demonstrated by Cardinale and Udenfriend (144145) in which prolyl hydroxylase catalyzes the decarboxylation of a-ketoglutarate in the absence of the peptidyl substrate. Oxygenase reactions are also coupled to decarboxylations in bioluminescent systems (401-403). One type of reaction is shown in Fig. 8, but in another one the excited state product molecule is produced in a decarboxylation step which fragments a four-membered peroxide ring (409). If a-ketoglutarate and various biological reducing agents do serve a similar function in hydroxylation, regulatory considerations may be important factors in rationalizing why a given hydroxylase is linked to a particular cofactor. In this vein, it may be interesting to test a-keto-y-hydroxyglutaric acid as an inhibitor and substitute for a-ketoglutarate in the prolyl hydroxylase reaction since t’his analog of a-ketoglutarate has

-

399. 0. Lockridge, V. Massey, and P. A. Sullivan, JBC 247, 8097 (1972). 400. M. S. Flashner and V. Massey, in “Molecular Mechanisms of Oxygen Activation’’ (0.Hayaishi, ed.), p. 245. Academic Press, New York, 1974. 401. M. DeLuca and M. E. Dempsey, in “Chemiluminescence and Bioluminescence” (M. J. Cormier, D. M. Hercules, m d J. Lee, eds.), p. 285. Plenum, New York, 1973. 402. M. DeLuca and M. E. Dempsey, in “Chemiluminescence and Bioluminescence” (M. J. Cromier, D. M. Hercules, and J. Lee, eds.), p. 345. Plenum, New York, 1973. 403. 0. Shimomura and F. H. Johnson, in “Chemiluminescence and Bioluminescence” (M. J. Cormier, D. M. Hercules, and J. Lee, eds.), p. 337. Plenum, New York, 1973.‘

3.

OXYGENASES : DIOXYGENASES

189

been established as an intermediate in the breakdown of hydroxyproline

(404,406). ACKNOWLEDGMENTS The author of Section 111 would like to thank Professor F. I. Tsuji for his critical reading of Section 111. The author of Section IV (Mitchel T. Abbott) wishes to thank Dr. G. J. Cardinale and Dr. S. Udenfriend for most helpful discussions and for sending him a preprint of their review article (144). The author is also indebted to his colleagues Dr. W. H. Richardson and Dr. S. A. Dahms for the stimulating suggestions and critical comments. In addition, the support of the National Institutes of Health, National Institute of Arthritis and Metabolic Diseases (AM0 9314) is gratefully acknowledged.

404. U. Maitra and E. E. Deliker, JBC 238, 3660 (1973). 405. E. Adams, Mol. Cell. Biochem. 2, 109 (1973).

This Page Intentionally Left Blank

Flavin and Pteridine Monooxygenases VINCENT MASSEY I . Introduction

PETER HEMMERICH

. . . . . . . . . . . . . . . . . 191

I1. Internal Flavoprotein Monooxygenaaes

. . . . . . . . .

A . Lactate Monooxygenase (Lactate Oxidase or Lactate Oxidative Decarboxylase) . . . . . . . . . . . . . . B . Lysine Monooxygenase . . . . . . . . . . . . C Arginine Monooxygenme (Arginine Decarboxylase) . . . . 111. External Flavoprotein Monooxygenases . . . . . . . . A. Salicylate Hydroxylase . . . . . . . . . . . . . B . pHydroxybenzoate Hydroxylase . . . . . . . . . C . Melilotate Hydroxylase . . . . . . . . . . . . D Phenol Hydroxylase . . . . . . . . . . . . . E Orcinol Hydroxylase . . . . . . . . . . . . . F. m-Hydroxybenzoate-6-hydroxylase . . . . . . . . . G. m-Hydroxybenzoate-4-hydroxylase . . . . . . . . . H Imidazolylacetate Monooxygenase . . . . . . . . . I Bacterial Luciferase . . . . . . . . . . . . . J . Microsomal Amine Oxidase . . . . . . . . . . . K . Kynurenine-3-hydroxylase . . . . . . . . . . . I V Pterin-Linked Monooxygenases . . . . . . . . . . . A Phenylalanine Hydroxylase . . . . . . . . . . . B . Tyrosine Hydroxylme . . . . . . . . . . . . . C . Tryptophan Hydroxylase (Tryptophan-5-monooxygenaae) . . V Model Studies and Possible Mechanisms . . . . . . . .

.

. . . .

.

.

.

193 194 199 203 204 206 211 217 221 223 224 225 225 226 229 230 231 232 238 240 241

.

1 Introduction

The reactivity with oxygen of reduced flavins and flavoenzymes is one of the most interesting problems in the field of flavin chemistry and will be reviewed in more detail a t the end of this chapter . Although the reac191

192

VINCENT MASSEY AND PETER HEMMERICH

tion is complex, involving the formation of highly reactive peroxydihydroflavins, flavin (Fl) radicals, and superoxide anion, 0,- (1-3), it is rapid (tl,, < 1 sec) and results is the stoichiometric production of H,O,: Fbed

4-Oz+ FI,,

+ HzOz

In the case of many flavoproteins, which are converted to the reduced form by the specific substrates whose oxidation they catalyze, the high rate of reaction with 0, may be retained or even enhanced, in which case they are classified as oxidases. On the other hand, many flavoenzymes react very slowly with 0, and much more rapidly with one-electron acceptors ; these enzymes are classified as dehydrogenases. Although the route of flavin reoxidation appears to be different in the two classes, involving the intermediate production of 0,- and flavin semiquinone in the case of the dehydrogenases and direct two electron oxidation in the case of the oxidases (4,6), the product of reaction with 0, in both classes, as with the free flavins, is H,O,. In contrast to these H,O,-producing enzymes, another group of flavoproteins has become recognized in recent years, the flavoprotein rnonooxygenases. This group of enzymes, also known as mixed function oxygenases or mixed function oxidases, in reaction with 0, cause the incorporation of one atom of the O2molecule into the substrate to yield an oxygenated product and the conversion of the other oxygen atom to H,O. The flavoprotein monooxygenases can be further conveniently subclassified on the basis of the electron donor involved in the catalytic reaction. In one group of enzymes the substrate itself serves as the electron donor; this group is therefore referred to as internal monooxygenases. I n the second group, in addition to the substrate to be hydroxylated, an external reductant such as NADH or NADPH is required for catalysis ; this group is therefore known as external monooxygenases. Since the reactions catalyzed by this group are hydroxylations, these enzymes are also often referred to as flavoprotein hydroxylases. I n addition to the numerous flavoprotein monooxygenases, there is a small group of enzymes which require tetrahydropteridines as cofactors. In view of the similar chemistry of pteridines and flavins, it is widely considered that the flavoprotein and pteridine-linked monooxygenases 1. Q. H. Gibson and J. W. Hastings, BJ 68, 368 (1962). 2. D. Ballou, G. Palmer, and V. Massey, BBRC 36, 898 (1969). 3. V. Massey, G. Palmer, and D. Ballou, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 5. Univ. Park Press, Baltimore, Maryland, 1973. 4. V. Massey, S. Strickland, S. G. Mayhew, L. G. Howell, P. C. Engel, R. G . Matthews, M. Schuman, and P. A. Sullivan, BBRC 36, 891 (1969). 5. V. Massey, F. Muller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G. Howell, S. G. Mayhew, R. G. Matthews, and G. P. Foust, JBC 244, 3999 (1969).

4.

193

FLAVIN AND PTERIDINE MONOOXYGENASES

may fanction by basically similar mechanisms. The similarity in structure of dihydroflavins and tetrahydropteridines is shown below:

Di hydrof lavin

Tetrahydrobiopterin

Although model studies support the concept that hydroxylation reactions involving dihydroflavins and tetrahydropteridines proceed via similar mechanisms (see Section V) there are two important pieces of experimental evidence which cast some doubt on this hypothesis. First, unlike the rapid reaction of dihydroflavins with 0, (tlIz< 1 sec), tetrahydropteridines in general are only sluggishly oxidized by O2 ( tlIz 5 min) . Unless the combination with protein dramatically enhances the 0, reactivity (for which no experimental evidence exists) it would appear unlikely that this autoxidation could be fast enough to be of catalytic significance. Second, in contrast to the flavoprotein monooxygenases, which have all been shown to have no metal involvement, the present evidence indicates strongly that the pteridine-linked monooxygenases are iron-containing enzymes. Thus, the possibility exists that the function of the tetrahydropteridines is to reduce the iron, and that it is the latter moiety which is responsible for oxygen activation, or that a tetrahydropteridine-Fe (111) complex is responsible. Several reviews concerned with flavoprotein monooxygenases (6-10) and pteridine-linked monooxygenases (11-13) have appeared previously. H

II. Internal Flavoprotein Monooxygenases

Internal flavoprotein monooxygenases catalyze oxidative decarboxylations in which an oxygen atom is incorporated into the substrate and 6. 0. Hayaishi, in. “Oxygenases” (0. Hayaishi, ed.), p. 1. Academic Press, New York, 1962. 7. 0. Hayaishi, Bacteriol. Rev. 30, 720 (1966). 8. 0. Hayaishi and M. Nozaki, Science 164, 389 (1969). 9. 0. Hayaishi, Annu. Rev. Biochem. 38, 21 (1969). 10. M. S. Flashner and V. Massey, in “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 245. Academic Press, New York, 1974. 11. S. Kaufman, in “Oxygenases” (0. Hayaishi, ed.), p. 129. Academic Press, New York, 1962. 12. S. Kaufman, Advan. Enzymol. 35, 245 (1971). 13. S. Kaufman and D. B. Fisher, in “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 285. Academic Press, New York, 1974.

194

VINCENT MASSEY AND PETER HEMMERICH

CO, and H,O are the other products. The substrate itself serves as a reductant of the flavin. Hence, these reactions in effect involve a double oxidation of the substrate, which is first oxidized at the expense of flavin reduction and then oxidized again by the insertion of oxygen. These reactions may be formalized as follows: H I R-C-COOH I

+

EFbx

R-C-COOH.EFlredHz

[::

R-C-COOH*EFlredH, II

X

XH

1

+

02-

EFlox

+

R-CZO

XH

+

C02

+

%O

It has been suggested that the mechanism by which the oxygen atom is incorporated into the product may be quite different with the internal flavoprotein monooxygenases than with the external monooxygenases (10). In the latter group there is growing evidence for the direct participation in the hydroxylation reaction of covalent adducts of oxygen and dihydroflavin. I n the case of the internal monooxygenases the active form of oxygen involved may be H,O,, the normal product of autoxidation of dihydroflavins. The nonenzymic decarboxylation of keto acids by H,O, is a well-established reaction, whose mechanism has recently been investigated (14). Similar oxidative decarboxylation of imino acids would be expected. As will be discussed in more detail in Sections II,A and B, the possibility exists that the decarboxylation reactions catalyzed by this group of enzymes result from the locally high concentration of H,Oz and keto acid (or imino acid) a t the active site. In this case the flavoprotein internal monooxygenases would be in fact typical flavoprotein oxidases ; the monooxygenase activity would then be an adventitious one because of the nature of the primary oxidase products and their mutual reactivity. A. LACTATE MONOOXYGENASE (LACTATE OXIDASEOR LACTATE OXIDATIVEDECARBOXYLASE) Lactate monooxygenase catalyzes the incorporation of molecular oxygen into lactate. Historically this was the first flavoprotein monooxygenase studied. In 1955, Sutton (15) reported the purification of the enzyme from Mycobactenurn phlei and established the prosthetic group to be FMN. The enzyme was obtained in crystalline form in 1957 (16) and 14. G. A. Hamilton, Progr. Bioorg. Chem. 1, 83 (1971). 15. W. B. Sutton, JBC 216, 749 (1955). 16. W. B. Sutton, JBC 226, 395 (1957).

4. FLAVIN AND PTERIDINE MONOOXYGENASES

195

reported to have a minimum molecular weight per flavin of 125,700. This value was based simply on the ultraviolet absorption; later workers have determined the minimum molecular weight to be 55,000-56,000 on the basis of dry weight and biuret determinations (17 ) . The oxygenase nature of the enzyme was established by Hayaishi and Sutton by ISO studies (18). They found that ISO was incorporated into acetate when ‘*02 gas and H2’“0was used, but not with ISO, gas and H21s0. I n the same year, Sutton demonstrated that pyruvate was the product when the enzyme was reduced anaerobically by lactate. Also, by the use of carbonyl trapping agents he was unable to detect any free pyruvate formed in aerobic catalysis, and concluded that any pyruvate formed must remain enzymebound (16). Beinert and Sands (19),with enzyme from M . phlei and a rapid scanning spectrophotometer, demonstrated the presence of a transient longwavelength intermediate which appeared under both aerobic and anaerobic conditions. I n 0.07 M phosphate buffer, pH 7.5, the intermediate was formed in a biphasic manner and disappeared slowly. Stopped-flow measurements performed in 0.05 M phosphate buffer showed a similar pattern ( 2 0 , d l ). These results, implying considerable mechanistic complexity, have since been demonstrated to occur from the use of phosphate as a buffer (see below). Sullivan (22) has purified and crystallized a lactate oxidase from Myco bacterium smegmatis with properties very similar to the enzyme from M . phlei. The enzyme has a minimum molecular weight of 49,500 per FMN. From sucrose gradient and sedimentation velocity data molecular weights of 300,000 and 341,000 were obtained, indicating that the enzyme is composed of multiple subunits. Using the enzyme from M . smegmatis, Lockridge et a,?. (23) have shown that lactate oxidase binds a large variety of anions, which inhibit enzymic activity. This inhibition is competitive with L-lactate, and most of the anions cause marked perturbations in the visible spectrum of the M at pH 7.0, 25O. I n enzyme. Phosphate binds with a Ka of 1.3 X 17. S. Takemori, K. Nakazowa, Y . Nakai, K. Suzuki, and M. Katagiri, JBC 243, 313 (1968). 18. 0. Hayaishi and W. B. Sutton, JACS 79,4809 (1957). 19. H. Beinert and R. H. Sands, in “Free Radicals in Biological Systems” (M. S. Blois, Jr., et al., eds.), p. 35. Academic Press, New York, 1961. 20. S. Takemori, Y . Nakai, M. Katagiri, and T. Nakamura, FEBS (Fed. Eur. Biochem. Soc.) Lett. 3,214 (1969). 21. M. Katagiri and S. Takemori, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 447. Univ. Park Press, Baltimore, Maryland, 1971. 22. P. A. Sullivan, BJ 110,363 (1968). 23. 0. Lockridge, V. Massey, and P. A. Sullivan, JBC 247, 8097 (1972).

196

VINCENT MASSEY AND PETER HEMMERICH

phosphate buffer the biphasic formation of a long wavelength absorbing intermediate similar to that reported for the M . phlei enzyme was also observed when enzyme was reduced anaerobically with L-lactate. However, in the absence of phosphate (and other inhibiting anions) a rapid monophasic formation of intermediate was observed, followed by a slow decay to fully reduced enzyme. The biphasic formation of intermediate was shown to result from the lactate being able to reduce only enzyme uncomplexed with phosphate (the rapid phase) ; the slow phase was determined by the rate of dissociation of phosphate from the inactive enzymephosphate complex. Lockridge et al. have also shown that the intermediate seen transiently is a complex of reduced enzyme and pymvate and that this is the species reacting with 0, in catalysis. Similar intermediates but of different stability and 0, reactivity were detected with a number of a-hydroxy acid substrates. In each case the spectrum corresponding to that of the intermediate produced transiently on anaerobic reduction could be obtained on addition of the corresponding keto acid to the free reduced enzyme. The rate of reaction of the reduced enzyme with O2 was markedly different depending on whether the reduced enzyme was uncomplexed or complexed with keto acid. I n the case of the reduced enzyme-pyruvate complex the rate of reaction with 0, was more than 200 times greater than with uncomplexed enzyme. On the basis of steadystate and stopped-flow kinetic analysis, the following reaction scheme for the enzyme is proposed (23) (cf. Table I) :

:.,

E * FMN * R-CHOH-

EmFMN

+

R-CHOH-COOH

COOH

/E

R- COOH co* H,O

*

k-31

+ +

E-FMNH,

+

E-FMN

+

R-CO-COOH

E * FMN R-CO-COOH 1

H,O,

I n this reaction scheme the catalytic pathway of oxidative decarboxylation follows the steps with rate constants k,, k,, k,, and k,. The step represented by k , is the slow release of a stoichiometric amount of pyruvate from the catalytic intermediate, in accord with the results first

TABLE I KINETICCONSTANTS OF LACTATE OXIDASE' Const ant Kd

=

k-i/ki

kz ka

ka ks

V,,,, catalytic K , (RCHOHCOOH) K , (02)

tLactate

8-Phenyl klactate

Ira-Hydroxyisovalerate

5 x M 14,000 min-l 2 . 5 min-1 1 . 1 X 108 M-1 min-1 (observed) 11,300 min-l 6,250 min-' 2.23 X M 7 . 1 x 10-544

0.34 M 6 , 7 0 0 min-I -2 min-1 5 . 7 X 106 M-1 min-1 (calculated) 1,060 min-l 910 min-1 5 x 10-2 M 1 . 6 x 10-4 M

9 x 10-2 M 3,700 min-' - 0 . 2 min-1 3 . 3 x 106 M-1 min-1 (calculated) 1,370 min-l 1,000'min-1 2.5 X M 3 x 10-4 M

ks (Free enzyme) a

5 . 4 X 106 M-1 min-1

Values were obtained in 0.01 M imidazole-HCl, pH 7.0, 25". From Lockridge et al. (23).

La-HydroxyB-methylvalerate 3 x M 590 min-' 2 . 7 X lo6 M-' min-l (observed) 350 min-l 220 min-1 1.13 X M 8 X M

198

VINCENT MASSEY AND PETER HEMMERICH

obtained by Sutton (16) and confirmed in the above work. It was also shown by Lockridge et al. that when the keto acid is allowed to dissociate from the reduced enzyme intermediate ( t n w 0.3 min) and 0, is introduced, a stoichiometric amount of H,Oz is produced. On the other hand, when free reduced enzyme was complexed with 2-14C-labeled pyruvate and 0, admitted, little HzO, was detected and a stoichiometric amount of 14CH,COOH was formed. The finding that the form of the enzyme which reacts with 0, is a complex of E-FMNH, and keto acid strongly suggests that the mechanism of decarboxylation may follow simply from the enzyme providing locally high concentrations of reactants. The nonenzymic decarboxylation of keto acids by H,O, has been known for many years; however, high concentrations of the reactants in free solution are required for rapid reaction. With lactate oxidase it seems reasonable to postulate that the E.FMNH,-pyruvate complex in its rapid reaction with 0, yields an ESFMN-pyruvate complex, and that the H20zproduced a t the active site reacts with pyruvate before either product has a chance to dissociate from the enzyme. In this case the primary products of the catalytic reaction, pyruvate and H,O,, are typical oxidase products, similar to those from the amino acid oxidases (imino acid and H,O,). The overall monooxygenase reaction is exhibited only because of the slow release of the products, which at the locally high concentration a t the active center can react rapidly in the secondary decarboxylation reaction. Thus, an “uncoupling” of monooxygenase function could result either from an enhanced rate of product dissociation from the reduced enzyme-product complex (k, in the above scheme) or by enhancement of the rate of release of either H,O, or product from the ternary complex of oxidized enzyme, product and H,O,. As will be discussed in the following section, numerous examples of such “uncoupling” have been documented. It is intriguing to speculate that the amino acid oxidases (which have been shown to operate by mechanisms basically similar to that of lactate oxidase) would also demonstrate such adventitious monooxygenase activity if the imino acid and H,O, products were only released slowly from the enzymes. It should be emphasized that this concept in no way detracts from the probability of a peroxydihydroflavin intermediate preceding the production of H,O, (cf. Section V ) . The postulate is made simply on the grounds that the same oxidative decarboxylation reaction occurs in the absence of enzyme. The possibility exists that a peroxydihydroflavin form of the enzyme is the active oxygenating agent reacting with keto acid; however, in that case step k, must result in the production of an E.FAD.acetate.COe.HzO complex, and the partially rate-limiting step k, must result in the slow release of at least one of the products.

4. FLAVIN

AND PTERIDINE MONOOXYGENASES

199

In further keeping with the basic similarity of lactate oxidase and Damino acid oxidase, i t has also been shown that proton abstraction from the a-carbon atom of the substrate by an enzyme base is an early step in the reaction mechanism of lactate oxidase. Thus, p-chlorolactate is a substrate for lactate oxidase under anerobic conditions resulting in the catalytic elimination of chloride ion and the formation of pyruvate ( 2 4 ) . Under aerobic conditions, as well as the elimination reaction, p-chlorolactate also behaves as a normal substrate, yielding chloroacetate, CO,, and H,O as products. Like the analogous reactions of D-amino acid oxidase with p-chloroalanine (65) the partition between the two catalytic pathways of elimination and oxidation depends strongly on the oxygen concentration, indicating competition of the two pathways for a common intermediate. This intermediate may very well be the complex of reduced enzyme and /3-chloropyruvate. Further evidence for the proton abstraction mechanism comes from recent studies with an acetylenic substrate. Walsh et al. (26) have shown that L-2-hydroxy-3-butynoic acid is a substrate for lactate oxidase as well as an inactivator. The rate of inactivation varies inversely with oxygen concentration, suggesting a partitioning of an enzyme-substrate intermediate between two reaction pathways, one leading to normal catalytic turnover, the other to inactivation. The inactivation has been shown to result from the formation of the covalent addition of a substrate moiety to the reduced flavin. Experiments with the acetylenic substrate labeled with tritium a t the a-carbon atom showed that the inactivated enzyme contained no tritium, indicating that the substrate must lose its a-hydrogen atom before inactivation occurs. When the label was a t carbon-4, inactivated enzyme contained 1 mole of tritium, associated with the flavin. Since inactivation may also be achieved by incubating reduced enzyme with the corresponding acetylenic keto acid (27), it appears likely that the partitioning between catalysis and inactivation occurs a t the level of the E.FMNH,-keto acid complex.

B. LYSINEMONOOXYGENASE Lysine monooxygenase has been isolated in crystalline form from Pseudomonas fluorescens (28).It ,catalyzes the oxidative decarboxylation of L-lysine to 8-aminovaleramide: 24. C. T. Walsh, 0. Lockridge, V. Massep, and R. H. Abeles, JBC 248, 7049 (1973). 25. C. T. Walsh, A . Schonbrunn, and R. H. Abeles, JBC 246, 6855 (1971). 26. C. T. Walsh, A. Schonbrunn, 0. Lockridge, V. Massey, and R. H. Abeles, JBC 247, 6004 (1972). 27. S. Ghisla, V. Massey, C. T. Walsh, and R. H. Abeles, unpublished observations. 28. H. Takeda and 0. Hayaishi, JBC 241, 2733 (1966).

200

VINCENT MASSEY AND PETER HEMMERICH

L- Lysine

6-Aminovaleramide

Studies with l*OZshowed that molecular oxygen is indeed incorporated into the product ( 2 9 ) . L-Arginine is also a substrate, being decarboxylated to y-guanidinobutyramide a t a rate approximately 10% that of L-lysine (SO). The crystalline enzyme exhibits a typical flavoprotein absorption spectrum with maxima at 274, 385, and 460 nm, and was found to contain FAD as prosthetic group ( 2 8 ) . Careful studies with metal chelating agents and metal analyses (31) indicate that metals play no role in catalysis and that the enzyme is a simple flavoprotein with FAD as the sole prosthetic group. The molecular weight has been calculated as 191,000 from sedimentation velocity experiments, and the FAD content was estimated to be two equivalents per molecule of protein (SO). However, a recent study indicates that this analysis may have been in error through contamination with nonflavoprotein impurities ; Flashner and Massey (S2) estimated the molecular weight to be 240,000-246,000with four FAD residues per molecule of protein. Lysine monooxygenase has several unusual properties as compared with other flavoprotein monooxygenases ; for example, the anaerobic reduction of the enzyme-bound FAD by equimolar concentrations of lysine is an extremely slow process requiring 7-9 hr for full reduction (SO). An explanation of this puzzling phenomenon comes from the finding (33) that lysine plays the role of an effector as well as that of a substrate. Evidence was found for there being a regulatory site as well as a catalytic site; i t was necessary for the regulatory site to be occupied by lysine (or nonsubstrate effectors such as c-aminocaproate) for the enzyme to be in a catalytically active form. These findings will be discussed further later in this section. Under anaerobic conditions, the reduction of the enzyme by lysine represults in the stoichiometric formation of A1-piperideine-2-carboxylate, sumably by the primary dehydrogenation of the substrate followed by 29. N. Itada, A. Ichihara, T. Makita, 0. Hayaishi, M. Suda, and N. Sasaki, J . Biochem. ( T o k y o ) 50, 118 (1961). 30. H. Takeda, S. Yamamoto, Y. Kojima, and 0. Hayaishi, JBC 244, 2935 (1969). 31. S. Yamamoto, H. Takeda, Y. Maki, and 0. Hayaishi, JBC 244,2951 (1969). 32. M. I. S. Flashner and V. Massey, JBC 249, 2579 (1974). 33. M. I. S. Flashner and V. Massey, JBC 249, 2587 (1974).

4.

201

FLAVIN AND PTERIDINE MONOOXYGENASES

hydrolysis of the resulting imino acid and cyclization of the intermediate a-keto r-aminocaproate (34-36‘).

L-Lysine

a-Imino, €-amino

caproic acid

a -Keto, €-amino

caproic acid

A’- Piperideine2-carboxylic acid

The last two steps of this reaction are presumably nonenzymic. The anaerobic formation of a-imino r-aminocaproate is analogous t o the anaerobic formation of pyruvate with lactate oxidase (16,dS) and suggests that a basically similar reaction pathway may exist for both enzymes. This suggestion is strengthened by the recent report of Nakazawa et al. (37) that aerobically lysine monooxygenase catalyzes a typical oxidase reaction with ornithine or 2,8-diaminooctanoate. I n these reactions the products are the corresponding keto acids and H,O,, without formation of CO,, analogous to the anaerobic reaction with lysine described above. Both the oxygenase-type reaction with lysine or arginine as substrate and the oxidase-type reaction with ornithine are kinetically complex, displaying sigmoidal substrate saturation curves (33,37). This phenomenon appears to result from the presence of two different types of substrate binding sites in the enzyme, a regulatory or effector site which must be occupied before the enzyme can react efficiently with substrate a t the catalytic site ( 3 3 ) . Flashner and Massey have shown that the regulator site may be occupied by lysine, arginine (both oxygenase substrates), L-ornithine (an oxidase substrate), or r-aminocaproate and L-lysine hydroxamate (both nonsubstrate effectors) to abolish the sigmoidal kinetic behavior with L-lysine concentration. It was also shown that lysine could act as an effector to permit the more rapid oxidation of c-N,N-dimethyl-L-ly sine. Recent work of Yamauchi et al. (38) showed that preincubation of 34. T. Nakazawa, S. Yamamoto, Y. Maki, H. Takeda, Y. Kajita, M. Nosaki, and 0. Hayaishi, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 214. Univ. of Tokyo Press, Tokyo, 1968. 35. S. Yamamoto, T. Nakarawa, and 0. Hayaishi, JBC 247, 3434 (1972). 36. S. Yamamoto, Y. Maki, T. Nakazawa, Y. Kajita, H. Takeda, M. Nosaki, and 0. Hayaishi, Advan. Chem. Ser. 77, 177 (1968). 37. T. Nakazawa, K. Hori, and 0. Hayaishi, JBC 247, 3439 (1972). 38. T. Yamauchi, S. Yamamoto, and 0. Hayaishi, JBC 248, 3750 (1973).

202

VINCENT MASSEY AND PETER HEMMERICH

the enzyme with sulfhydryl reagents transforms the activity toward L-Iysine or L-arginine from an oxygenase to an oxidase. Further work from the same laboratory (39) pertaining to the nature of the active site has shown that L-alanine, itself without activity, is made an efficient substrate if the other moiety of the L-lysine molecule, n-propylamine, is added. The activity so induced is an oxidase one; i.e., the products are pyruvate and H,O,. Other a-monoamino acids were also oxidized in the presence of alkylamines of various chain length; the highest oxidase activity was observed when the total chain length of both amino acid and amine was nearly identical to that of lysine. These striking results have been interpreted by Hayaishi and his colleagues as indicating two types of enzyme-substrate complexes: one of the correct type which leads to oxygenation, the other of an approximate fit in which the dehydrogenation of the substrate by FAD does not couple with oxygen activation, and the reduced flavin is simply oxidized by 0, to produce H,02. While the synergistic effect of a-monoamino acids and alkylamines certainly indicates a requirement for both the a-amino and a terminal basic residue, the results are capable of other interpretations. Perhaps the major effect is on the regulatory site and that a-amino acids could be oxidized a t the catalytic site without simultaneous occupation of this site by an alkylamine. An alternative hypothesis to explain the “uncoupling” phenomenon, i.e., the ability of the one enzyme to catalyze either oxygenase or oxidase reactions, has already been considered in detail in the previous section. According to this iiadventitious” monooxygenase hypothesis, the reaction with all substrates may be regarded as proceeding in two distinct halfreactions. The first of these involves the dehydrogenation of the substrate to produce an enzyme: FADH,.imino acid complex. The second step involves the reaction of this complex with 0, to produce an enzymeFAD-imino acid-HzOz complex. If both the imino acid and H,O, are only slowly dissociated from the complex, oxidative decarboxylation occurs (oxygenase activity). On the other hand, oxidase activity would be exhibited if either product dissociated rapidly from the quaternary complex, or alternatively if imino acid dissociated rapidly from the enzymeFADH,-imino acid complex. Yamamoto et al. (40) have used rapid reaction techniques to investigate the transient intermediates involved in both the aerobic and anaerobic reactions with lysine as substrate. They observed, during the anaerobic 39. S. Yamamoto, T. Yamauchi, and 0. Hayaishi, Proc. N u t . Acad. Sci. U . S. 69, 3723 (1972). 40. S. Yamamoto, F. Hirata, T. Yamauchi, M. Nozaki, K. Hiromi, and 0. Hayaishi, JBC 246, 5540 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

203

reduction of the enzyme, a transient long wavelength absorbing species which appeared in a biphasic manner and disappeared very slowly to yield reduced enzyme. The spectral characteristics of the intermediate are reminiscent of those first reported for D- and L-amino acid oxidases, i.e., complexes of E- F A D H , and imino acid (41,42). The significance of the biphasic appearance of the intermediate is unclear. However, this behavior is also similar to that found with D-amino acid oxidase in the presence of inhibitors (41) and of lactate oxidase when inhibitory ions are present ( 2 3 ) . Indeed, lysine monooxygenase has been found to be very susceptible to inhibitory ions ( 3 2 ) .Hence, the possibility is very real that the observed complexities in the kinetics of formation of the intermediate are related to buffer inhibition effects.

c. ARGININE MONOOXYGENASE (ARGININE DECARBOXYLASE) Arginine monooxygenase and lysine monooxygenase are closely related in properties and function. Both enzymes catalyze the oxidative decarboxylation of basic amino acids. However, substrate specificity appears to be stricter for arginine monooxygenase, which does not attack L-lysine ( 4 3 ) .On the other hand, lysine monooxygenase will also use L-arginine as an oxygenase substrate at 10% the rate of its reaction with L-lysine. Both enzymes exhibit sigmoidal saturation curves with their substrates and both have pH optima in the region of p H 9. The enzyme was partially purified from the washed mycelium of Xtreptomyces griseus (43) and shown to convert L-arginine to 7 -guanidinobutyramide.

F C=NH CHNH, I COOH L

- Arginine

y

2

C=NH

C -NH, II

0

y-Guanidinobutyramide

Two other compounds similar in structure to arginine were also found to be substrates (canavanine and homoarginine), also being converted to their corresponding amides but a t lower rates (4). 41. V. Massey and Q. H. Gibson, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 23, 18 (1964). 42. V. Massey and B. Curti, JBC 242, 1259 (1967). 43. N. V. Thoai and A. Olornucki, BBA 59, 533 (1962). 44. N. V. Thoai and A. Olomucki, BBA 59,545 (1962).

204

VINCENT

MABSEY

AND PETER HEMMERICH

The oxygenase nature of the enzyme was shown by

1802

experiments

(46). Further studies on a more purified preparation revealed the enzyme to be a flavoprotein with FAD as prosthetic group (46). ThomB-Beau et al. (47) have shown that treatment with diethylpyrocarbonate inacti-

vates the enzyme with the modification of one histidyl residue per mole of enzyme-bound flavin. Arginine protects the enzyme completely from this inactivation. These data, together with a study of the pH dependence of the I(, for L-arginine, indicate that one histidyl residue per flavin is required for catalytic activity, and that this histidyl residue is part of the active center of the enzyme. In light of the finding that proton abstraction from substrate appears to be an early step in catalysis by several flavoproteins (24-26) , the possibility exists that the essential histidyl residue of arginine monooxygenase may function by abstracting a proton from the a-carbon atom of the substrate.

111. External Flavoprotein Monooxygenases

With the external flavoprotein monooxygenases, an external reductant such as NADH or NADPH is required for catalytic activity. A large number of enzymes has been found to belong to this group, and a t least for those cases which have received considerable experimental attention, remarkable similarities in properties have emerged. Most of the enzymes are of bacterial origin and may be induced by growing the microorganism in cultures containing the substrate as sole carbon source. Most of the enzymes catalyze hydroxylation reactions (and hence are often called hydroxylases) which utilize an aromatic compound as substrate. I n most cases the hydroxylated product is either more soluble or more readily metabolized than the substrate ; hence, these enzymes aid considerably in the detoxification process of aromatic compounds and constitute an important part of pollution control and detoxification carried out by soil microorganisms. On the molecular level, the chief and remarkable common property of this group of enzymes is the dual role played by the substrate, which a t the same time is an effector as well as a substrate. The effector role is exhibited chiefly in the enormous stimulation of the rate of reduction of the enzyme flavin by NADH or NADPH when the substrate is bound. 45. D. B. Pho, A. Olomucki, and N. V. Thoai, BBA 118,299 (1966). 46. A. Olomucki, D. B. Pho, R. Lebar, L. Delcambe, and N. V. Thoai, BBA 151, 353 (1968). 47. F. ThomCBeau, Le-Thi-Lan, A. Olomucki, and N. V. Thoai, Eur. J . Biochem. IS, 270 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

205

This simulation in rate is generally of the order of 103 to lo4 over that of the reduction of free enzyme by the pyridine nucleotide. In a few cases an increased rate of reaction of the reduced enzyme with 0, has also been reported when the substrate is complexed to the enzyme. The effector role has been corroborated by the finding of many examples of substrate analogs which increase the rate of reduction of the enzymebound flavin by pyridine nucleotide but do not serve as hydroxylatable substrates and remain unchanged during the reaction. I n these cases the effector role is displayed by an increased rate of the reduced pyridine nucleotide-oxygen reductase activity. The reduction product of 0, in these cases is H,O, rather than H,O, the normal product with hydroxylatable substrates. Thus an increased rate of 0, consumption and reduced pyridine nucleotide oxidation, coupled with the production of H,O,, is a diagnostic test for nonsubstrate effectors. Some effectors have been found which combine both substrate and nonsubstrate properties, i.e., they may be hydroxylated but not quantitatively, also leading to increased pyridine nucleotide oxidation with H,O, production. I n nearly all cases it has been shown that the combination of the oxidized enzyme and the effector (be it a substrate or a nonsubstrate) results in marked changes in the physical properties of the enzymes (absorption, fluorescence, and circular dichroism spectra). Such changes have permitted the ready determination of the dissociation constants for the cornplexes, which are in all cases one to one complexes. While not so much attention has been paid to this aspect, it is also clear that complexing of the reduced enzyme with substrate also results in similarly detectable spectral changes. It is often found that complexing the oxidized enzyme with the substrate leads to a markedly decreased efficiency of the photochemical reduction of the enzyme flavin by EDTA ; hence, in experiments where the enzyme is photochemically reduced it is the usual practice to add the substrate anaerobically after reduction. The effector role of the substrate may be expected t o play a n important role in the metabolic control of the cell. If these flavoenzymes could be rapidly reduced by NADH or NADPH in the absence of substrate, pyridine nucleotide oxidation would be continuously fast and uncontrolled. The fact that rapid reaction between reduced pyridine nucleotide and enzyme flavin is permitted only when the substrate to be hydroxylated is present thus provides an elegant control mechanism of pyridine nucleotide oxidation. While the physiological reductant in all cases is either NADH or NADPH it is important from a mechanistic viewpoint to state that hydroxylation reactions can be accomplished with these enzymes using nonphysiological reductants. For the oxygenation reaction the important

206

VINCENT MASSEY AND PETER HEMMERICH

thing is to have a complex of reduced enzyme flavin and the substrate. This can be accomplished also by reducing the enzyme with chemical reductants such as dithionite, or photochemically with EDTA. Another feature of the external flavoprotein hydroxylases as a class appears to be emerging, although there are not yet sufficient examples to make it a firm generalization. When these enzymes, in their reduced form and complexed with their specific substrates, are mixed with 0, there appears very rapidly an intermediate with distinctive spectral characteristics which can be ascribed to that of a peroxydihydroflavin, i.e., a covalent adduct of molecular oxygen and reduced flavin. It is this species which is undoubtedly responsible for the insertion of one atom of oxygen into the substrate to form the hydroxylated product. The role of such adducts will be considered in more detail in the appropriate sections following and in the final section dealing with model studies.

HYDROXYLASE A. SALICYLATE Salicylate hydroxylase was the second flavoprotein monooxygenase to be discovered and the first example of the group of external monooxygenases (48). It was first purified by Katagiri and co-workers from Pseudomonas putida (49,50) and shown to have a molecular weight of 57,000 with one molecule of FAD per molecule of protein. It was also shown to contain no significant amounts of metals. By the use of 1 8 0 2 it was shown (51) that the reaction catalyzed by the enzyme is + K

O

H

NADH

+

H++

180,-

a’:+ NAD’

f

CO,

+

H,”O

The enzyme is thus unique among this group in that it is the only one which simultaneously with introducing a new hydroxyl function also results in the elimination of CO,. Recently, White-Stevens and Kamin (52-54) have reported on a salicylate hydroxylase isolated from a soil micoorganism which they obtained by enrichment culture. This enzyme differs significantly in many of its 48. M. Katagiri, S. Yamamoto, and 0. Hayaishi, JBC 237, PC2413 (1962). 49. S. Yamamoto, M. Katagiri, H. Maeno, and 0. Hayaishi, JBC 240, 3408 (1965). 50. S. Takemori, H., Yasuda, K . Mihara, K. Suzuki, and M. Katagiri, BBA 191, 58 (1969). 51. M. Katagiri, H. Maeno, S. Yamamoto, 0. Hayaishi, T. Kitao, and S. Oae, JBC 240, 3414 (1965). 52. R. H.White-Stevens and H. Kamin, BBRC 38, 882 (1970). 53. R. H. White-Stevens and H. Kamin, JBC 247, 2358 (1972). 54. R. H. White-Stevens, H. Kamin, and Q. H. Gibson, JBC 247, 2371 (1972).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

207

properties from that isolated from Pseudomonas putida. Chief among these differences are its molecular weight of 91,000 and its content of two molecules of FAD and two subunits of molecular weight 46,000. This enzyme provided the first example of a nonsubstrate effector; benzoate was found to greatly stimulate the NADH-0, reductase activity (producing H,O,), the benzoate not being hydroxylated but serving as a substratelike effector (52,53).It was originally thought that the P. putida enzyme did not exhibit this property ( 4 9 ) ,but more recent studies have shown that a t high concentrations benzoate does indeed act as a nonsubstrate effector ( 5 5 ) . The Pseudomonas putida enzyme is quite specific in its requirement for NADH as external reductant; NADPH is ineffective (49).In addition to salicylate, the following hydroxybenzoates were reported as substrates: 2,3-dihydroxybenzoate, 2,4-dihydroxybenzoate, 2,5-dihydroxybenzoate, 2,6-dihydroxybenzoate, p-amino salicylate, 1-hydroxy-2-naphthoate ( 4 9 ) , and 3-methyl salicylate ( 5 0 ) . However, of this series only 3-methyl salicylate was shown by product analysis to be hydroxylated ; the remaining compounds were tested merely by stimulation of NADH-0, reductase activity and so could be either true substrates or nonsubstrate effectors. On the other hand, White-Stevens and Kamin have made this distinction with their enzyme ( 5 3 ) .I n addition to benzoate, they found that o-nitrobenzoate, rn-hydroxybenzoate, p-hydroxybenzoate, and salicylamide were nonsubstrate effectors. A number of substituted salicylates were found to act both as hydroxylatable substrates and nonsubstrate effectors. This list includes p-amino salicylate, 3-methyl salicylate, 2,3-, 2,4-, 2,5-, and 2,6-dihydroxybenzoates ( 5 3 ) . WhiteStevens and Kamin have also reported that their enzyme will utilize NADPH as electron donor with the same V,,, value as NADH; however, the K , value for NADPH is an order of magnitude higher than that for NADH (53). On mixing with salicylate, a marked perturbation of the absorption spectrum of salicylate hydroxylase is obtained ( 5 0 ) . Titration experiments using this property established that a 1:1 complex of enzyme and M at salicylate was formed, with a dissociation constant of 3.5 X pH 7.0 and room temperature. Spectral changes were also observed in the complexing of the enzyme with substituted salicylates. Similarly, complex formation of the holoenzyme with salicylate was shown by fluorescence studies, where decreases in the fluorescence of tryptophan residues of the protein and of salicylate were detected on binding salicylate, 55. M. Katagiri, S. Taltemori, M. Nakamura, and T. Nakamura, in “Oxidases and

Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 163. Univ. Park Press, Baltimore, Maryland, 1973.

208

VINCENT MASSEY AND PETER HEMMERICH

and where increases in the flavin fluorescence of the enzyme were also M , in excellent followed ( 5 6 ) .These results provided a Kd of 3.2 X agreement with that calculated from absorbance studies. It was also shown that dramatic fluorescence changes result on mixing apoprotein with salicylate and various substituted salicylates. The dissociation constant for the apoprotein-salicylate complex (1.8 X M ) was even lower than that for the holoenzyme-salicylate complex. The fluorescence technique was also used to determine the dissociation constant for FAD ; a value of 4.5 X M was found. It was also found that NADH and NADPH formed complexes with the apoenzyme with Kd values of 1.1 X M and 1.5 X M , respectively. Product analysis from reactions involving stoichiometric amounts of enzyme demonstrated in a very clear fashion that the hydroxylation reaction is a result of reaction of the reduced enzyme-salicylate complex with 0,. The role of NADH was shown simply to be the reduction of the enzyme-bound FAD ; in these stoichiometric studies it could be replaced by dithionite or by photochemical reduction of the flavin with EDTA. The production of catechol was independent of the order of addition of reducing agent and salicylate, but was dependent upon 0, being added last (50,57). A reaction pathway for the catalytic reaction has been proposed on the basis of these results and rapid reaction studies. Takemori et a2. (58) have recently reported in detail on their rapid reaction studies. This publication has clarified a number of incompletely documented preliminary communications. The catalytic pathway is described by the following reactions: E-FAD

+

kon = 1.8

X

lo7 M-'sec-'

salicylate

E FAD salicylate

koff = 62 sec-'

-

E FAD. salicylate

+ NADH + H+

E * FADH,. salicylate

+ 4

b e d = 230 sec-'

XI

&OX

P

x

,

= 21 sec-'

salicylate - E - FADH,. + NAD'

-

E FAD t catechol CO,

+

H,O

56. K. Suauki, S. Takemori, and M. Katagiri, BBA 191,77 (1969). 57. S. Takemori, H. Yasuda, K. Mihara, K. Suauki, and M. Katagiri, BBA 191, 69 (1969).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

209

The rate-limiting step in catalysis has been found to be in the reoxidation of the reduced enzyme-salicylate complex with 0,, which was found to reach a limiting value of 21 sec-I as the 0, concentration was increased. This is identical with the value of V,,, found in the catalytic reaction. The finding of a limiting rate in flavin reoxidation indicates the formation of an intermediate X,, which may well be a complex (or compound) between the reduced flavin and 0,. Unfortunately, no spectral characterization of this intermediate was attempted; however, it is clear from the results that i t has comparatively low absorbance a t 450 nm and so may be the same type of intermediate found with p-hydroxybenzoate hydroxylase, melilotate hydroxylase, and phenol hydroxylase (see later sections on the individual enzymes). Similarly, an intermediate, XI, was detected in the reduction of the oxidized enzyme-salicylate complex by NADH; this intermediate was converted to the reduced enzyme-salicylate complex a t a rate of 230 sec-I. By pulsed-flow studies the rate of combination of oxidized enzyme with salicylate was estimated to be 1.8 X lo7 M-l sec-l. Using the value of X d of this complex nf 3.5 X M , obtained from static titration experiments, the off velocity constant was therefore calculated as 62 sec-l. Fluorescence quenching experiments were used to determine the K a value of binding of salicylate to reduced enzyme; a value of 1.7 x 10-5 M was found. Thus, salicylate binds more weakly to reduced enzyme than to the oxidized form by a factor of approximately 5-fold. Although a complete study was not made, it was evident from rapid reaction studies that salicylate binds much more slowly, by a factor of approximately 2000, to the reduced enzyme than to the oxidized form. This result rules out the possibility that reduction of enzyme by NADH might precede salicylate binding; salicylate binds to reduced enzyme at a rate only one-seventieth of that required to sustain the catalytic velocity of 21 sec-'. It was further shown in these studies that NADH reacts with enzyme in the absence of salicylate a t a comparatively slow rate and in an apparently second-order fashion, k:ed = 2.5 x lo3 M-l sec-'. At the standard concentration of NADH employed in the catalytic assays of 3X M , this reduction rate is some 2000 times slower than that of the enzyme-salicylate complex. It was also found that when the oxidized enzyme was complexed with benzoate, a nonsubstrate effector, rate stimulation of the reduction of the enzyme flavin by NADH was similar to that with salicylate. When uncomplexed reduced enzyme was mixed with 0, the reaction appeared to be second order with a rate constant of 2.9 X lo4 M-' sec-I. Complexing with benzoate did little to change this pattern; a rate constant of 4.5 x lo4 M-l sec-1 was found. These results should be contrasted

210

VINCENT MASSEY AND PETER HEMMERICH

to the limiting first-order rate of 21 sec-l following a rapid equlibrium described for the reduced enzyme salicylate complex. White-Stevens et al. (54) have also reported rapid reaction studies with their enzyme. As with the P. putida enzyme, marked spectral perturbations were observed on mixing the oxidized enzyme with substrate or nonsubstrate effectors, permitting the evaluation of dissociation constants for these complexes. I n general, the same basic reaction scheme as that of Takemori et al. (58) was proposed. However, there were many quantitative differences between the two enzymes. The forward and reverse constants for the formation of the oxidized enzyme-salicylate complex were determined independently as 9 X lo7 M-I sec-I and 130 sec-l, yielding a Kd value of 1.5 X M . This compares favorably with the Kd of 2-5 X M estimated from static titration experiments. The reduction by NADH of the E-FAD-salicylate complex was found t o be a multistep M ) followed process: the formation of a ternary complex (& = by two first-order reactions with rate constants of 150-600 sec-l and 42 sec-l. No long wavelength absorbance changes were found to be associated with reduction, and hence the two first-order processes were ascribed to formation of fully reduced flavin (the fast step) followed by an isomerization reaction (the slow step). The reaction of molecular oxygen with the reduced enzyme was also studied. In marked distinction to the results obtained with the P. putida enzyme, no limiting first-order rate was detected when salicylate was present. With free reduced enzyme, or in the presence of salicylate, benzoate, p-hydroxysalicylate, or p-aminosalicylate, the rate of reaction was the same and describable by a second-order rate constant of 1.7 X lo4 M-l 6ec-I. At atmospheric oxygen equilibration, this corresponds to a pseudo-first-order rate constant of 23 sec-l. While this rate constant is sufficient to account for the catalytic V,, with salicylate as substrate, it appears inadequate to account for the 3.6-fold greater rate found catalytically with p-hydroxysalicylate. The finding of a constant reoxidation rate led White-Stevens et al. (54) to propose the formation of a “nascent H20,” bound to enzyme, which could either react further with bound substrate to yield H,O and hydroxylated product or to dissociate yielding H,O, and unmodified effector. Flashner and Massey (10) have proposed a structure for the “nascent H,O,”; thus it could be a peroxydihydroflavin intermediate as has been observed with other flavoprotein hydroxylases, and that this was not detected experimentally possibly because the reduced enzyme was not snturated with substrate. This objection now seems to be removed by the finding with the P. putida enzyme that salicylate, while bound less tightly 58. S. Takemori, M. Nakamura, K . Suauki, M. Katagiri, and T. Nakamura, BBA 284, 382 (1972).

4. FLAVIN

21 1

AND PTFBIDINE MONOOXYGENASES

to reduced enzyme than to oxidized, has a K d value which is only five times greater for reduced than oxidized enzyme. If the same sort of relationship applies with the White-Stevens and Kamin enzyme, the concentrations of salicylate used should have been sufficient to ensure that most of the enzyme was complexed with substrate. Clearly more work is required to elucidate this problem. The finding of a limiting rate of reoxidation with the P. putida enzyme offers the clearest possibility of determining the nature of the oxygenating species since rapid reaction studies could reveal whether the intermediate determining the rate-limiting reaction has the spectrum of a peroxydihydroflavin.

B. p-HYDROXYBENZOATE HYDROXYLASE p-Hydroxybenzoate hydroxylase enzyme has been obtained in crystalline form from four different species of Pseudomonas: P. desmolytica (59), P. putida A 3.12 (60),P. putida M-6 (61), and P. fluorescens (62). In all cases the enzyme was obtained from p-hydroxybenzoate-adapted cells. The enzymes from these sources show great similarities to each other in terms of the strict requirement for NADPH as physiological external reductant, the Michaelis constant for p-hydroxybenzoate, excess substrate inhibition, and the content of 1 mole of FAD per mole of protein. The molecular weight has been estimated a t 65,000 for the P. fluorexens enzyme (62), 68,000 for the P. desmolytica enzyme (59), and 83,000-93,000 for the P. putida enzyme (60,sl). An important difference, however, has been found in the stability of the enzymes. The crystalline preparations from both P. putida species are extremely unstable requiring addition of p-hydroxybenzoate, EDTA, and a small molecular weight thiol to protect against inactivation (60,61). On the other hand, the enzyme from P. fluorescens is stable for months, even in the absence of stabilizing agents, greatly facilitating experimental work ( 6 2 ) . Working with the P. desmolytica enzyme, Yano et al. (63) demonstrated the following stoichiometry for the catalytic reaction: COOH

COOH I

I

+ NADPH + H' + Oz-

QOH

f

NADP'

f

H,O

I

OH

59. 60. 61. 62. 63.

OH

K. Yano, N. Higashi, and K. Arima, BBRC 34, 1 (1969). K. Hosokawa and R. Y. Stanier, JBC 241, 2453 (1966). B. Hesp, M. Calvin, and K. Hosokawa, JBC 244, 5644 (1969). L. G. Howell, T. Spector, and V. Massey, JBC 247,4340 (1972). K. Yano, N. Higashi, S. Nakamura, and K. Arima, BBRC 34, 277 (1969).

212

VINCENT MASSEY AND PETER HEMMERICH

By spectral titration experiments stoichiometric binding of p-hydroxybenzoate to the holoenzyme was shown (63). Convincing evidence was also given that enzyme-bound FADH, was the direct electron donor to 0, in the oxygenation reaction ; amounts of 3,4-dihydroxybenzoate were found stoichiometric with the amount of E .FADH, taken, independently of whether the latter had been produced by NADPH or dithionite. I n anaerobic stopped-flow experiments performed with enzyme containing less than a stoichiometric amount of p-hydroxybenzoate, reduction by NADPH occurred in a distinctIy biphasic fashion, related to the mole ratio of p-hydroxybenzoate to enzyme. The value for k r e d for the fast reaction was 96 sec-', some lo4 times larger than that for the slow reaction. These results demonstrate the activator or effector role of p-hydroxybenzoate and that the species reacting in catalysis with NADPH is the oxidized enzyme-p-hydroxybenzoate complex. With enzyme isolated from P. fluorescens, Howell et al. (62) have also demonstrated a perturbation of the flavin absorption spectrum by p-hydroxybenzoate. A K d of 2.9 X M was calculated from such spectral perturbations. Addition of p-hydroxybenzoate also resulted in a 75% decrease in the flavin fluorescence of the enzyme. The Kd and 1 :1 stoichiometry of binding from fluorescence titrations were similar to those found spectrally. As with the enzyme from other species, the rate of anaerobic reduction of the enzyme flavin by NADPH is greatly enhanced in the presence of p-hydroxybenzoate. With or without p-hydroxybenzoate the reduction rate with NADPH shows saturation kinetics indicating the formation of oxidized enzyme-NADPH complexes prior to reduction; the limiting first-order rate constant was 0.41 min-l in the absence and 1.52 X lo4 min-l in the presence of p-hydroxybenzoate, representing a rate stimulation of approximately 40,000 ( 6 2 ) . Another effect of prior complexing of the enzyme with p-hydroxybenzoate is to decrease the dissociation constant of the enzyme-NADPH complex by approximately 13-fold. While it is generally considered that the effector role of the substrate in facilitating the reduction of the enzyme by NADPH must result from some conformational change in the enzyme on binding the substrate, such changes must be rather small and restricted since only minor changes have been detected in the circular dichroism spectra of the enzyme on binding either substrate or competitive inhibitors (61,64). A large number of analogs of p-hydroxybenzoate has been screened as possible substrates or inhibitors. Teng et al. (64) drew a positive correlation between the strength of inhibition and the Hammett substituent constant, U , of 64. N. Teng, G . Kotowycz, M. Calvin, and K. Hosokawa, JBC 246, 5448 (1971).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

213

the substituent on the inhibitory benzoate compound. Recently, however, Spector and Massey (65) examined a larger group of inhibitors of the P . fluorescens enzyme and found that many inhibitors deviate greatly from this type of correlation. Halogen ions, in particular C1- and I-, have been found to be inhibitors of the enzyme (66). The inhibition was found to be competitive with NADPH, and a mixture of competitive and noncompetitive with p-hydroxybenzoate. Of special interest in this study was the finding that C1binds more strongly to the enzyme-p-hydroxybenzoate complex (Ed = 8 x M ) than to the free enzyme (I& = 0.11 M ) . Several compounds as well as p-hydroxybenzoate have been found to enhance the rate of flavin reduction by NADPH and hence activate the NADPH oxidase activity. Howell and Massey showed that 6-hydroxynicotinate is an acfivator without itself being a substrate (66‘,67).This was the second example of “uncoupling” of flavin reduction from substrate hydroxylation found with the external monooxygenases ; a similar uncoupling was first described for salicylate hydroxylase (52) (see Section II1,A) and has since been shown for practically every enzyme in this class. Activation of NADPH oxidation has also been found with 2,4dihydrobenzoate, 3,4-dihydroxybenzoate, and benzoate ( 6 5 ) . Product analysis showed that 2,4-dihydroxybenzoate is a substrate effector, being hydroxylated to 2,3,4-hydroxybenzoate. On the other hand, 3,4-dihydroxybenzoate, the product of hydroxylation of p-hydroxybenzoate, is not hydroxylated further, and is therefore a nonsubstrate effector. It was originally thought that benzoate was hydroxylated to a small extent to m-hydroxybenzoate, but recent work has not substantiated this (68).Interestingly, p-amino benzoate, a strong competitive inhibitor of p-hydroxybenzoate, has also been found to be a substrate, being converted to 3-hydroxy 4-amino benzoate (69). All of the effectors mentioned above perturb the absorption and fluorescence spectra of the enzyme, permitting titration experiments to determine the various dissociation constants. Rapid reaction studies (62,70)with the P. fluorescens enzyme in the anaerobic reduction of the enzyme by NADPH in the presence of p-hydroxybenzoate or 2,4-dihydroxybenzoate have allowed the characterization of several intermediates. These intermediates appear to be of the 65. T. Spector and V. Massey, JBC 247, 4679 (1972). 66. P. J. Steennis, M. M. Cordes, J. G. H. Hilkens, and F. Muller, FEBS (Fed. Eur. Biochem. Soc.) Lett. 36,177 (1973). 67. L. G. Howell and V. Massey, BBRC 40,887 (1970). 68. S. Strickland, L. Schopfer, B. Entsch, and V. Massey, unpublished observations. 69. B. Entsch, D. P. Ballou, and V. Massey, unpublished. 70. T. Spector and V. Massey, JBC 247,5632 (1972).

214

VINCENT MASSEY AND PETER HEMMERICH

charge transfer type, exhibiting long wavelength bands and probably representing charge transfer complexes between the oxidized flavin and NADPH and between the reduced flavin and NADP'. Rapid reaction studies with the enzyme from P. desmolytica did not detect such intermediates (71), possibly indicating that with this enzyme these intermediates are kinetically less stable than with the P. fluorescens enzyme. Rapid reaction studies with the P. fluorescem enzyme have permitted the spectral characterization of oxygenated intermediates when the reduced enzyme-substrate complex is reacted with molecular oxygen. Distinctly different intermediates vere detected with p-hydroxybenzoate (72) and 2,4-dih;.droxybenzoate (70) as substrates. No oxygenated intermediates were detected with the free reduced enzyme or enzyme complexed with a nonsubstrate effector such as 6-hydroxynicotinate or 3,4-dihydroxybenzoate. These results were initially interpreted as possibly indicating a different site of addition of 0, to the dihydroflavin depending on the particular substrate complexed to the enzyme. However, more recent studies have shown that when 2,4-dihydroxybenzoate is the substrate, three different intermediates can be detected (73). The first intermediate formed, whose formation rate is directly proportional to 0, concentration, has a spectrum very similar to that formed with p-hydroxybenzoate as substrate (A,,, 370 nm, E 8000 M-l cm-I). I n the presence of 2,4-dihydroxybenzoate this intermediate is converted rapidly to a second intermediate with a maximum at 410 nm and a very high extinction coefficient, c = 13,600 M-l cm-l. The rate of formation of this intermediate is much less dependent on 0, concentration, and saturates a t high 0, concentrations, with a limiting rate constant (at pH 8.5, temperature 1.5O) of 5300 min-'. I n turn, this intermediate is converted to a third a t a rate independent of 0, concentration, lc, = 21 min-l. Finally, this intermediate disappears with the reformation of the spectrum of oxidized enzyme, with the 0,-independent rate lc, = 8.3 min-'. Figure 1 shows the spectra of the above intermediates. Studies in which the reaction of the reduced enzyme 2,4-dihydroxybenzoate complex with 0, was quenched rapidly with HC1 after finite time periods and then analyses performed for the product 2,3,4-trihydroxybenzoate revealed that oxygen atom insertion into the substrate occurs at the stage of conversion of intermediate I to intermediate 11. While the mechanistic details are still far from clear, i t is tempting r~

-

71. S. Nakamura, Y. Qgura, K. Yano, N. Higashi, and K. Arima, Biochemistry 9, 3235 (1970). 72. T. Spector and V. Massey, JBC 247, 7123 (1972). 73. B. Entsch, V. Massey, and D. P. Ballou, BBRC 57, 1018 (1974).

4.

2 15

FLAVIN AND PTERIDINE MONOOXYGENASES

I

I

I

c.-

- 12000

0.4 -

- 10000

- 8000

03 a,

E

0 C

m

n L

I

140000

.

I1

05

- 6000

0.2 -

0

In

- 4000

n

a

o.,

-Reduced enzyrn$,,

- 2000

*dihyroxybenzoate

Wavelength (nm)

FIQ.1. Intermediates in the reaction of oxygen with reduced p-hydroxybenzoate hydroxylase complexed with 2,4-dihydroxybenzoate. Data of Entsch et al. (79).

to speculate on the chemical nature of the intermediates. Intermediate I, with an absorption maximum a t 385 nm and an extinction coefficient of 8500 M-l cm-l, together with the second-order dependence on O2 concentration of its formation, would most logically represent a covalent adduct of reduced flavin and 0,. The important question of the site or sites of addition in the isoalloxazine ring system cannot be answered a t this time. However, comparison with the spectral properties of model compounds suggests that the intermediate may have the structure of a substituted C (4a)-N ( 5 )-dihydroflavin (see Section V) ; for example,

H

C(4a) - Peroxydihydroflavin

The rapid quench experiments show that oxygen transfer to the substrate is accomplished coincident with the conversion of intermediate I to intermediate 11. Hence, it is tempting to speculate that the unusual

216

VINCENT MASSEY AND PETER HEMMERICH

spectral characteristics of intermediate I1 result from a complex of the enzyme with the flavin now in a hydroxydihydroflavin form and with the product not yet in its final form; for example, this could be some dienonole-type intermediate such as

0

which should aromatize readily to yield 2,3,4-trihydroxybenzoate. Thus, intermediate 111 could represent a complex of 2,3,4-trihydroxybenzoate with the hydroxydihydroflavin form of the enzyme. The latter, by dehydration of the hydroxydihydroflavin and dissociation of the product, would then return the enzyme to the oxidized form ready for the next catalytic cycle. Extension of such studies to other substrates (69) has revealed that the sequence of intermediates described above is not unique to 2,4-dihydroxybenzoate; for example, p-aminobenzoate, previously considered to be merely a competitive inhibitor, has been found to be a substrate for the enzyme, and elicits a similar sequence of intermediates. Studies with the reduced enzyme p-hydroxybenzoate complex a t low pH values and low temperature have also permitted the positive identification of two intermediates in the reaction of this complex with 0,, with spectral characteristics similar to those of I and I11 in the reaction with 2,4-dihydroxybenzoate as substrate (69). It would thus appear likely that the following reaction scheme must be a minimal one for the oxygen reaction: EFHzArH

+

ki 0 2 -+

I

kz --*

I1

2 TI1 2 EF + ArOH + HzO

In the case of 2,4-dihydroxybenzoate and p-aminobenzoate as substrates, the various rate constants are such that three intermediates can be seen. However, if k , were slower than Ic, and kq, only intermediate I would be expected to be detected. This would appear to be the case for p-hydroxybenzoate as a substrate a t pH 8.5. If k , were slower than lc, but not slower than Ic,, then intermediates I and 111 would be expected to be detected. This would appear to be the case for p-hydroxybenzoate as substrate a t pH 6.5.

4.

217

FLAVIN AND PTERIDINE MONOOXYGENASES

C. MELILOTATE HYDROXYLASE In 1965,in the course of investigating the metabolism of coumarin by an Arthrobacter species, Levy and Frost discovered an enzyme which uses NADH and 0, to hydroxylate melilotate to 2,3-dihydroxyphenylpropionate (74): FH,CH,COOH

FH,CH&OOH

+ NAD+ + q o

+ NADH + H+ + o*-

The enzyme was subsequently purified and found to be a flavoprotein with a molecular weight of 65,000 containing one molecule of FAD per molecule protein (75). More recently, Strickland and Massey (76) have reported the isolation of melilotate hydroxylase from a Pseudomonas species. Like the Arthrobacter enzyme this was also found to contain FAD as prosthetic group (1 molar equivalent per 65,000 g protein) but differs from the Arthrobacter enzyme in having a considerably higher molecular weight (238,000-250,000) and in having four protein subunits. This enzyme has been the subject of a comprehensive kinetic study ( 7 7 ) .I n a conventional steady-state analysis, in which the concentrations of each of the three substrates, melilotate, NADH, and 0, were varied systematically, the following unique pathway was deduced: NADH E . FAD + mel.-.

kl

/FAD

E,

k,

E-FADH, k4

me 1

A .~

/FADH,

k,

E

'me1

+ NAD+

The validity of this reaction scheme was demonstrated by rapid reaction studies in which changes in the enzyme itself were monitored by stoppedflow spectrophotometry and fluorimetry, which permitted determination 74. C. C. Levy and P. Frost, JBC 241, (1967). 75. C. C. Levy, JBC 242, 747 (1968). 76. S.Strickland and V. Massey, JBC 248, 2944 (1973). 77. S. Strickland and V. Massey, JBC 248,2953 (1973).

218

VINCENT MASSEY AND PETER HEMMERICH

of most of the individual rate constants. The formation of an oxidized enzyme-melilotate complex was readily detected by perturbation of the flavin absorption spectrum of the enzyme ; titration experiments showed that under standard conditions (pH 7.3, l o ) the Kd for this complex was 3.8 x M . That the reaction pathway follows the ordered sequence shown above was demonstrated by the fact that enzyme in the absence of melilotate is reduced very slowly by NADH (limiting rate 1.4 min-l extrapolated to infinite NADH concentration). This is slower by more than two orders of magnitude than the catalytic turnover number of the enzyme (735 min-I). I n contrast the E. FAD-melilotate complex reacts extremely rapidly with NADH, in an apparently second-order reaction, k , = 1.4 x lo8 M-I min-'. Determination of the absorption spectrum showed that the product of this reaction was a ternary complex of reduced enzyme, melilotate, and NAD', which proceeded to dissociate the NAD+ a t a rate k , = 1300 min-I. The ternary complex was shown to exhibit a charge transfer absorption spectrum (with a long wavelength band centered round 750 nm) as a result of interaction between EaFADH, (donor) and NAD' (acceptor). The same ternary complex could be formed in static equilibrium experiments in which the E.FADH,-melilotate complex was titrated anaerobically with NAD'. These titrations permitted the determination of the dissociation constant (Kd = 1.45 X M ) and hence the calculation of k,. Stopped-flow turnover data (in which all three substrates are mixed with the enzyme) also indicate that NAD+ dissociates from the ternary complex prior to reaction with 0, (77). As well as the dramatic enhancement of the reduction rate, complexing the enzyme with melilotate also results in a significant increase in the rate of reaction of the reduced flavin with 0,. I n the absence of melilotate the rate is 9.7 X lo5 M-l min-l; in the presence of saturating melilotate this is increased some 16fold to 1.6 x lo7 M-l min-'. Of particular interest is the detection of an oxygenated flavin intermediate in the reaction of the reduced enzymemelilotate complex with 02. This intermediate has a spectrum similar to that of intermediate I found with p-hydroxybenzoate hydroxylase, as described in the preceding section. The rate of appearance of the intermediate ( k , = 1.6 X lo7 M-l min-l is directly dependent on 0, concentration; its subsequent breakdown to yield oxidized enzyme and products is independent of 0, concentration and proceeds a t a rate k , = 1400 min-l. These independently measured rate constants were then used to predict values for the Michaelis constants of the substrates and the maximum catalytic velocity of the hydroxylation reaction, using kinetic equivalents appropriate for the reaction mechanism shown. Table I1 lists the individual rate constants determined from stopped-flow and equilibrium studies, the predicted Vn,,, and K , values, and those actually observed in the

4. FLAVIN

219

AND PTERIDINE MONOOXYGENASES

TABLE I1 KINETICANALYSISOF MELILOTATE HYDROXYLASE (pH 7.3, 1”) Step ki k2

ka kr

Rate constant

Method of determination

5 . 7 X lo8 M-I min-1 2 . 2 X l o 4 min-1 1 . 4 X 108 M-1 min-1

From Vmsxand K m (mel) From k, and Kd (mel) Stopped flow Stopped flow Stopped flow From k5 and Kd (NADf) Stopped flow Stopped flow Stopped flow

0 1.3

k7

x 103 min-1 9 . 0 X lo5 M-I min-1 1.6 X l o 7 M-l min-1

ks ko k I o,ki 1,k I 2

1 . 4 X 103 min-1 Not determined

ks ka

Kinetic constant

0

Kinetic equivalent

Observed steady-state value 735 min-1

K , (NADH)

+

(k4 k5) kaks . 40

4.7

x 10V M

Predicted from rate constants 680 min-l

4.8 X

M

steady-state analysis. The predicted and observed values are in remarkably good agreement and provide very strong evidence for the validity of the proposed mechanism. The intramolecular migration of ring substituents during hydroxylation of aromatic substrates (the NIH shift) is a phenomenon which has been investigated extensively by Jerina, Daly, and co-workers (78). Their incisive experiments have led to the mechanistic conclusion that hydroxylations catalyzed by the liver microsomal P 450 system proceed via an arene oxide intermediate (79). It is the subsequent protonation of this arene oxide followed by rearrangement which leads to the final aromatic 78. D. M. Jerina and J. W. Daly, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 143. Univ. Park Press, Baltimore, Maryland, 1973. 79. D. M. Jerina, J. W. Daly, B. Witkop, P. Zaltzman-Nirenberg, and S. Udenfriend, Biochemistry 9, 147 (1969).

220

VINCENT MASSEY AND PETER HEMMERICH

hydroxylated product. One of the most important questions concerning these results is whether the arene oxide pathway is an obligatory route for aromatic hydroxylations occurring via reductive activation of molecular oxygen. Confirming the presence or absence of the NIH shift in flavoprotein-catalyzed hydroxylations would thus lead to important extrapolations concerning similarities in the mechanism of hydroxylations effected by different biological systems. In a recent study, Strickland et at. (80) have employed combined gas chromatography-mass spectrometry to analyze the reaction products when 3,5-dideuteromelilotate was employed as a substrate with melilotate hydroxylase. No retention of deuterium indicative of the NIH shift could be detected (limits of detection 0.5%). Unfortunately, these results cannot rule out definitely the occurrence of the arene oxide pathway since it has been shown that substituents on phenols generally show little or no migration during aromatic hydroxylations with systems known to give the NIH shift with other substrates (81).This is presumed to result from a catenoid intermediate being stabilized effectively through loss of a proton from the phenolic-OH to yield a stable dienone without migration of a substituent (Scheme 1). R

R

R

~

&

d

R

dienonc stabilization

*He

0

,

1

D

-Y@orD"

-De

NI H SHIFT

1

NO NIH SHIFT

SCHEME 1 80. S. Strickland, L. Schopfer, and V. Maasey, Biochemistry 14,2230 (1975). 81. J. W. Daly, G. Guroff, 5. Udenfriend, and B. Witkop, ABB 122, 218 (1967).

4. FLAVIN AND PTERIDINE MONOOXYGENASES

221

While such stabilization effects would make it difficult to interpret a negative result, it should be pointed out that small amounts of migration, 1-5%, have been observed during hydroxylation of phenols (81). Thus, it is possible from the complete absence of migration observed with melilotate hydroxylase that the arene oxide pathway is not operational in this flavoprotein-catalyzed reaction. In the above study it was also found that use of 3,5-dideuteromelilotate instead of the normal protium form led to no changes in the rate of formation ( k , ) of the oxygenated intermediate or in its subsequent breakdown (k9).As discussed in the previous section, recent work with a similar flavoprotein p-hydroxybenzoate hydroxylase (73) has demonstrated the existence of a t least three intermediate steps when reduced enzyme-substrate complex is reacted with molecular oxygen. This suggests the possibility that similar intermediates exist with melilotate hydroxylase which are however invisible for kinetic reasons. It could be one of these latter intermediates which is responsible for the fission of the carbon-hydrogen bond, thus accounting for the lack of an observed isotope effect.

D. PHENOL HYDROXYLASE Recently, Neujahr and Gaal (82) have reported the purification of a phenol hydroxylase from a yeast, Trichosporon cutaneum. This yeast strain was induced to produce the enzyme in response to growth on phenol or resorcinol as the major carbon source. The enzyme was obtained as a protein homogeneous by disc gel electrophoresis. It has a molecular weight of 148,000and was reported to contain one molecule of FAD per molecule of protein. However, this value was probably low, as a result of some loss of FAD from the enzyme during purification; more recent analyses of 17 different preparations of the enzyme gave an average FAD content of 1.74 molecules per 148,000molecular weight. Sodium dodecyl sulfate (SDS) gel electrophoresis also indicates the presence of two subunits of molecular weight approximately 76,000 (Dr. H. Y. Neujahr, personal communication). The enzyme has an unusually broad substrate specificity, although the requirement for NADPH as external electron donor is strict. Phenol is hydroxylated to yield catechol. Catechol is also hydroxylated to yield

82. H. Y. Neujahr and A. Gaal, Eur. J . Bbchem. 35, 386 (1973).

222

VINCENT MASSEY AND PETER HEMMERICH

pyrogallol (1,2,3-trihydroxybenzene). Other dihydroxybenzenes are also substrates ; resorcinol (1,3-dihydroxybenzene) and quinol (1,4-dihydroxybenzene) are both hydroxylated to yield hydroxyquinol ( 1,2,4-trihydroxybenzene) . Various substituted phenols also act as substrates, with rates between 10 and 100% that of phenol. Thus, 2-F-, 3-F-, and 4-Fphenols; 2-C1-, 3-C1-, and 4-Cl-phenols ; 2-methyl, 3-methyl, and 4methyl phenols; and 2-amino, 3-amino, and 4-amino phenols are substrates. While complete stoichiometric studies have not been carried out in all cases, it is evident that substantial hydroxylation of these substrates occurred. Thus, the possibility exists that the enhanced rate of NADPH oxidation with these substances may in part result from their also acting as nonsubstrate effectors (cf. Section II1,A on salicylate hydroxylase) . The enzyme is susceptible to pronounced inhibition by chloride ions,

0.25

1

0.20

0.15 E

m

n

5

2

0.10

0.05

I

350

LOO Anrn L50

500

FIG.2. Phenol hydroxylase, 2.13 x 10.' M with respect, to enzyme-bound flavin, in 0.1 M phosphate pH 7.6, containing 10 mM dithiothreitol, 15 mM EDTA, and 5 mM phenol, was reduced photochemically in an atmosphere of N, and mixed in the stopped-flow apparatus. with an equal volume of 02-saturated buffer mixture. The absorbance changes (path length 2 crn) were followed with time a t the wavelengths shown 6-10 nm intervals from 330 to 550 nrn). The biphasic changes in absorbance were analyzed to determine the spectrum of the intermediate. Temperature of observation, 2" (V. Massey and H. Y. Neujahr, unpublished results).

4.

223

FLAVIN AND PTERIDINE MONOOXYGENASES

this inhibition effect being much more marked on the acid limb of the pH-activity profile than on the alkaline limb. The catalytic reaction is also inhibited by high concentrations of phenol. Metal chelators are without effect on activity; however, a requirement for intact thiol residue (s) is indicated by the susceptibility of the enzyme t o inhibition by heavy metal ions and p-mercuribenzoate ( 82) . Like the other flavoprotein external monooxygenases so far investigated, phenol hydroxylase also shows distinctive changes in the visible absorption spectrum on complexing with its phenolic substrates (82). Preliminary studies also indicate that with such complexes the rate of reduction of the enzyme flavin by NADPH is considerably faster than with uncomplexed enzyme (H. Y. Neujahr and V. Massey, unpublished results). Preliminary rapid reaction studies have shown that phenol hydroxylase, like p-hydroxybenzoate hydroxylase and melilotate hydroxylase (see previous sections), forms a transient peroxydihydroflavin species when the reduced enzyme-phenol complex is mixed with oxygen. The spectrum of this species, together with those of the complexes of phenol with the oxidized and reduced enzyme, is shown in Fig. 2.

E. ORCINOLHYDROXYLASE Orcinol hydroxylase has been isolated in crystalline form from Pseudomonas putida and shown to be a flavoprotein consisting of a single polypeptide chain of molecular weight 60,000-70,000 and containing one molecule of FAD per molecule of protein (83).The reaction catalyzed by the enzyme is

+ HO

NADH

+

-

H+ + 0,

+ NAD+ + H,O

HO

Orcinol

2,3,5-Trihydroxytoluene

The weak NADH oxidase activity of the enzyme is stimulated by resorcino1 and m-cresol. Product analysis showed that these compounds were functioning chiefly as nonsubstrate effectors. When resorcinol is used some hydroxylated product (hydroxyquinol) is formed but not in sufficient quantities to account for the NADH and O2 consumption. Furthermore, addition of catalase to the assay mixture results in the return of 40% 83. Y . Ohta and D. W. Ribbons, FEBS (Fed. Eur. Bioehem. Soc.) Lett. 11, 189 (1970).

224

VINCENT MASSEY AND PETER HEMMERICH

of the 0, consumed. With m-cresol, no hydroxylation occurs, and catalase returns 50% of the consumed 0,. Addition of catalase has no effect on the NADH oxidation in the presence of either substrate analog. These results show that orcinol is the only true substrate with its hydroxylation being tightly coupled to NADH oxidation, while resorcinol is hydroxylated to only a limited extent and m-cresol not a t all. Orcinol hydroxylase can utilize both NADH and NADPH as well as reduced 3-acetylpyridine nucleotide as electron donors, although NADH is the best donor (84). Ribbons et al. have studied the stereospecificity of hydride transfer from NADH to the enzyme-bound FAD using 4Rand 4S-[SH]-NADH (84). The pro-R protium (A side) of NADH is stereospecifically transferred when orcinol is the substrate. With m-cresol as effector, transfer of hydride rather than tritide appears to be preferred from either side of the reduced pyridine nucleotide, while with resorcinol the specificity for the pro-R tritium is less than with orcinol yet more than with m-cresol.

F.

m-HYDROXYBENZOATE-6-HYDROXYLASE

m-Hydroxybenzoate-6-hydroxylaseis an inducible enzyme which has been purified from Pseudomom aeruginosa. It catalyzes the initial hydroxylation reaction in the gentisate pathway for the metabolism of mhydroxybenzoate (85): COOH

rn-Hydroxybenzoic acid

COOH

Gentisic acid

It is a flavoprotein using either NADH or NADPH as electron donor. As with the other external flavoprotein monooxygenases previously discussed, reduced pyridine nucleotide oxidation is greatly facilitated by the presence of the substrate effector, m-hydroxybenzoate (D. W. Ribbons, personal communication). The unusual feature of the reaction catalyzed by this enzyme is the site of hydroxylation in the aromatic ring. With all other flavoprotein hydroxylases, the new hydroxyl function is introduced in the position 84. D. W. Ribbons, Y. Ohta, and I. J. Higgins, in "The Molecular Basis of Electron Transport" (J. Schultz and B . F. Cameron, eds.), p. 251. Academic Press, New York, 1972. 86. E. E. Groseclose and D. W. Ribbons, Bacteriol. Proc. p. 273 (1972).

4. FLAVIN

225

AND PTERIDINE MONOOXYGENASES

ortho to the existing hydroxyl group. This feature has been used by Hamilton (86) t o propose a mechanism for flavoprotein-catalyzed hydroxylation reactions involving a 4a-peroxydihydroflavin which is converted to a ring-opened form, the latter then being attacked by the nucleophilic center ortho to the phenolic group of the substrate. While this mechanism lacked experimental basis, it would seem unable to accommodate the para insertion of a hydroxyl function catalyzed by m-hydroxybenzoate-6-hydroxylase.

G.

m-HYDROXYBENZOATE-4-HYDROXYLASE

Prema Kumar et al. (87) have reported the partial purification from Aspergillus niger of a m-hydroxybenzoate hydroxylase, utilizing NADPH as electron donor and producing protocatechuate: COOH

COOH

OH

The prosthetic group was found to be FAD. The same workers (88) have shown that the above reaction can be completely inhibited by reasonably low levels of superoxide disrnutase. This result is surprising in view of the lack of inhibition by superoxide dismutase of other flavoprotein hydroxylases (4,SS). The positive effect of superoxide dismutase found with this enzyme indicates the possibility of involvement of the superoxide in the hydroxylation reaction. This possibility will be conanion, 02-, sidered further in the last section. m-Hydroxybenzoate-4-hydroxylase has also been obtained recently fr0m.P. testosteroni. In addition, this enzyme uses NADPH as the preferred electron donor, although NADH will also function less efficiently (D. W. Ribbons, personal communication).

H. IMIDAZOLYLACETATE MONOOXYGENASE Imidazolylacetate monooxygenase has been isolated from a pseudomonad species and forms part of the histidine catabolic pathway in which 86. G. A. Hamilton, i n “Molecular Mechanisms of Oxygen Activation” (0.Hayaishi, ed.), p. 405. Academic Press, New York, 1974. 87. R. Prema Kumar, P. V. Subba Rao, N. S. Sreeleela, and C. S. Vaidyanathan, Can. J. Bioehem. 47, 825 (1969). 88. R. Prema Kumar, S. D. Ravindranath, C. S. Vaidyanathan and N. Appaji Rao, BBRC 49, 1422 (1972).

226

VINCENT MASSEY AND PETER HEMMERICH

imidazalonylacetate is converted to aspartic acid (89). The reaction catalyzed by the enzyme is H HC =C I 1 HN,C+N

,CH,COOH

%

+NADH+H++O,-

H

Imidazolylacetate

C

&

,

~

~

Z

1 I HKC’,N

~ f

~

~

NAD’

+ H,O

H

Imidazolonylacetate

The monooxygenase nature of the enzyme was shown by Rothberg and Hayaishi with l8O experiments (90). Crystallization of imidazolylacetate monoxygenase (89,91) revealed the flavoprotein nature of the enzyme with FAD as the only prosthetic group. The molecular weight was estimated to be between 87,000 and 90,000, with one molecule of FAD per molecule of protein. NADH is the preferred electron donor, although NADPH can also function in a less effective manner, Okamoto et al. have studied the role of thiol groups in the enzyme (92). They found that two thiol groups could be titrated in the native enzyme by silver nitrate or by p-mercuribenzoate. I n the presence of imidazolylacetate only one thiol group could be titrated and the presence of the substrate protected against inactivation by the mercurial. While the monooxygenase activity was lost completely on reaction with mercurial (in the absence of substrate) the weak NADH oxidase activity was not; in fact, a two- to threefold stimulation of this activity (which results in H,O, formation) was observed. In addition to imidazolylacetate, imidazolylpropionate and imidazolyllactate were found to stimulate the rate of NADH oxidation; however, in the absence of product analysis it is not clear whether these compounds function as true substrates or nonsubstrate effectors (91). In common with other external flavoprotein monooxygenases, imidazolylacetate monooxygenase is not inhibited by metal chelators nor does it contain any significant quantities of trace metals. It is unusual among this group of enzymes however inasmuch as the addition of substrate is without effect on the visible absorption spectrum and on the CD and ORD spectra (911.

I. BACTERIAL LUCIFERASE The phenomenon of bioluminescence emission exhibited by many bacteria has attracted much experimental attention. No attempt will be made 89. Y. Maki, S. Yamamoto, M. Nosaki, and 0. Hayaishi, BBRC 25,609 (1966). 90.S. Rothberg and 0. Hayaishi, JBC 229, 897 (1957). 91. Y . Maki, S. Yamamoto, M. Nosaki, and 0. Hayaishi, JBC 244,2942 (1969). 92. H. Okamoto, M. Noraki, and 0. Hayaishi, BBRC 32, 30 (1968).

4.

227

FLAVIN AND PTERIDINE MONOOXYGENASES

here to review the rather extensive literature on the subject or to consider the possible mechanisms of the light emitting process. The reader is referred to a recent review article for such information (93). It has long been known that the light emission depends on the presence with the bacterial luciferase of FMNH,, 0,, and a long-chain aldehyde (94,95). Although the aldehyde is essential for high quantum yields, its fate during the reaction remained obscure until recently. It was proposed by McElroy and Green in 1955 (96) that the aldehyde is converted to the corresponding long-chain acid since this reaction would provide sufficient energy for the emission of a quantum of light of 490 nm. Experimental support for this proposal came recently from the detection of acid production by mass spectrometry (97). Further definitive proof of acid production was obtained independently by McCapra and Hysert (98) and by Dunn et d.(99). These studies have thus established the luciferase reaction as a hydroxylation of the long-chain aldehyde: FMNHz

+ RCHO +

0

2

4

FMN

+ RCDOH + HzO

Luciferase may therefore be classified as a somewhat unusual flavoprotein monooxygenase, its unique feature being that FMNH, appears to function as a substrate rather than a prosthetic group. However, this is clearly a matter of semantics. The dissociation constant for binding of FMNH, to the enzyme from P. fischeri has been estimated as 9.7 X lo-? M ; a value of 8.0 x lo-? has been found for the so-called MAV-luciferase (100). Baldwin has recently reported that F M N also binds to the latter enzyme, although much less strongly; a Kd value of 2.4 X M was calculated (101). Although most experiments with the isolated enzyme have employed added FMNH, as the electron donor, the luminescent bacteria utilize a NADH-FMN reductase to produce the FMNH, used in the luciferase reaction (93). Thus, the combined effects of the two enzymes is formally the same as for all of the external flavoprotein monooxygenases so far discussed. 93. J. W. Hastings, Annu. Rev. Biochem. 37,597 (1968). 94. M. J. Cormier and B. L. Strehler, JACS 75, 4864 (1953). 95. B. L. Strehler, E. N. Harvey, J. J. Chang, and M. J. Cormier, Proc. Nut. Acad. Sci. U . S . 40, 10 (1954). 96. W. D. McElroy and A. A. Green, ABB 56, 240 (1955). 97. 0. Shimomura, F. H. Johnson, and Y. Kohama, Proc. Nut. Acad. Sci. U . S . 69, 2086 (1972). 98. F. MeCapra and D. W. Hysert, BBRC 52, 298 (1973). 99. D. K. Dunn, G. A. Michaliszyn, I. G. Bogacki, and E. A. Meighen, Biochemistry 12, 4911 (1973). 100. E. A. Meighen and J. W. Hastings, JBC 246, 7666 (1971). 101. T. 0. Baldwin, BBRC 57, lo00 (1974).

228

VINCENT MASSEY AND PETER HEMMERICH

From their early work employing rapid reaction spectrophotometry, Hastings and Gibson (102) concluded that a long-lived intermediate is formed on mixing luciferase, FMNH,, and 02,which reacts further, with concomitant light emission, in the presence of a long-chain aldehyde. Competing with the formation of this intermediate is the rapid reaction of free FMNH, and 0, to yield F M N and H,O,. I n a recent paper the existence and spectral characterization of the proposed enzyme intermediate was shown in a very elegant fashion (103).A mixture of luciferase, FMNH,, and 0, in 50% ethylene glycol-phosphate buffer was allowed to react at 4 O for 10 sec and then the temperature was lowered rapidly to -2OO. The mixture was then chromatographed on a Sephadex L H 20 column a t -20° to separate enzyme-bound intermediates cleanly from free FMN. The results are very nicely consistent with the following scheme:

FMN

.+1

H,O,

E

lkd

+ FMN + H,O,

+

Ikb

E FMN+ H,O +acid hu

+

The intermediate I1 isolated by low temperature chromatography was shown to have an absorption spectrum with a maximum a t 370 nm and comparatively little absorption at 450 nm. Its spectral characteristics are in fact very similar to those of the oxygenated flavin intermediates found with p-hydroxybenzoate hydroxylase and phenol hydroxylase described in the previous sections. The intermediate is stable for long periods a t -20° ; on warming in the absence of aldehyde, enzyme, FMN, and H20, are formed (pathway k d ). I n the presence of aldehyde however the characteristic light emission of the luciferase reaction (490 nm) is observed, consistent with the formation of a further intermediate I I a and breakdown to products via the pathway labeled kb. The physical isolation of an oxygenated flavin intermediate in this case, as opposed to the kinetic “isolation” with the other hydroxylases, offers possibilities of incisive mechanistic studies. Extension of this isolation technique t o other enzymes, particularly with substrates being turned over only slowly, is 102. J. W. Hastings and Q. H. Gibson, JBC 238,2537 (1963). 103. J. W. Hastings, C. Balny, C. Le Peuch, and P. Douznu, Proc. N a t . Acad. Sci. U.8. 70, 3468 (1973).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

229

clearly an attractive experimental approach to investigating further the reaction mechanisms of these enzymes.

J. MICROSOMAL AMINEOXIDASE All of the flavoprotein monooxygenases so far discussed are of microbial origin. I n recent years, Ziegler and his colleagues have reported on a monooxygenase present in liver microsomes of many vertebrate species (104-107).This enzyme appears to be a simple flavoprotein containing no significant quantities of metal ions or heme residues. The enzyme has been isolated from pig liver microsomes and is reported to have a molecular weight of approximately 500,000 (106). The minimum molecular weight per FAD prosthetic group is 71,000. Thus, the enzyme would appear to he a polymeric, aggregating species. The enzyme catalyzes the NADPH- and 0,-dependent N-oxidation of a variety of secondary and tertiary amines. The secondary amines are oxidized to the corresponding hydroxylamines and the tertiary amines to amine oxides, e.g.,

N-Methylaniline

N, N-Dimethylaniline

N - Methylpheny Lhydroxylamine

N, N-Dimethylaniline - N-oxide

The primary amines, 1-naphthylamine and 2-naphthylamine, are also oxidized but a t lower rates (107). With one substrate, dimethyloctylamine, sigmoidal kinetics are found, suggesting the existence of an effector 104. J. M. Machinist, E. W. Dehner, and D. M. Ziegler, ABB 125, 858 (1968). 105. D. M. Ziegler, D. Jollow, and D. E. Cook, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 507. Univ. Park Press, Baltimore, Maryland, 1971. 106. D. M. Ziegler and C. H. Mitchell, ABB 150, 116 (1972). 107. D. M. Ziegler, L. L. Poulson, and E. M. McKee, Xenobiotica 1, 523 (1971).

230

VINCENT MASSEY AND PETER HEMMERICH

site in addition to the catalytic site. This idea is supported by the finding of a variety of compounds, which are themselves not substrates, but which increase the rate of oxidation of true substrates (106).

K.

KYNURENINE-3-HYDROXYLASE

Kynurenine-hydroxylase has been purified partially from rat liver mitochondria (108).Okamoto et al. (109) showed that the enzyme is localized in the outer membrane of the mitochondrion. Several workers, have, as a result, used kynurenine-hydroxylase activity as a mitochondria1 marker (see, for example, 110,111) . Experiments with lsO2 have established the enzyme as a true monooxygenase (112,115): 0 a I ; CII H r

CH-COOH I NH,

+

NADPH

+

HC

+

NADP’

+

H,O

+

0,

~-Kynurenine

qC!Z,C&0 II

CH-CCOOH I

NH*

OH 3 - Hydroxy - ~-kynurenine

The enzyme can use NADH as an alternative electron donor (114). Acid ammonium sulfate treatment of the partially purified enzyme resulted in a decrease in activity to 60% the original level. Addition of FAD restored almost completely the initial activity, while FMN was ineffective (109,11$1l5). There is thus circumstantial evidence that the 108. H. Okamoto, “Methods in Enzymology,” Vol. 174, p. 460,1970. 109. H. Okamoto, S. Yamamoto, M. Nozaki, and 0. Hayaishi, BBRC 26, 309 (1967). 110. D. S. Beattie, BBRC 31, 901 (1968). 111. C. A. Schnactman and J. W. Greenawalt, J. Cell Biol. 38, 158 (1968). 112. Y. Saiton, 0. Hayaishi, and S. Rothberg, JBC 229, 921 (1957). 113. H. Okamoto, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 223. Univ. of Tokyo Press, Tokyo, 1968. 114. 0. Hayaishi and H. Okamoto, Amer. J . Clin. Nutr. 24, 805 (1971). 115. .H. Okamoto and 0. Hayaishi, BBRC 29, 394 (1967).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

23 1

enzyme is a FAD-containing flavoprotein. Whether or not FAD is the only prosthetic group will obviously have to await more extensive purification of the enzyme.

IV. Pterin-Linked Monooxygenases

A .small group of metabolically important enzymes has been shown to require a reduced unconjugated pterin cofactor as electron donor in the aromatic ring hydroxylation reactions which they catalyze. Because of the rather similar chemistry of flavins and pteridines i t is a widely considered possibility that these enzymes may function by mechanisms basically similar to those of the flavoprotein monooxygenases. While this possibility is real, there are several important distinctions which may be made between the properties of the pteridine and flavin monooxygenases and between the properties of tetrahydropteridines and dihydroflavins. 1. While the flavoprotein monooxygenases have been shown to be free of trace metal involvement, the pteridine monooxygenases all seem to contain and require protein-bound iron as an additional cofactor. 2. I n the case of the flavoprotein monooxygenases the flavin is tightly bound to the protein, and the protein is very selective in binding only FAD or FMN. With the pteridine enzymes on the other hand, the pteridine is bound only weakly to the protein, and hence can be considered more as a mobile substrate than as a prosthetic group. I n addition, a large number of unsubstituted tetrahydropteridines will function in the catalytic reaction. In this connection the pteridine monooxygenases are more akin to bacterial luciferase (see Section II1,I) than to the rest of the flavoprotein monooxygenases. 3. With the exception of bacterial luciferase, the reactions catalyzed by the flavoprotein monooxygenases involve a single enzyme. In the case of the pteridine-linked hydroxylations a t least two enzymes are involved in the catalytic reaction, the hydroxylase proper, where the tetrahydropteridine is oxidized to a dihydropteridine in the couse of the hydroxylation reaction, and a tetrahydropteridine regenerating enzyme, NADHdihydropteridine reductase. 4. The reactivity of dihydroflavins with molecular oxygen is rather high (tM < 1 sec) while that of tetrahydropteridines is rather low ( t ~ in the order of minutes). Unless there is a dramatic enhancement of this rate on binding the tetrahydropteridine to the protein, such rates would appear unlikely to be great enough to sustain catalysis.

232

VINCENT MASSEY AND PETER HEMMERICH

A. PHENYLALANINE HYDROXYLASE Phenylalanine hydroxylase was the first discovered and most widely studied enzyme of this group. Studies from several laboratories in the early 1950’s showed the conversion of phenylalanine to tyrosine in rat liver extracts supplemented with pyridine nucleotides (116,117). I n 1957 Kaufman (118) established the following stoichiometry in a partially purified system: GPhenylalanine

+ NADPH + H+ + O2+ Gtyrosine + NADPf + H20

This stoichiometry suggested that the reaction is of the monooxygenase type, i.e., that the phenolic group of tyrosine is derived from 0, rather studies; when than H,O. This conclusion was shown to be correct by 1802 the hydroxylation reaction was carried out in the presence of lRO2the phenolic group was found to be labeled, but no labeling was found when the reaction was carried out with l6OZand H2180 (119). The specificity of this reaction for NADPH rather than NADH remained a matter of uncertainty for many years. Kaufman and his colleagues found that NADPH was the more efficient electron donor in his system (f18), whereas Mitoma reported NADH to be more effective (117). This discrepancy now appears to be satisfactorily explained by the recognition of two enzymes being involved in the overall reaction in addition to the actual phenylalanine hydroxylase. Kaufman established that an unconjugated pterin, tetrahydrobiopterin, is an essential cofactor in the hydroxylation reaction (120).In the course of the reaction this is converted to an oxidized form, identified as a quinonoid isomer of dihydrobiopterin (lal).

Tetrahydrobiopterin

o-Quinonoid dihydrobiopterin

116. S.Udenfriend and J. R. Cooper, JBC 194, 503 (1952). 117. C.Mitoma, A B B 60, 476 (1956). 118. 8. Kaufman, JBC 226, Fill (1957). 119. S.Kaufman, W.F. Bridgers, F. Eisenberg, and S. Friedman, BBRC 9, 497 (1962). 120. S.Kaufman, Proc. Nat. Acad. Sci. U . S. 50, 1085 (1963). 121. S.Kaufman, JBC 239, 332 (1964).

4.

233

FLAVIN AND PTERIDINE MONOOXYGENASES

Hence, the primary hydroxylase reaction may be written : H I CH,- C-COOH

H I CH,- C-COOH +

pheny lalanine tetrahydrobiopterin + 0, hydroxylase

OH L-Pheny lalanine

L-Tyrosine

+ quinonoid dihydrobiopterin

+

H,O

In order for this enzyme to function catalytically, the quinonoid dihydrobiopterin has to be reduced again. This is accomplished by dihydropteridine reductase, an enzyme long recognized t o participate in the overall reaction but only recently purified (122). This enzyme has been found to function much more efficiently with NADH (lower K , values and higher V,,, values) than with NADPH: Quinonoid dihydrobiopterin

+ NADH + H+ dihydropteridine reductase >

tetrahydrobiopterin

+N A P

The quinonoid dihydrobiopterin can rearrange nonenzymically to 7,8-dihydrobiopterin, which is the form of the cofactor isolated from rat liver (120).

7'

cH$-c-c'?XN&N,, I

H

I

H

I

NyNH2 0

When this form of the cofactor is present a third enzyme, identified as dihydrofolate reductase, is required for reduction to the physiologically active tetrahydrobiopterin (13). This enzyme is NADPH-specific: 7,ti-Dihydrobiopterin

-

+ NADPH + H+ dihydrofolate tetrahydrobiopterin + NADP+ reductase

Hence, the pyridine nucleotide specificity depends on the form of the cofactor present. However, it should be pointed out that the role of dihydrofolate reductase is as a scavenger of any dihydropteridine which has 122. J. E. Craine, E. S. Hall, and S. Kaufman, JBC 247, 6082 (1972).

234

VINCENT MASSEY AND PETER HEMMERICH

escaped from the quinonoid form and that in the presence of sufficient dihydropteridine reductase it plays only a minor role (13).It would therefore be more proper to restate the overall stoichiometry carried out hy phenylalanine hydroxylase and dihydropteridine reductase to be Phenylalanine

+ N A D H + Hf + 0 %+ tyrosine + N A D + + H20

The substrate specificity of phenylalanine hydroxylase is complicated by the fact that the rate of substrate oxidation is dependent on the nature of the tetrahydropteridine employed as cofactor (123), on the presence or absence of lysolecithin, whose effect is also dependent on the tetrahydropteridine cofactor employed ( l a d ) , and on the presence or absence of another protein recently isolated from liver known as the phenylalanine hydroxylase stimulating factor (PHS) (125,126).The effect of this protein also depends on the nature of the pteridine cofactor used. An additional complication arises from the fact that with several substrates increased rates of pyridine nucleotide oxidation occur (in the two enzyme system) without complete coupling to hydroxylation of the substrate. I n this case the product of 0, reduction is H,O,, a situation analogous to that found with several flavoprotein monooxygenases (see previous sections). I n fact, the phenomenon of “uncoupling” hydroxylation from oxidation with the phenylalanine hydroxylating system predated by several years (127) the discovery of this phenomenon with the flavoprotein monooxygenases. Again this effect depends on the nature of the substrate, the tetrahydropteridine, the presence or absence of lysolecithin, or the presence or absence of the PHS protein (128). The following compounds have been shown to be hydroxylated, a t least partially : tryptophan (129) p-2-thienylalanine (130), 4-chlorophenylalanine, 2-fluorophenalalanine1 3-fluorophenylalanine1 and 4-fluorophenylalanine (131). With the 4-fluorophenylalanine the products have been identified as tyrosine and F- (131). In addition, a number of p-substituted phenylalanines have been found to be hydroxylated with migration and retention of the p substituent (13%’).This demonstration of the “NIH 123.C.B. Storm and S. Kaufman, BBRC 32, 788 (1968). 124.D.B. Fisher and S. Kaufman, JBC 248,4345 (1973). 125. C.Y.Huang, E. E. Max, and S. Kaufman, JBC 248,4235 (1973). 126. C.Y.Huang and S. Kaufman, JBC 248, 4242 (1973). 127. S. Kaufman, BBA 51, 619 (1961). 128. D.€9. Fisher and S. Kaufman, JBC 248, 4300 (1973). 129. R.A. Freedland, I. M. Wadzinski, and H.A. Waisman, BBRC 5, 94 (1961). 130. S. Kaufman, “Methods in Enzymology,” Vol. 5, p. 802, 1962. 131. S. Kaufman, BBA 51, 619 (1961). 132. G. Guroff, C. A . Reifsnyder, and J. W. Daly, BBRC 24, 720 (1966).

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

235

shift” phenomenon (133) strongly implies the formation of arene oxide intermediates in the hydroxylation reaction. This will be considered further in the final section. B y varying the structure of the tetrahydropteridine cofactor, the extent of coupling the pteridine oxidation and hydroxylation of phenylalanine was determined. I n addition to the natural cofactor, 6-methyl and 6,7-dimethyl tetrahydropterin show tight coupling. I n contrast, 7-methyl, 7-phenyl, or unsubstituted tetrahydropterin show loose coupling (123). These results might imply that an alkyl group a t the 6 position is necessary for tight coupling to occur. However, substantial uncoupling of hydroxylation is observed with 4-fluorophenylalanine or tryptophan when 6-methyl tetrahydropterin is used as cofactor (123), making such generalizations somewhat untenable. When 6,7-dimethyl tetrahydropterin is employed as cofactor the initial rate of hydroxylation vs. phenylalanine concentration is hyperbolic. I n contrast, when the natural tetrahydrobiopterin is used as cofactor, the substrate saturation curve is sigmoidal ( l a d ) , Similar sigmoidal kinetics were reported with tryptophan as substrate, and the abolition of this complex behavior by 1-propanol ( 1 3 4 ) . This observation led Fisher and Kaufman (12-4) to explore the effects of various fatty acids and derivatives on the rates and kinetics of various reactions catalyzed by the hydroxylase. It was found that 1-propanol, 1-butanol, and a variety of fatty acids containing 16 carbon atoms or more gave substantial increases in catalytic activity when tetrahydrobiopterin was the cofactor, but were without effect with 6,7-dimethylpterin as cofactor. Long-chain acyl-CoA derivatives were also effective, as well as mixtures of bile salts and fatty acids. The greatest stimulations (of the order of 20-fold) were observed with phospholipids such as lysolecithin and lysophosphatidylserine. While the stimulating effect of lysolecithin on phenylalanine hydroxylation was exhibited only when tetrahydrobiopterin was used as cofactor, increased hydroxylation rates of other substrates were found even with 6,7-dimethyl tetrahydropterin as cofactor. Thus tryptophan hydroxylation is increased dramatically, especially a t low concentration of tryptophan ( 1 2 4 ) . The extent of uncoupling of hydroxylation and oxidation with this substrate remains unchanged, however. I n the presence of lysolecithin, with either tetrahydropteridine cofactor, the rate of hydroxylation of m-tyrosine was increased some 40-fold. Dopa, the product of 133. G. Guroff, D. Jerina, J. Rensen, S. Udenfriend, and B. Witkop, Science 157, 1524 (1967). 134. P. A . Sullivan, N. Kester, and S. J. Norton, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 30, 1067 (1971).

236

VINCENT MASSEY AND PETER HEMMERICH

this hydroxylation reaction, was formed stoichiometrically with respect to oxidation of NADPH, and the reaction is therefore tightly coupled (124). I n contrast, when (p)-tyrosine was tested as a substrate in the presence of lysolecithin, the rate of NADPH oxidation was greatly stimulated, but the tyrosine remained unchanged. Hence, in the presence of lysolecithin, tyrosine behaves as a nonsubstrate effector. This effect was shown to be the direct result of an increased rate of oxidation (producing H202)of the tetrahydropteridine cofactor (128). The stimulating effects of lysolecithin can also be mimicked by preincubation of phenalalanine hydroxylase with chymotrypsin (134).Lysolecithin exposes a thiol group of the enzyme which is unreactive to DTNB in the untreated enzyme (124). Fisher and Kaufman have interpreted these results as indicating that the hydroxylase contains a polypeptide portion which can act as an internal regulator of enzymic activity. It was proposed that the polypeptide can be either displaced reversibly from its inhibitory site by the detergent action of a lipid or can be irreversibly removed by chymotrypsin. If such a regulatory role does indeed operate it must be rather subtle, in view of the big differences in effects found depending on the nature of the tetrahydropteridine cofactor and the substrate used. Further insight into the details of the hydroxylation reaction is promised by the finding of yet another regulator of the enzymic activity. Studies by Kaufman and his colleagues have shown that even in the presence of lysolecithin, the specific activity of phenylalanine hydroxylation decreases sharply if the concentration of enzyme is increased (124). This effect (which was exhibited only with tetrahydrobiopterin as cofactor) was shown to be abolished by a phenylalanine hydroxylase stimulator (PHS) present in liver extracts. This factor has now been purified and shown to be a protein of molecular weight 51,500 (125). Evidence has been presented that PHS is an enzyme which catalyzes the breakdown of an intermediate in the hydroxylation reaction (126). The evidence presented, mainly of a kinetic nature, shows that the effect of PHS is unlikely to result from removal of an inhibitor in the hydroxylase preparations or from effects on the state of aggregation of the hydroxylase. Persuasive evidence is given for the reversible release from the hydroxylase of an intermediate which is subsequently converted nonenzymically to the products of the hydroxylation reaction. In this model PHS serves as an auxiliary enzyme catalyzing the breakdown of the intermediate : ki

ki

E+S=ES=E+S’ k-i

S’

k-z

k 2 p’

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

237

Under the experimental conditions where the phenomenon is observed (approximately equimolar concentrations of enzyme and tetrahydrobiopterin) the reverse step k-, can become significant, thus accounting for the observed decrease in specific activity as the hydroxylase concentration is increased. The fact that this effect is not found with 6,7-dimethyl tetrahydropterin would be explained simply by a lower value of k-, and/or an increased value of k , for this cofactor. Such arguments clearly rule out the possibility that S’ is a simple arene oxide intermediate of phenylalanine/tyrosine and suggest that S’ represents some complex or compound of the pterin, substrate, and O2 which is an intermediate in the hydroxylation reaction. The surprising conclusions from these studies are that such an intermediate is released from the enzyme before undergoing conversion to products, and the slow rate of its breakdown ( k , estimated as approximately 2.4 min-l). This latter value is almost 20 times slower than the calculated rate of release of S’ from the enzyme. Under suitable conditions it might therefore be possible to test this hypothesis directly ; for example, by initiating the reaction and then cooling rapidly, followed by low temperature chromatography (see results with luciferase, Section II1,I) it would be possible to isolate the intermediate and study its properties. Phenylalanine hydroxylase has been purified from rat liver to a state close t o homogeneity (135). The enzyme appears t o have a molecular weight around 100,000, and to be composed of two subunits of molecular weight 55,000. The enzyme has recently shown to contain iron a t the level of 1-2 atoms per molecule of protein (136‘). The enzyme is inhibited by metal chelators and substantial removal of the iron was achieved by incubation with o-phenanthroline and precipitation with ammonium sulfate. The activity was restored only by the addition of FeCl,. The enzyme was found to exhibit an EPR signal a t g = 4.23 attributable to high-spin ferric iron. This signal was substantially abolished when all three substrates were present under turnover conditions, suggesting that the iron may be reduced in the catalytic cycle, or, less probably, that a change in its ligand field results in a change from high- to low-spin state of the ferric iron (136). Unfortunately, no studies were carried out with tetrahydropteridine or phenylalanine added separately, which would be expected to better define the role of the iron. A steady-state kinetic study of the enzyme by Kaufman and Fisher (IS) indicated that a random order quaternary complex mechanism was operational (i.e., that a complex of all three substrates with the enzyme is 135. S. Kaufman and D. B. Fisher, JBC 245,4745 (1970). 136. D. B. Fisher, R. Kirkwood, and S. Kaufman, JBC 247, 5161 (1972).

238

VINCENT MASSEY AND PETER HEMMERICH

formed in the catalytic cycle). It should be noted, however, that these results are a t variance with other studies on the enzyme, which indicated a ping-pong type of mechanism (137).Kaufinan and Fisher (13) have reasoned that their kinetic analysis is inconsistent with reduction of the enzyme. However, their results would be consistent with a mechanism in which the quaternary complex was composed of reduced enzyme, an oxidized form of tetrahydropteridine (perhaps the semiquinone) , phenylalanine, and 02.Clearly, rapid reaction studies, to follow the rate of disappearance of the EPR signal in the presence of various combinations of the substrates, would be very desirable. Equally desirable would be experiments in which enzyme, tetrahydropteridine, and phenylalanine would be mixed with oxygenated buffer, and spectral and EPR changes monitored by rapid reaction techniques. Such experiments have proved very useful in investigating the mechanisms of flavoprotein monooxygenases and should be applicable to the present system.

B. TYROSINE HYDROXYLASE I n 1964 the enzymic conversion of L-tyrosine to 3,4-dihydroxyphenyIalanine (dopa) was demonstrated in particles isolated from adrenal medulla, brain, and other sympathetically innervated tissues (138). With partially purified preparations from adrenal medulla, the requirement in the catalytic activity of a tetrahydropteridine cofactor was demonstrated (158,139).Direct proof of the monooxygenase nature of the enzyme came from lSO studies by Daly et al. (140), who showed that the oxygen atom inserted at the 3 position of the benzene ring is derived from molecular oxygen : CH,-

C -COOH

CH,- C-COOH ~

tetrahydropteridine

+

+

O2-

dihydropteridine

+ H,O

OH OH

OH

The fact that tyrosine hydroxylase can be coupled to NADH oxidation by dihydropteridine reductase (139) strongly suggests that the dihydropteridine product is the quinonoid form (see Section IV,A) . The intracellular location of the enzyme has been a matter of some 137. V. G. Zannoni, I. Rivkin, and B. N. LaDu, Fed. Proc., Fed. Amer. SOC.Ezp. Biol. 26, 840 (1967). 138. T.Nagatsu, M.Levitt, and S. Udenfriend, JBC 239,2910 (1964). 139. A. R.Brennemann and S. Kaufman, BBRC 17, 177 (1964). 140. J. W. Dsly, M. Levitt, G. Guroff, and S. Udenfriend, ABB 126, 593 (1968).

4. FLAVIN

AND PTERIDINE MONOOXYGENASES

239

controversy. Nagatsu e t al. (138) , in their pioneering studies, concluded that in guinea pig brain and beef adrenal medulla, most if not all of the enzyme was particle bound, and that soluble enzyme found was the result of release by the homogenization method employed. More recent evidence indicates that in rat brain the enzyme exists in two forms, a soluble and a membrane-bound form (141). Soluble tyrosine hydroxylase has never been purified to any great extent: Almost all studies on the enzyme have been carried out with partially purified enzyme from bovine adrenal medulla which was solubilized by limited proteolysis with either trypsin (142) or chymotrypsin (143). The enzyme is inhibited by iron chelators such as ap-dipyridyl (138) and o-phenanthroline (1&) , but not by the nonchelating analog, m-phenanthroline (145). In addition, the enzymic activity is stimulated by Fe2+ (138). Shiman et al. (143) have pointed out that this stimulation is no valid evidence for participation of iron in the catalytic reaction since catalase has a similar effect. The activation by Fez+was therefore considered to result from removal of H,O, formed by autoxidation of the tetrahydropteridine. However, more recent studies, while admitting the validity of this argument, have produced convincing evidence that iron atoms are required in the catalytic function of the enzyme. Petrack e t al. (142,146') have demonstrated activation of the enzyme by ferrous iron; no other metal ion tested had a similar effect, nor did catalase or peroxidase. I n analogy with phenylalanine hydroxylase, a number of tetrahydropteridines will serve as electron donors with tetrahydrobiopterin being the most efficient in terms of lower K , value and higher V,,,, value. 6-Methyl and 6-7-dimethyl tetrahydropterin are both good donors (139). As would be expected, N(5)-substituted pteridines are inactive as cofactors; the tetrahydropteridine also has to be substituted with either a 2-amino or a 4-hydroxy group in order to function in the hydroxylation reaction (144). Like phenylalanine hydroxylase, the substrate specificity of tyrosine hydroxylase is fairly broad with relative rates depending on the pteridine cofactor used ; for example, phenylalanine is also hydroxylated to 141. 142. 143. 144. 145.

R. T. Kucaenski and A. J. Mandell, JBC 247, 3114 (1972). B. Petrack, F. Sheppy, and V. Fetaer, JBC 243, 743 (1968). R. Shiman, M. Akino, and S. Kaufman, JBC 246, 1330 (1971). L. Ellenbogen, R. J. Taylor, and G. B. Brundage, BBRC 19, 708 (1965). R. J. Taylor, C. S. Stubbs, and L. Ellenbogen, Biochem. Pharmacol. 18, 587

(1966). 146. B. Petrack, F. Sheppy, V, Fetaer, T. Manning, H. Chertock, and D. Ma, JBC 247, 4872 (1972).

240

VINCENT MASSEY AND PETER HEMMERICH

tyrosine a t a rate approximately the same as that of tyrosine hydroxylation when tetrahydrobiopterin is employed as the electron donor (143). However, with 6,7-dimethyltetrahydropterinthe rate is only about onetwentieth that of tyrosine hydroxylation (143). Tong et al. (147,148) have also reported that the conversion of m-tyrosine to dopa occurs at about 50% the rate of conversion of L-tyrosine to dopa. In addition, they found that phenylalanine hydroxylation resulted in approximately 15% formation of m-tyrosine and 85% of p-tyrosine. Studies with isotopically labeled substrate have demonstrated the N I H shift to be operational with this enzyme ( I . @ ) . When [4-aH]phenylalanine was employed as substrate, the tyrosine formed was found to have the tritium retained, indicating migration to the 3 and 5 positions. I n the subsequent conversion of [3,5JH] tyrosine to dopa, however, 50% of the tritium was lost (cf. Scheme 1, Section 111,~). Partial steady-state kinetic studies have been carried out with the enzyme. Using a partially purified soluble enzyme from adrenal medulla, Ikeda et al. (149) have found the kinetic behavior to be of the Ping Pong type, and concluded that a reduced form of the enzyme was involved in catalysis (presumably iron in the ferrous state). Omitting intermediate complexes, their results indicate the following sequence : E,.

+ tetrahydropteridine -+ Ered + dihydropteridine Ered

+ tyrosine + Oz+ E,, + dopa + H20

Working with a solubilieed enzyme, Joh et al. (150) concluded that the mechanism did not involve a reduced enzyme intermediate but rather a quaternary complex. Subsequent work by Shiman and Kaufman (quoted in 13) on both the particulate enzyme from bovine adrenal medulla and a purified solubilized enzyme also showed kinetic behavior indicative of the participation of a quaternary complex. While these experimental discrepancies are hard to rationalize, it should be pointed out that the finding of participation of a quaternary complex does not rule out reduction of the enzyme in the course of catalysis; such a complex could consist of reduced enzyme, oxidized pteridine, tyrosine, and 0,.

c. TRYPTOPHAN HYDROXYLASE (TRYPTOPHAN-5-MONOOXYGENASE) Much less information concerning the properties of tryptophan hydroxylase is available than with phenylalanine and tyrosine hydroxylases. Much of the literature deals with controversies about the subcellular loca147. J. H. Tong, A. D’Iorio, and N. L. Benoiton, BBRC 43, 819 (1971). 148. J. H. Tong, A. D’Iorio, and N. L. Benoiton, BBRC 44, 229 (1971). 149. M. Ikeda, K. A. Fahien, and S. Udenfriend, JBC 241,4452 (1966). 150. T. H. Joh, R. Kapit, and M. Goldstein, BBA 171, 378 (1969).

4.

241

FLAVIN AND PTERIDINE MONOOXYGENASES

tion of the enzyme and about whether a tetrahydropterin is indeed a cofactor. These aspects have been discussed extensively in a recent review by Kaufman and Fisher (13). The enzyme was first detected in brain by Grahame-Smith (151),who also partially purified the enzyme and demonstrated the requirement for a tetrahydropteridine cofactor (168). The relative paucity of information on the enzyme is the result of difficulties encountered in its assay and purification; the most highly purified preparation so far reported has been enriched only 10-fold over the starting material (153).Friedman et al. demonstrated convincingly that in the presence of reduced pyridine nucleotide and dihydropteridine reductase, tetrahydropteridines function catalytically in the reaction. They established the following stoichiometry (153): Tetrahydropteridine

+ tryptophan + 02

4

dihydropteridine 5-hydroxytryptophm

+

+ HtO

Like the other pterin-linked hydroxylases, several tetrahydropteridines were found to function as electron donors. However, tetrahydrobiopterin, the naturally occurring cofactor for phenylalanine hydroxylase, seems to be the most efficient donor, having itself a lower K , value than other tetrahydropteridines and also exhibiting lower K,, values for O2 and tryptophan than found in the presence of other pteridine cofactors (153,154). Although a definitive answer will have to await the availability of a more highly purified enzyme, there is evidence that iron atoms may be associated with the enzymic activity. Friedman et al. (153)have demonstrated that stimulation of the activity by Fez+is the result of removal of deleterious H,O,, but other workers have demonstrated substantial inhibition of the enzyme by the iron chelators, Tiron, a,a-dipyridyl, and o-phenanathroline (154,155).

V. Model Studies and Possible Mechanisms

In a series of papers, Mager and Berends (151-161) have proposed a common pathway for hydroxylation reactions catalyzed by flavopro151. D.G.Grahame-Smith, BBRC 16, 586 (1964). 152. D.G.Grahame-Smith, BJ 105, 351 (1967). 153. P. A. Friedman, A. H. Kappelman, and S. Kaufman, JBC 247, 4165 (1972). 154. E.Jequier, D.S. Robinson, W. Lovenberg, and A. Sjoerdsrna, Biochem. Pharmacol. 18, 1071 (1969). 155. A. Ichiyama, S.Nakamura, Y.Nishisuka, and 0. Hayaishi, JBC 245, 1699 (1970). 156. H.I. X. Mager and W. Berends, Rec. Trav. Chim. Pays-Bas 84, 1329 (1965). 157. H. I. X. Mager, R. Addink, and W. Berends, Rec. Trav. Chim. Pays-Bas 86, 833 (1967).

242

VINCENT MASSEY AND PETER HEMMERICH

teins and the pteridine-linked hydroxylases. Their proposal involves the formation of an intermediate hydroperoxide on reaction of a tetrahydropteridine or dihydroflavin with 02: H

I n the above general formulation the N(8) of the pteridines is equivalent to the N(10) of the flavins. The evidence of Mager and Berends for this formulation came from measurements of the stoichiometry of 0, consumption in oxidation of N (8)-substituted tetrahydropteridines and N (10)-substituted dihydroisoallaxazines. Less than stoichiometric 0, consumption was observed, which was formulated to result from the following reactions :

+ +

AH2 02 4 AHOOH AH2 AHOOH -+ 2 A 2 HzO (nonpolar medium) AHOOH + [AH+ OOH-] + A H a 2 (polar medium)

+

+

+

(1) (2)

(3) where AH, represents either tetrahydropteridine or dihydroflavin. In this formulation a complete coupling of reactions (1) and (2) would lead to a stoichiometry of one molecule of 0, consumed for each two molecules of AH,, and a coupling of reactions (1) and (3) to a stoichiometry of one molecule of 0, for each molecule of AH, oxidized. While such a formulation is consistent with their results, the observed stoichiometries could also be explained : AH2 AH1

+ Oz+ A + HzOz + HzOz+

A

+ 2 HzO

(4)

(5) Indeed, there are several reports in the literature that at physiological pH values H,O, is a comparable or even more efficient oxidant of tetrahydropteridines than is 0, (13,f62). However, with dihydroflavins, H,O, is a much poorer oxidant than molecular oxygen (163); thus, a t least 158. H. I. X. Mager and W. Berends, Rec. Trav. Chim. Pays-Bas 91,611 (1972). 159. H. I. X. Mager and W. Berends, Ree. Trav. Chim. Pays-Bas 91, 630 (1972). 160. H. I. X. Mager and W. Berends, Rec. Trav. Chim.Pays-Bas 91, 1137 (1972). 161. H. I. X. Mager and W. Berends, Tetrahedron 30, 917 (1974). 162. J. A. Blair and A. J. Pearson, JCS, Perkin Trans. I p. 80 (1974). 163. M. Dixon, BBA 226,2.59 (1971).

4.

243

FLAVIN AND PTERIDINE MONOOXYGENASES

with flavins, reactions (1)-(3) provide a reasonable interpretation of t,he results. Mager and Berends (157) proposed that the hydroperoxide might act. as a hydroxylating agent in the presence of a suitable acceptor:

'AHOH' (AHOH)

-H*O

Evidence in favor of a peroxydihydro intermediate bearing the OOH residue a t the bridge carbon between two nitrogen atoms was claimed by Mager and Berends ( 1 5 7 ) , using an N(1)-alkylated flavin model ( l-RFlredH,Scheme 2) as starting material for autoxidation. First, such an alkylated intermediate can be handled safely in aprotic solution be-

244

VINCENT MASSEY AND PETER HEMMERICH

cause of enhanced solubility. Second, spontaneous splitting of HzOz is overcome in this case, since this would require a proton a t N (1). In the presence of even slight amounts of water, the reaction sequence in this autoxidation will be HZO RFlredH

+

0,-

RF1- ?

-

O

O

HzO2

H

u RF1-10a-OH-

spirohydrantoin

while in the absence of water the decay of peroxydihydroflavin may be assumed to be RFl-?-OOH

2

[RFl-lOa-OH]-

spirohydrantoin

Henbe, the spirohydantoin (SPH) is the final product of l-alkyl-dihydroflavin autoxidation in any case (164). It should be pointed out that the SPH rearrangement is an entirely irreversible reaction which destroys the fla‘vin system. N (1)-C (10a) cleavage in a potential biocatalytic intermediate RF1-lOa-XH, XH being any protic nuelophile, must, therefore be strictly excluded biologically. In the above formulation a question mark has been put as to the position of OOH fixation in the 1-RFl,,-nucleus since (cf. Schemes 2 and 3) obviously any of the vinylogous positions 6, 8, 9a, and 10a might do. If there exists a rapid equilibrium between those possible isomers, i.e., a low activation barrier for shifts of nucleophiles like OOH- and OHon the flavin surface, the product SPH arising from irreversible decay of RF1-lOa-OH does not give any answer as to the structure of the actual oxygenating intermediate RFl-?-OOH. Such a low activation barrier for group migrations on the flavin surface has indeed been demonstrated by numerous papers of the Hemmerich group (for review, cf. 166,166). Since SPH, as mentioned above, is formed in any case as final product, the incorporation of lSO from lSO2 in the SPH-carbonyl could only then have evidential value for a RF1-10a-00H isomer if it were occurring efficiently in the presence of water. Under aqueous conditions, however, Blair and Pearson (162) demonstrated clearly that the carbonyl of the SPH is derived from water, not from 02. Under anhydrous conditions, as used by Mager and Berends (167),lSO from lSOz must trivially be found in SPH since no other relevant source of oxygen atoms is available. 164. K. H. Dudley and P. Hemmerich, J . Org. Chem. 32, 3049 (1967). 165. P. Hemmerich and W. Haas, in “Structure and Properties of Reduced Flavins”

(K. Yagi, ed.). Univ. of Tokyo Press, Tokyo (in preas). 166. P. Hemmerich and M. Schuman-Jorns, in “Enzymes: Structure and Function” (C. Veeger, J. Drenth, and R. A. Oosterbaan, eds.), p. 95. North-Holland Publ., Amsterdam, 1973; FEBS Symp. 29,95 (1973).

4.

245

FLAVIN AND PTERIDINE MONOOXYGENASES

,

OXIDASES

I

DEHYDROGENASES

nonessential {I; red

essential w k , t i u e

PEROXIDE heterolytic cleavage

H*

+G

SUPEROXIDE

I

N(I) blocked

N(51 blocked

SCHEME 3

Hence, the question as to the isomer structure of RF1-?-00H remains open : Miiller (personal communication) has meanwhile excluded positions 6 and 8 to be involved be means of thorough NMR studies of his “alcohol adducts” Rl-?-OR’(168), for which he initially claimed a 10a structure in line with Mager and Berends, but again without substantial evidence. Unfortunately, no facile differentiation between positions 9a and 10a can be made by NMR. A proposal as to the solution of this problem will be made below. Mager and Berends have found hydroxylation of phenylalanine using either tetrahydropteridines or dihydroflavins and molecular oxygen or hydrogen peroxide (I61) . They postulated the hydroxylating species to be hydroxy radicals. However, no convincing evidence is given for the 167. H. I. X. Mager and W. Berends, Tetrahedron Lett. 41, 4051 (1973). 168. F. Miiller, in “Flavins and Flavoproteins” (H. Kamin ed.), p. 363. Univ. Park

Press, Baltimore, Maryland, 1971.

246

VINCENT MASSEY AND PETER HEMMERICH

involvement of hydroxy radicals. But if it is assumed that the OH hypothesis is correct, two flavin molecules would be required for OH generation from molecular oxygen:

+

RFhH 02 + RFl-?-00H RFI-7-00H RFlredH + RFlOH

+

+ RFI + OH

This stoichiometry, involving stoichiometric flavin radical formation and “interflavin contact,” would make unlikely any biological importance of this “model” reaction. Furthermore, OH is such a reactive species that biocatalytic specificity of OH-involving processes could only be maintained if the radical was not allowed to diffuse away from the center of formation. But in that case it would be difficult to judge whether spin decoupling was indeed occurring in the catalytic pathway. Hence, oxygenation must occur in a quaternary complex made up from flavin, oxygen, substrate, and apoprotein, and the differentiation of OH- transfer on the one hand and oxygen at,om (“oxene” or OH+) transfer on the other hand turns into semantics. The true problem is as to the chemical structure of this active complex. The first experimental evidence indicating formation of oxygenated flavins came from rapid kinetic studies on the chemical reaction of reduced flavins with 0, (1-3), which was found to be an unexpectedly complex process involving the formation of peroxydihydroflavin and its decay into flavin radical and superoxide anion. The latter was found to be an even beter oxidant of reduced flavin than 0,, resulting in an autocatalytic reaction. The following series of steps was found to be minimal in describing the overall reaction (1,3,169) :

In a study of the reaction of various flavoproteins with 0, it has been found (4,s) that all dehydrogenases tested yield 0,- and the neutral flavin radical. On the other hand, no evidence for intermediate flavin 169. P. Hemmerich, A. P. Bhaduri, G. Blankenhorn, M. Brustlein, W. Haaa, and

W.-R. Knappe, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 3. Univ. Park Press, Baltimore, Maryland, 1973.

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

247

radicals or of 0,- could be found with flavoprotein oxidases, suggesting that with this group of enzymes the peroxydihydroflavin must undergo a heterolytic cleavage to yield directly the oxidized flavin and H,O,. This group of flavoproteins can also be characterized by their ability to stabilize the red-colored flavin semiquinone anion a t artificial half-reduction (170). The possibility of participation of 0,- in flavoprotein-catalyzed hydroxylation reactions has been considered. Massey and co-workers could find no evidence for 02-production with p-hydroxybenzoate hydroxylase or melilotate hydroxylase ; furthermore, superoxide dismutase had no effect on the hydroxylation reaction ( 4 ) .Similarly, White-Stevens and Kamin found no evidence with salicylate hydroxylase for the participation of 0,- or of hydroxy radicals ( 5 3 ) .On the other hand, Prema Kumar et al. (88) found that m-hydroxybenzoate-4-hydroxylase was inhibited completely a t fairly low levels of superoxide dismutase. While their interpretation of this result was that 0,- in some way functions as a hydroxylating agent we want to emphasize the possibility that superoxide dismutase may serve to catalyze the breakdown of a peroxydihydroflavin intermediate, e.g., by displacing the equilibrium (b) above. I n this case one would expect little or no change in the rate of NADPH oxidation, which should then be “uncoupled” from the hydroxylation reaction. Unfortunately, no information was given concerning this point. Strickland and Massey (171) found that hydroxylation of aromatic compounds such as p-hydroxybenzoate could be achieved a t physiological pH in the presence of model dihydroflavins and 0,. This reaction was inhibited substantially (75-100%) by superoxide dismutase. It seems unlikely that 0,- was the active hydroxylating agent however, since no hydroxylation was obtained by infusion of electrolytically generated 0,into a solution of p-hydroxybenzoate. These results again suggest that the active hydroxylating species must be the peroxydihydroflavin and that the inhibiting effect of superoxide dismutase in some way results from the destruction of this species, e.g., by removing 0,- from the equilibrium reaction (b) shown above for the autoxidation of dihydroflavins. Whatever the mechanism, there is little doubt that peroxydihydroflavins are active participants in flavoprotein-catalyzed hydroxylation reactions. As detailed in previous sections, transient species with very similar spectral properties have now been detected with three flavoprotein hydroxylases and shown to participate in the hydroxylation reactions. 170. V. Massey and G. Palmer, Biochemistry 5, 3181 (1966). 171. S. Strickland and V. Massey, in “Oxidases and Related Redox Systems” (T. E. King, H. S.Mason, and M. Morrison, eds.), Vol 1, p. 189. Univ. Park Press, Baltimore, Maryland, 1973.

248

VINCENT MASSEY AND PETER HEMMERICH

I n addition, a similar oxygenated flavin derivative has actually been isolated by low temperature chromatography in the case of bacterial luciferase (103). The important question yet to be answered definitely is the position of substitution in the flavin ring system and the mechanism by which oxygen is withdrawn from this intermediate and inserted into the aromatic substrate. Hamilton (14,86) has proposed a “vinylogous ozone” mechanism involving a nucleophilic attack of the aromatic substrate to a ring-opened form of a 4a-peroxydihydroflavin. No experimental evidence exists to support this hypothesis. While it is consistent with the introduction of a hydroxyl residue ortho to the original hydroxy group of the substrate (a phenomenon found with most flavoprotein hydroxylases) it could not account for the para substitution occurring with m-hydroxybenzoate hydroxylase (see Section II1,F). The theoretically possible HF1-00H isomers (cf. Scheme 3) must ab initio be separated into two subgroups, according to whether the position of proton fixation is N(5) or N ( l ) , i.e., one subgroup containing only the isomer 5-HF1-4a-OOH and the second containing four isomers l-HF1-6,8,9a,lOa-OOH. The first group isomer yields upon homolytic cleavage the blue (172,173), chemically stable, and biologically essential radical 5-HFlH, which is clearly associated with the “dehydrogenase” subclass of flavoproteins (5,170). This homolytic cleavage is easy because the spin density a t C(4a) is high (174) in the radical 5-HF1. The isomers of the second group would yield upon homolytic cleavage the tautomeric red (175,176), chemically unstable, and biologically nonessential radical l-HF1 or-the latter being strongly acidic-its anion F1-. This HF1-00H subgroup must, therefore, be associated with the oxidase and oxygenase subclasses of flavoproteins in keeping with their stabilization of the red flavin radical (5,170) arising from artificial l-e- oxidation or reduction, but not from reaction of the reduced enzymes with 0,. Furthermore, the question of which possible HF1-00H isomer is actually involved in flavin-dependent oxygenation requires at first the characterization of alkylated “model” flavin derivatives substituted in the respective positions 4a, 6, 8, 9a, 10a (cf. Scheme 3) by nucleophiles less 172. F. Muller, P. Hemmerich, A. Ehrenberg, G . Palmer, and V. Massey, Eur. J . Biochem. 14, 185 (1970). 173. F. Miiller, M. Briistlein, P. Hemmerich, V. Massey, and W. H. Walker, Eur. J. Biochem. 25, 573 (1972). 174. W. H. Walker, A. Ehrenberg, and J. M. Lhoste, BBA 215, 166 (1970). 175. A. Ehrenberg, F. Muller, and P. Hemmerich, Eur. J. Biochem. 2, 286 (1967). 176. F. Miiller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Press, Baltimore, Maryland, 1971.

4.

FLAVIN AND PTERIDINE MONOOXYGENASES

249

reactive than OOH-. Such model compounds RF1-X will exhibit chromophores practically identical with the respective HF1-00H chromophores and allow their structural assignment by comparison of absorption and fluorescence spectra. Walker et al. (177) were first to find OH- as a stable model nucleo350-370 nm. Rephile, characterizing 5-RF1-4a-OH ( R = benzyl) A, placement of OH- by mercaptide RS- indeed gives rise to facile homolytic C (4a)-SR cleave, yielding the blue radical 5-RFl, as required for a flavoprotein dehydrogenase model. Muller (168) used CH,O- as a model nucleophile characterizing 1-RFl?-OCH, (arising from 1-RFlox+ OCH,-, Scheme 2) hmax 410430 nm, as mentioned above. After F . Muller’s (personal communication) exclusion of positions 6 and 8 by NMR (cf. above) the question mark stands for either 9a or 10a. Experimental evidence for discrimination between these two possibilities is not available presently, but the following chemical reasoning should apply. Hemmerich and Miiller (178) pointed out that in l-RFlos+ the azomethine-type center 9a will be thermodynamically favored for nucleophile attack over the amidine-type center 1Oa. Furthermore, it is clear from the spin density map (179) of flavin radicals that C(10a) has a negligible spin density as compared to C (4a) , since 14N(1) does not contribute by spin polarization to the E P R hyperfine pattern with a splitting of more than 1 Hz. The same should be true for reasons of symmetry with C( 9a ) , though direct EPR-evidence is hampered by the lack of a magnetically active nucleus at C (9). Closer inspection of the 1-HFl-9aOOH structure reveals an acidic center at N ( l ) H with a pK estimated to be <7. Hence, the 9a isomer should be present under physiological conditions as the anion HOO-ga-Fl-, thus giving rise to facile polar elimination of OOH-, which, a t least theoretically, would suggest the 9a-dihydroperoxide t o be the isomer associated with flavoprotein oddases. Flavoprotein oxygenases, however,-requiring a “peracid-type” active intermediate-should involve a HF1-00H intermediate where the carbon center bearing the OOH group is sp2 rather than sp3, as pointed out by McCapra and Hysert (98). Such an intermediate can most easily arise from HF1-lOa-OOH, e.g., by reversible opening of one of the C-N bonds adjacent to C (10a). Since, as mentioned previously, the C (10a)-N (1) bond can only be opened irreversibly, because of spirohydantoin forma-

+

177. W. H. Walker, P. Hemmerich, and V. Massey, Eur. J. Bwchem. 13, 258 (1970). 178. P. Hemmerich and F. Muller, Ann. N. Y. Acad. Sci. 212, 13 (1973). 179. F. Muller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Press, Baltimore, Maryalnd, 1971.

250

VINCENT MASSEY AND PETER HEMMERICH

tion (cf. Scheme 2 ) , the ring opening must occur a t the C (10a)-N (10) bond. Such ring opening is known to be strictly reversible from the mechanism of chemical flavin synthesis (180). Thus, the oxygenation reaction is envisaged to occur as shown in Scheme 4. “Oxene” insertion into the aromatic substrate (ArH) yields

AroH-l 0

”PER ACID”

- Ar H-complex

ArH

SCHEME 4

finally the hydroxylated product (ArOH), presumably through a series of intermediates (see Sections II1,B and C ) . The flavin is left in an “alloxane-aminoanil” state of overall composition HFlOH ; such compounds are known to recyclize very rapidly, with elimination of water, to yield reoxidieed flavin catalyst (180). Hemmerich and Miiller (178)have collected arguments for a bZue color of the l-HF1-10a-OOH chromophore, and indeed a blue (Amax 600 nm) derivative arising upon aprotic autoxidation of l-RFlredHhas been demonstrated as early as 1960 by Hemmerich et al. (181), and has been 180. P. Hemmerich, C. Veeger, and H. 4, 671 (1965).

C. 5. Wood, Angew. Chem., Int. Ed. Engl.

4.

FLAVLN AND PTERIDINE MONOOXYGENASES

251

readily mistaken for the radical 1-RF1 (181,182), later on shown to be red (cf. above). Unpublished measurements of magnetic susceptibility with Ehrenberg proved this very labile blue intermediate to be diamagnetic, and very recent unpublished data of Hemmerich revealed a composition RF1-O-O-FIR, where the peroxo-linkage between the two flavin halves is thought to occur through C (10a). This chemical reasoning has been summarized in Scheme 3 and it would indeed allow the differentiation of the dehydrogenase, oxidase and oxygenase subclasses of flavoproteins according not only to their radical forms but also to their respective flavin oxygen “complexes” HF100H. It must be borne in mind, however, that isomerization of the three possibly “relevant” dihydroperoxides, l-HF1-9a/lOa-OOH and 5-HF1-4a-O0H, may be an intrinsically activationless process governed solely by apoprotein conformation. Hence, if HFlOOH stabilization is reached in an enzyme process experimentally, for example, at low temperature (103) or by complexing the active site with a “nonsubstrate effector’’ (cf. above), the intermediate detected might not be the true active HF1-00H but a ‘‘storage” isomer. While model studies indicate that hydroxylation reactions mediated by dihydroflavins and tetrahydropteridines may be the same (157), there are rather big differences between the properties of flavoprotein hydroxylases and pteridine-linked hydroxylases. The NIH shift has been demonstrated with the latter enzymes (see Section IV) , while there is no evidence yet produced for this happening with the flavoproteins. The pteridine-linked enzymes appear to contain iron, and a t least with phenylalanine hydroxylase there is evidence that the iron functions in the catalysis (136). The rate of reaction of 0, with tetrahydropteridines, while sufficiently fast t o complicate experimental work, is embarrassingly slow from an enzyme point of view, with half-lives a t physiological pH values in the order of minutes (161,157,162).This latter objection may be removed if further experimental work substantiates the conclusions of Kaufman and colleagues that an intermediate is liberated relatively rapidly from the enzyme and then decays slowly to yield products (196). However, this intermediate would have to possess unusual chemical properties consisting of a stable complex made up from oxygenated tetrahydropteridine and phenylalanine. If this were indeed the case it is difficult to envisage what the role of the iron might be. An alternative possibility is that it is the iron in the ferrous state which 181. P.Hemmerich, B.Prijs, and H. Erlenmeyer, Helv. Chim. Acta 43, 372 (1960). 182. K.H.Dudley, A. Ehrenberg, P. Hemmerich, and F. Miiller, Helv. Chim. Acta 47, 1354 (1964).

252

VINCENT MASSEY AND PETER HEMMERICH

serves to activate molecular oxygen for the hydroxylation reaction. I n this case the tetrahydropteridine may function simply to reduce the iron, in the same way as the dihydronicotinamide coenzymes serve t o reduce the flavin in flavoprotein hydroxylases. This possibility would eliminate the objection raised above concerning the slow rate of autoxidation of tetrahydropteridines but would not offer an explantion of the slowly decaying intermediate released from the enzyme. A third possibility suggested by Viscontini (183) is that the tetrahydropteridine complexes with the enzyme-bound Fe3+to form a Fez+-pteridine radical complex, which then reacts with 0, to form an oxygenated complex in which the 0, is liganded to the iron atom. The latter is then proposed to yield hydroxy radicals, the active hydroxylating species. While the concept of an iron-tetrahydropteridine complex being the active hydroxylating species is an interesting speculation, there does not appear to be any substantial evidence in favor oi hydroxy radical involvement since this would fail to provide any explanation for the NIH shift, which requires the formation of an arene oxide intermediate. In conclusion, it is clear that the detailed mechanisms of hydroxylation catalyzed by either the flavoproteins or the pteridine-linked enzymes are far from being understood despite the large amount of experimental effort so far expended. However, the most attractive explanation would seem to be that in both classes, whatever the nature of the primary oxygenated species may be, an oxene residue is transferred to the substrate with subsequent rearrangement to the hydroxylated product. ACKNOWLEDGMENT This review was written while Vincent Massey was on sabbatical leave a t the University of Konstanz. He wishes to acknowledge with gratitude a Senior U. S. Scientist Award of the Alexander von Humboldt Foundation. 183. M. Viscontini, in “Chemistry and Biology of Pteridines” p. 217. Int. Acad. Printing Co., Tokyo, 1970.

(K. Iwai et al., eds.),

Iron- and Copfier-Containing Monooxygenases V. ULLRICH

0

W. DUPPEL

I. Introduction. . . . . . . , . . . . . . . A. Historical Aspects , . . . . . . . . . . . B. Role of Iron and Copper Complexes in Oxygen Activation 11. Occurrence and Biological Importance . . . . . . . A. Nomenclature . . . . . . . . . . . . . B. Functions of Monooxygenases . . . . . . . . 111. Iron-Containing Monooxygenases . . . . . . . . A. Heme-Containing Monooxygenases . . . . . . . B. Nonheme Iron-Containing Monooxygenases . . . . IV. Copper-Containing Monooxygenases . . . . . . . A. Dopamine 8-Monooxygenase . . . . . . . . . B. Phenol o-Monooxygenase . . . . . . , , . .

. . . . . . . . . . .

. . . .

253 253 256 256 . 256 . 257 . 258 . 258 . 285 . 294 . 294 296

. .

1. Introduction

A. H I S ~ R I C A ASPECTS L The unique role of molecular oxygen, or dioxygen, as an electron acceptor in the respiratory chain is so predominant in cell metabolism that other functions of this molecule were not realized for a long time. Such functions are well established in the chemistry of oxygen and are referred to as “oxygenation” reactions (1). This term defines the oxidation of an 1. 0. Hayaishi, in “Oxygenases” (0. Hayaishi, ed.), p. 1. Academic Press, New York, 1962. 253

254

V. ULLRICH AND W. DUPPEL

organic or inorganic compound by direct introduction of one or two oxygen atoms of the dioxygen molecule. By the aid of the heavy isotope lSO and mass spectroscopy Mason et al. ( 9 ) in 1955 found proof that enzymes can use molecular oxygen to oxygenate their substrates. When phenolase (in this chapter referred to as "phenol o-monooxygenase") was incubated with ls02-enriched air and the phenolic substrate, the oxygen atom of the hydroxyl group formed was shown to originate from dioxygen according to the equation (3): RH

+ ''0~+ DH2 + R"0H + D + Hz"0

(1) where RH is the substrate and DH, the hydrogen donor. Although phenolase was early recognized as a copper-containing enzyme ( 4 ) ,the function of this metal ion in the reaction mechanism was far from being understood a t that time. Hydroxylation reactions with dioxygen as a cosubstrate were also identified in the microsomal fractions of the adrenal cortex and liver leading to the formation of C-21-hydroxylated steroids (6,6)or to hydroxylated drugs (7), respectively. Interestingly, these microsomal hydroxylations were inhibited by carbon monoxide in the dark, but not in the light, suggesting a possible role of ferrous ions in the reaction mechanism (5,8).Earlier spectroscopic investigations of the pigments in liver microsomes had revealed the presence of a carbon monoxide binding ferrous cytochrome with a Soret absorption band a t 450 nm wavelength ( 9 , I O ) . Since the same pigment was also found in adrenocortical microsomes, a possible role of this cytochrome in the steroid 21-hydroxylation appeared very likely (8).By the technique of the photochemical action spectrum, first used by Warburg (11) for the identification of the terminal oxidase in the respiratory chain, Estabrook et al. (8) identified the cytochrome with the unusual CO absorption band as the oxygen activating component of the steroid 21-hydroxylase system (see Fig. 1). Soon thereafter the cytochrome nature of the pigment in liver was H. S. Mason, W. L. Fowlks, and E. Peterson, JACS 77, 2914 (1955). H. S. Mason, Nature (London) 177, 79 (1956). F. Kubowitz, Biochem. Z . 299, 32 (1937). K. J. Ryan and L. L. Engel, JBC 225, 103 (1957). D. Y. Cooper, R. W. Estabrook, and 0. Rosenthal, JBC 238, 1320 (1963). 7. B. B. Brodie, J. Axelrod, J. R. Cooper, L. Gaudette, B. N. LaDu, C. Mitoma, and S. Udenfriend, Science 121, 603 (1955). 8. R. W. Estabrook, D. Y. Cooper, and 0. Rosenthal, Biochem. 2. 338, 741 (1963). 9. M. Klingenberg, ABB 75, 376 (1958). 10. D. Garfinkel, ABB 77, 193 (1958). 11. 0. Warburg, in "Heavy Metal Prosthetic Groups and Enzyme Action." Oxford Univ. Press (Clarendon), London and New York, 1949. 2. 3. 4. 5. 6.

5.

255

IRON- AND COPPER-CONTAINING MONOOXYGENASES

n a400

420

440

460

480

500

Wavelength [nm]

FIO.1. Photochemical action spectrum for the light reversal of CO inhibition of steroid 21-hydroxylation. Taken from Estabrook et al. ( 8 ) . established by the work of Omura and Sat0 (19J.3). They tentatively suggested the name “cytochrome P-450” which has not yet been replaced by a more systematic nomenclature. Shortly after, a P-450-like cytochrome was also found in adrenal mitochondria (14) and identified as a part of the steroid llp-hydroxylating system (16). An important step forward toward the elucidation of the molecular function of the cytochrome was its isolation and purification from a Pseudomonas putida strain grown on camphor as the sole carbon source (16-18). It is now evident that many cytochrome P-450-like hemoproteins exist which function as oxygen activating components in a variety of hydroxylating systems. The special coordination sphere and environment of the porphyrin iron seems to be directly related to this function. However, other iron-dependent hydroxylating systems with different chelating structures for the iron ion do exist and certainly more will be found in the future. 12. T. Omura and R. Sato, JBC 239, 2370 (1964). 13. T. Omura and R. Sato, JBC 239, 2379 (1964). 14. D. C. Sharma, E. Forchielli, and R. I. Dorfman, JBC 237, 1495 (1962). 15. L. D. Wilson and B. W. Harding, Biochemistry 9, 1615 (1970). 16. J. Hedegaard and I. C. Gunsalus, JBC 240,4038 (1965). 17. J. A. Peterson, ABB 144,678 (1971). 18. C. A. Yu, 1. C. Gunsalus, M. Katagiri, K. Suhara, and S. Takemori, JBC 249, 94 (1974).

256

V. ULLRICH AND W. DUPPEL

B. ROLEOF IRON AND COPPERCOMPLEXES IN OXYGEN ACTIVATION Besides some bacterial flavoproteins all other oxygenating enzymes contain iron or copper ions that are intimately involved in the activation process of the oxygen molecule. This fact deserves some general considerations before going into a more detailed description of monooxygenases. As a consequence of its unique electron configuration the dioxygen molecule is metastable in the presence of most organic compounds although the free energy of its reduction to water is extremely favorable (19). Once reduced it loses its metastability and the reduced intermediates, the 0,- ion, hydrogen peroxide, and the OH radical, represent highly reactive species. During the aerobic metabolism of the cell these intermediates may be formed, but they are most efficiently removed by the action of copper-dependent dismutases and catalase (20). The two transition elements copper and iron are essential cofactors of oxygen utilizing enzymes, such as hemoglobin, myoglobin, hemocyanins, and erythrocruorins as biological oxygen carriers, the iron-dependent dioxygenases, cytochrome oxidase, and the peroxidases. The cofactor function of iron and copper is based on their pronounced redox properties, their ability to form various types of metal-oxygen bonds, and the wide variation of their redox properties caused by complexation with biological chelates and proteins. Thus, conformational changes as intrinsic properties oi enzyme proteins can trigger changes in the redox behavior of the metal ion or vice versa.

11. Occurrence and Biological Impo,rtance

A. NOMENCLATURE Hayaishi (1) first proposed the name “oxygenases” for enzymes which can introduce molecular oxygen into organic substrates. If both atoms of the dioxygen molecule appear in the substrate the corresponding oxygenases are now called “dioxygenases.” Consequently, Hayaishi (21) has suggested the term “monooxygenases” for those enzymes that incorporate only one atom into the substrate with concomitant reduction of the second atom to water. Linguistic simplification with reference to “monoxide” 19. P. George, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 3. Wiley, New York, 1965. 20. E. M. Gregory and I. Fridovich, J . Bucteriol. 114, 1193 (1973). 21. 0.Hayaishi, Proc. Plen. Sese., Int. Congr. Biochem. 6th, 1964 Vol. 33, p. 31 (1984).

5.

IRON- AND COPPER-CONTAINING

MONOOXYGENASES

257

has led to “monoxygenases,” a term that should be recommended for use in the official nomenclature ( 2 l a ) . Historically, the group of hydroxylating enzymes was first called “hydroxylases.)’ This turned out to be misleading and insufficient for a characterization since a hydroxyl group may be derived either from water or dioxygen. Moreover, not all monooxygenases catalyze hydroxylations but also epoxidations, lactonizations, or oxidations a t heteroatams. I n 1957,Mason (22)had proposed the name “mixed function oxidases” in order to characterize the mixed function of the oxygen molecule. The official nomenclature of the Enzyme Commission uses the term “hydroxylases” as a subgroup of the oxidoreductases under E C 1.14 and distinguishes i t from the “oxygenases” (EC 1.13) which incorporate both oxygen atoms. This nomenclature is neither logical nor practical. Since the electron donor for the reaction is to be included the official name for the enzyme that converts tyrosine to 3,4-dihydroxyphenylethylamine (DOPA) (EC 1.14.2.1) is “3,4-dihydroxyphenylethylamine: ascorbate: oxygen-oxidoreductase (hydroxylating) .” I n this chapter the term “monooxygenase” will be used. It will be preceded by the main substrate of the enzyme and the position to be attacked. The system mentioned above can then be called tyrosine 3-monooxygenase. Addition of the electron donor in brackets is possible, but often the reductant is unspecific or acts indirectly, and thus no better characterization of the monooxygenase is achieved.

B. FUNCTIONS OF MONOOXYGENASES Monooxygenases play an important role in many biosynthetic and metabolic pathways and often have regulatory properties since their molecular activities are usually low. They are involved in amino acid metabolism, e.g., in the biosynthesis of tyrosine, DOPA, hydroxyproline, hydroxylysine, or serotonin, and in the formation of steroids and bile acids. Equally important is the function of monooxygenases in the degradation of lipophilic organic compounds. This includes the inactivation of steroid hormones and the oxidation of drugs and foreign compounds. The introduction of a hydroxyl group in one of these lipophilic substrates greatly increases their water solubility and thus enhances their elimination from the body. The corresponding monooxygenases are found not only in highly specified mammalian tissues but also in yeasts, plants, fungi, and 21a. Although the editor finds value in the author’s recommendation to use the

term “monoxygenase,” in this chapter the designation “monooxygenase” is used to conform with usage in other chapters in the volumes. 22. H. S. Mason, Adwan. EnzymoZ. 19, 79 (1957).

258

V. ULLRICH AND W. DUPPEL

bacteria. The importance of monooxygenases in adaptive microorganisms has only recently been recognized. Bacteria and also yeasts can grow even on nonphysiological carbon sources such as aromatic or aliphatic hydrocarbons ( 2 3 ) . These organisms contain inducible monooxygenases for the corresponding hydrocarbons and convert their substrates into phenols or alcohols which undergo further oxidative attack until norma1 metabolic pathways can be used for the complete oxidation of the molecules. In this way the conversion of petroleum into animal food has been achieved (24).The stereospecificity of most monooxygenases in adaptive bacteria has also been made use of for the biotechnical production of hydroxylated steroids which are difficult to obtain by chemical synthesis

w’6)

The biological importance of monooxygenases in genera1 can be summarized by recalling the fact that no other biochemical mechanism of attack a t a nonactivated carbon-hydrogen bond is known. The following chapters contain a survey of sufficiently characterized monooxygenases from various sources.

111. Iron-Containing Monooxygenases

A. HEME-CONTAINING MONOOXYGENASES

A series of reactions is involved until the product is released from a monooxygenase system. The most important of these molecular events is the activation of the dioxygen molecule to the (‘active oxygen” species and this must take place a t the catalytic site of the enzyme. In the case of iron-containing monooxygenases the catalytic center consists of the ferrous ion in a special coordination sphere. At least one coordination bond is required for the fixation of the metal to the protein, the others may be provided by additional amino acids or can be formed by biological chelators like porphyrins..Indeed, the heme ring is known at present to be the most common prosthetic group of monooxygenases ; however, its binding to the protein must occur quite differently from other hemoproteins since all heme-containing monooxygenases have in common an unusual Soret band of their carbon monoxide complexes at about 450 nm. This characteristic absorption band has led to the designation ‘(cyto23. A. C. van der Linden and G. J. E. Thijsse, Advan. Enzymol. 27, 469 (1965). 24.J. Someya, T. Murakami, N. Tagaya, N. Futai, and Y. Sonoda, J. Ferement. Technol. 48, 291 (1970). 25. H. Iizuka and A. Naito, “Microbial Transformations of Steroids and Alkaloids.” Univ. Park Press, Baltimore, Maryland, 1968.

5.

259

IRON- AND COPPER-CONTAINING MONOOXYGENASES

chrome P-450” for the hemoprotein in the monooxygenase system of liver microsomes (12 ) . After the discovery of more P-450-like hemoproteins another problem of nomenclature arises. It seems inappropriate to keep the name ‘‘cytochrome P-450” for all heme-containing monooxygenases with different specificities for substrates and electron donors. Indices like “P,-450” or “P-45OC,,” which are occasionally used in literature do not principally solve the problem. However, for ‘it more systematic designation more details on the heme ligands must be known. The various functions of cytochrome P-450 species have been reviewed by Lemberg and Barrett ( 2 6 ) . When looking a t heme-containing monooxygenases from different sources i t becomes apparent that the electron transport from the pyridine nucleotides to the hemoprotein is mediated by two different mechanisms. One group uses iron-sulfur proteins together with a flavoprotein for the reduction of the cytochrome. These systems occur in mitochondria and bacteria. The other type uses flavoproteins directly and is associated with endoplasmic membranes. Therefore, a subdivision of the heme-containing monooxygenases into these two groups was made in this chapter.

1. Iron-Sulfur Proteins as Electron Donors a. Camphor 5-exo-Monooxygenase of P . putida. A bacterial monooxygenase system that converts D-camphor to the 5-exo-alcohol according to the equation &Camphor

+ NADH + H+ + O2-+

em-5-OH-camphor

+ NAD+ + H20

(2)

was reported by Hedegaard and Gunsalus (16)in 1965. The monobxygenase activity was only present when the P. putida strain was grown on camphor and the corresponding genes were found to be localized on an episome which contained all genetic information for the camphor-isobutyrate pathway (27). The spectral properties of bacterial extracts clearly showed the presence of a cytochrome P-450 species, but in contrast to other known heme-containing monooxygenases all components of the system proved to be soluble and could be purified to homogeneity (18,28,29). An extract of camphor-grown cells contained the hemoprotein, also called “cytochrome P-45OC,,,,” in a concentration of about 1 nmole/mg of protein. After DEAE-cellulose chromatography, ammonium sulfate fractionation ( 30 4 5 % saturation), gel filtration, calcium phosphate gel 26. R. Lemberg and J . Barrett, “Cytochromes,” p. 73. Academic Press, New York, 1973. 27. A. M. Chakrabarty and I. C. Gunsalus, Genetics 68, 510 (1971). 28. R. L. Tsai, I. C. Gunsalus, and K. Dus, BBRC 45, 1300 (1971). 29. P. W. Trudgill, R. DuBus. and I. C. Gunsalus, JBC 241, 1194 (1966).

260

V. ULLRICH AND W. DUPPEL

adsorption, and a second DEAE-chromatography the specific content increased 22- to 27-fold (18). All purification steps were carried out in the presence of camphor, which finally could be removed by dialysis or chromatography with 20 mM Tris-chloride buffer on Sephadex gels (17,18).The isolated hemoprotein, which could be crystallized (1430) as the camphor complex as well as in the substrate-free form, has a molecular weight of 44,000-46,OOOin a single polypeptide chain. The amino acid composition is not unusual and shows a total of six cysteines and one tryptophan. The amino terminal residue is asparagine or aspartate and valine forms the carboxyl end (18,28). According to the spectral properties the cytochrome is different from other known hemoproteins as can be seen from Fig. 2 and Table I (31-33) . The blue shift of the Soret absorption band after reduction and the resulting low extinction coefficient of the Soret band is rather unusual. Most characteristic, however, is the y-absorption band a t 447 nm of the carbon monoxide complex in the reduced state. Interestingly, metyrapone (2-methyl-1,2-di-3-pyridyl-l-propanone)(33) and N-phenylimidazole

X (nm)

FIG.2. Absorption spectra of cytochrome P450,., (15). 30. C. A?.Yu and I. C. Gunsalus, BBRC 40, 1431 (1970). 31. C. A. Tyson, J. D. Lipscomb, and I. C. Gunsalus, JBC 247, 5777 (1972). 32. B. Griffin and J. A. Peterson, ABB 145, 220 (1971). 33. J. A. Peterson, V. Ullrich, and A. G. Hildebrandt, ABB 145, 531 (1971).

5.

261

IRON- AND COPPER-CONTAINING MONOOXYGENASES

TABLE I SPECTRAL PROPERTIES OF CYTOCHROME P-450,,,

AND

ITS LIQANDCOMPLEXES ~

System

Maxima (nm).

P-4500, P-450,, camphor P-4.50red P-450red - Oa P-45Or,d - CO P-4500, - ethyl isocyanide P-450red - ethyl isocyanide P-450,, - metyrapone P-450r.d - metyrapone

+

(I

Extinction coefficients

(e,~)

417(105) 391 (87) 411 (71) 418 (62) 447(106) 430(114) 453(104) 421 (99) 442(107)

535(10) 571(10.5) 540(10) 646(4.5) 540(14) 552(14) 550(12) 548 549, 576 539(21) 566(22)

~~~

Ref. 31 31 31 31 31 3.2 32 33 33

in parentheses.

(34) in stoichiometric amounts form ligand complexes with y bands also around 445 nm in the reduced form. Ethyl isocyanide binds more strongly to the reduced state than to the ferric cytochrome and competes with camphor for the binding site ( 3 2 ) . The presence of camphor induces a dramatic change in the absorption spectrum. The y band shifts to 394 nm and a new band appears a t 645 nm (17,35).The spectral dissociation constant (K,) of this conversion is close to the K , of the overall hydroxylation reaction (17,S6), thus indicating the formation of an enzyme-substrate complex. Cations potentiate the affinity for the camphor binding ( 1 7 ) . Only D ( + ) - and L(-)-camphor and some closely related compounds can cause this spectral change ( S 5 ) .Concomitant with the change in the optical spectrum a conversion of the spin state occurs. The camphor-free enzyme is low-spin with g values a t 2.45, 2.26, and 1.91 ( 3 7 ) . Addition of camphor produces a high-spin EPR signal with g values a t 8.4 and 1.8 which are seen only at liquid helium temperature (Fig. 3 ) . From the g values a very strong rhombicity of the heme was derived ( S 7 ) . The redox potential of the low-spin ferric hemoprotein has been reported to be around -380 mV whereas the potential of the high-spin form is about 200 mV higher (38). By isoelectric focusing the isoelectric point of the low-spin form was found at 4.55 which increased to 4.67 34. R. L. Tsai, P1i.D. Thesis, University of Illinois, Urbana, 1969. 35. I. C. Gunsalus, Hoppe-Seyler’s Z. Physiol. C h e m . 349, 1610 (1969). 36. I. C. Gunsalus, C. A. Tyson, R. L. Tsai, and J. D. Lipscomb, Chem.-Biol. Interact. 4, 75 (1971). 37. R. L. Tsai, C. A. Yu, I. C. Gunsalus, J. Peisnch, W. Blumberg, W. H. OrmeJohnson, and H. Beinert, Proc. Nut. Acarl. Sci. U. S . 66, 1157 (1970). 38. J. A. Peterson, cited in Peterson et al. (33).

V. ULLRICH AND W. DUPPEL

262 79OK

15'K 226 194

P - 450' CAMPHOR

FIQ.3. EPR spectra of cytochrome P-450,.,. Effect of substrate and phenylimidarole ( 5 7 ) .

after the binding of camphor (39). This has been interpreted as the release of a weakly basic group or the masking of an acid. Urea, acetone, mercurials, detergents, and other denaturating conditions convert cytochrome P-450,,, to an inactive form which has an absorption of the carbon monoxide complex a t 420 nm in the reduced state (40). Reactivation to cytochrome P-450 by sulfhydryl compounds is possible only with freshly prepared cytochrome P-420. The heme group of the cytochrome can be dissociated from the apoprotein by incubation with apomyoglobin and hemopexin (41,4.2). The concomitant loss of the hydroxylase activity is partly restored by addition of ferriprotoporphyrin IX. NADH is the electron donor for the monooxygenase system, and an electron transport system is required for the reduction of the hemoprotein. 39. K. Dus, M. Katagiri, C. A. Yu, D. L. Erbes, and I. C. Gunsalus, BBRC 40, 1423 (1970). 40. C. A. Yu and I. C. Gunsalus, JBC 249, 102 (1974). 41. U. Muller-Eberhard, H. H. Liem, C. A. Yu, and I. C. Gunsalus, BBRC 35, 229 (1989). 42. C. A. Yu and 1. C. Gunsalus, JBC 249, 107 (1974).

5.

IRON- AND COPPER-CONTAINING MONOOXYGENASES

263

This system consists of a flavoprotein and an iron-sulfur protein which were isolated as pure proteins (28,29,43). The reductase has a molecular weight of 43,500 and contains in a single polypeptide chain one mole of FAD perhaps attached through a cysteine sulfur (28). Extensive investigations have been carried out on the iron-sulfur protein which was called “putidaredoxin.” This protein was easily purified and has a molecular weight of 12,500 (28;43). The prosthetic group of oxidized putidaredoxin includes two ferric irons which are linked t o two atoms of acid-labile sulfur and four cysteines of the protein (28). The natural-abundance iron has been replaced by “Fe and 59Feand the acidlabile sulfur by selenide with retention of activity (44-46). This isotopic replacement has allowed EPR and ENDOR studies (47,48). The primary sequence (28) resembles that of adrenodoxin and also the redox potential of about -240 mV a t pH 7 (49) is similar. Putidaredoxin is only a oneelectron acceptor, and the electron resides in one of two irons (47). Mossbauer measurements in strong magnetic fields show that the sign of the magnetic hyperfine tensor has apposite signs for the ferric and the ferrous site (60). Although both irons in the oxidized state are high-spin they are antiferromagnetically coupled to give a diamagnetic ground state. In the presence of camphor the cytochrome P-450,,, can be reduced by stoichiometric amounts of NADH provided that the flavoprotein and putidaredoxin are also present (26). Without the substrate no reduction is seen, which may be explained by the very low potential of the low-spin state of the cytochrome. When an excess of reducing equivalents is avoided and oxygen is admitted one can observe the formation of an oxygenated species with a Soret absorption band a t 418 nm ( 6 1 6 5 ) . This 43. D. W. Cushman, R. L. Tsai, and I. C. Gunsalus, BBRC 26, 577 (1967). 44. D. V. Dervartanian, W. H. Orme-Johnson, R. E. Hansen, H. Beinert, R. L. Tsai, J. C. M. Tsibris, R. C. Bartholomaus, and I. C. Gunsalus, BBRC 26, 569 (1967). 45. J. C. M. Tsibris, R. L. Tsai, I. C. Gunsalus, W. H. OrmeJohnson, R. E. Hansen, and H. Beinert, Proc. Nat. Acad. Sci. U . S. 59, 959 (1968). 46. J. C. M. Tsibris, M. J. Namtvedt, and I. C. Gunsalus, BBRC 30, 323 (1968). 47. J. Fritz, R. Anderson, J. Fee, G. Palmer, R. H. Sands, J. C. M. Tsibris, I. C. Gunsalus, W. H. OrmeJohnson, and H. Beinert, B B A 253, 110 (1971). 48. W. H. Orme-Johnson, R. E. Hansen, H. Beinert, J . C. M. Tsibris, R. C. Bartholomaus, ahd I. C. Gunsalus, Proc. Nat. Acad. Sci. U. S. 60, 368 (1968). 49. G. S. Wilson, in ‘‘Mossbauer Spectroscopy in Biological Systems” (P. Debrunner, J. C. M. Tsibris, and E. Miinck, eds.), Allerton House Meet. Proc., 1969. 50. E. Miinck, P. G. Debrunner, J. C. M. Tsibris, and I. C. Gunsalus, Biochemistry 11, 855 (1972). 51. I. C. Gunsalus, Con!. P&O Struct. Funct., 1970 (1970). 52. Y. Ishimura, V. Ullrich, and J. A. Peterson, BBRC 42, 140 (1971). 53. J. A. Peterson, Y. Ishimura, and B. W. Griffin, ABB 149, 197 (1972).

264

V. ULLRICH AND W. DUPPEL

species decays with a first-order rate constant of 0.01 6ec-I and the 391-nm Soret band characteristic of the high-spin cytochrome-camphor complex reappears ( 2 6 ) . The oxygenated cytochrome is diamagnetic and also in its spectral properties very similar to the oxy forms of hemoglobin and myoglobin ( 4 6 4 ) . No product is obtained with the oxy form alone unless reduced putidaredoxin is added ( 3 1 ) .The reduction of the oxy complex is strictly dependent on putidaredoxin although slow reduction of the ferric cytochrome can also be achieved by other iron-sulfur proteins ( 3 1 ) . These results point to a specific interaction of putidaredoxin with the cytochrome and, indeed, the CD spectra of a mixture are different from the sum of the components (26). Electrofocusing experiments have been reported in which 6: 1, 3: 1, and 1:1 aggregates of putidaredoxin and cytochrome P-450,,, were detected a t high concentrations (55). A titration of the cytochrome with oxidized putidaredoxin followed by EPR showed a distinct shift in the position and intensity of the g tensors until a 1 :1 ratio was approached. The ratio of the components that are active under biological conditions is therefore suggested to be 1:1:1. According to kinetic measurements of the single steps in the reaction cycle and rapid scan spectroscopy the transfer of the second electron from reduced putidaredoxin to the oxycomplex represents the rate-limiting step ( 3 1 ) .I n Table I1 the reported kinetic data of the individual reactions are listed.

RATECONSTANTS

OF

REACTION8

TABLE I1 CAMPHOR &MONOOXYOENATION CYCLEa

I N THE

First-order rate constant (sec-l)

Roaction

+

-

RH Camphor (RH) oxidized cytochromeP-450,., --$ P-450,, e (putidaredoxin) -+ P-450,,d - RH P-450,, - RH RH 09 -+ P-450 - RH P-45Or.d

7000 35 470

- RH + e (putidaredoxin) -+P-450red - RH

17

+

-

+

A 2

P-450

I

0 2

I

0 2

aP-450,.,, reductase, and putidaredoxin in a ratio of 1: 1: 2. Under these conditions the reductase is not rate limiting.

54. M. Sharrock, E. Munck, P. G. Debrunner, V. Marshall, J. D. Lipscomb, and I. C. Gunsalus, Biochemistry 12, 258 (1973). 55. K. Dus, J. D. Lipscomb, and I C. Gunsalus, Abstr., 169nd Meel., Amer. Chem. Soc. (1971).

5.

IRON- AND COPPER-CONTAINING MONOOXYGENASES

265

b. 1 Ip-Steroid Monooxygenase of Adrenal Mitochondria. The high activity in steroid hormone biosynthesis of the adrenal cortex has led to early studies on the mechanism of biological hydroxylations. In 1951, Sweat (56) had shown by i n vitro studies that adrenocortical mitochondria were able to hydroxylate steroids a t the 11 position. Estabrook et al. ( 8 ) reported the occurrence of a CO binding hemoprotein in bovine adrenal cortex microsomes and established its function in C-21 steroid hydroxylation by means of the photochemical action spectrum of the carbon monoxide inhibited reaction. Harding e t d.(57) reported the presence of a GO binding pigment in the mitochondria1 fraction of the adrenal cortex and assumed its involvement in steroid 1lp-hydroxylation, which could be established by Wilson and Harding (58). Like the soluble bacterial camphor monooxygenase system, the steroid llp-monooxygenase of adrenal mitochondria consists of the cytochrome P-450 containing terminal oxidase, an iron-sulfur protein (adrenodoxin or adrenal ferredoxin) , and a flavin-containing reductase (59). In contrast to the bacteria1 system, these components are bound to the inner mitochondrial membrane and require NADPH as an electron donor. The system could be solubilized by sonication of intact mitochondria and was separated into its components (59). Most of the P-450 hemoprotein of the adrenal mitochondria could be found in the precipitate after the first centrifugation, from where it was solubilized by treatment with the nonionic detergent Triton NlOl (60).The subsequent procedure involved lyophiliaation and extraction with acetone and l-butanol to remove lipids. Further purification by ammonium sulfate precipitation and subsequent chromatography on DEAE-cellulose yielded a clear and stable solution. A similar procedure was used by Kinoshita et al. (61) but with sodium cholate as a solubilizing agent. Improved preparations resulted in a threefold purification of the hemoprotein to a specific content of 6-8 nmoles/mg of protein ( 6 2 ) .Eighty percent of the total lipid and 99% of the cholesterol were removed. The preparations had hardly, if any, cytochrome P-420 contamination and were still active in llp-hydroxylation after recombination with both electron transport components 56. M.L. Sweat, JACS 73, 4056 (1951). 57. B. W. Harding, S. H. Wong, and D. H. Nelson, BBA 92,415 (1964). 58. L. D.Wilson and B. W. Harding, Biochemistry 9,1615 (1970). 59. T.Omura, E . Sanders, R. W. Estabrook. D. Y. Cooper, and 0. Rosenthal, ABB 117, 660 (1966). 60. H.Schleyer, D.Y. Cooper, and 0. Rosenthal, JBC 247, 6103 (1972). 61. T. Kinoshita, S. Horie, N. Shimazono, and T. Yohro, J. Biochem. (Tokyo) 60, 391 (1966). 62. F. Mitani and S. Horie, J. Biochem. (Tovko) 65, 269 (1969).

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( 6 3 ) .The spectral properties of the adrenal hemoprotein closely resemble lthose of soluble cytochrome P-450 of the camphor monooxygenase system. Low temperature spectra have been employed to better evaluate the maxima of the weaker absorption bands ( 6 3 ) . The spin state of the isolated hemoprotein was reported to be low spin in the Triton preparation (6 4 ), while in the cholate preparation the coexistence of a high- and low-spin form was shown (65). Jefcoate e t al. (66) succeeded in separating the high-spin and low-spin cytochrome P-450 by ammonium sulfate fractionation after isooctane extraction of the cholate solubilized enzyme. It turned out that the high-spin form exhibited a reversed substrate binding spectrum with 20a-hydroxycholesterol and was active in side chain cleavage in the reconstituted system, while the low-spin form showed the typical substrate binding spectrum with ll-deoxycorticosterone and was active in llpmonooxygenation. Another report showed an increase in the optical spectrum of the high-spin form by addition of deoxycorticosterone, while pregnenolone caused an increase of the low-spin form (66). Alfano e t al. (67) have also described two forms of high-spin cytochromes, one with a g value of 7.9 in the EPR spectrum, which was attributed to the substrate-bound high-spin form of the llp-monooxygenase cytochrome P-450, and the other with a g value at 8.2, which represents the high-spin form of cytochrome P-450 involved in cholesterol side chain cleavage. Elevated blood ACTH levels increased the 8.2 signal, indicating a regulating hormonal effect of ACTH on the utilization of cholesterol by the side chain cleavage monooxygenase system. Two spectral dissociation constants for deoxycorticosterone have been determined by titration analysis, a tight binding near the active center of M and a second unspecific one, hydroxylation with a Kd of 1.8 X M ( 6 8 ) . These numbers are in good agreement with a K d of 40 X with the K,,, value of about 30 pM for the steroid hydroxylation in the reconstituted system, showing that cytochrome P-450 is the terminal oxidase in the steroid 11b-monooxygenation system. 63. D. Y. Cooper, H. Schleyer, and 0. Rosenthal, Drug Metab. Disposition 1, 21 (1973). 64. D. Y. Cooper, H. Schleyer, and 0. Rosenthal, Hoppe-Seyler’s 2.Physiol. Chem. 349, 1592 (1968). 65. F. Mitani and S. Horie, J . Biochem. (Tokyo) 88,139 (1969). 66. C. R. Jefcoate, R. Hume, and G. S. Boyd, FEBS (Fed. Eur. Biochem. Soc.) Lett. 9, 41 (1970). 67. J. Alfano, A. C. Brownie, W. H. OrmeJohnson, and H. Beinert, JBC 248, 7860 ( 1973). 68. H. Schleyer, D. Y. Cooper, S. S. Levin, and 0. Rosenthal, in “Biological Hydroxylation Mechanisms” (G. S. Boyd and R. M. S. Smellie, eds.), p. 187. Academic Press, New York, 1972.

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The redox potential of the cytochrome was found to be -400 & 10 mV in a Triton-solubilized preparation ( 6 9 ) .A K , of 9 p M for oxygen in the reconstituted llp-monooxygenase system has been reported ( 7 0 ) . The reduction of the hemoprotein follows biphasic kinetics. Deoxycorticosterone stimulates the initial phase while pregnenolone is inhibiting (70). The electron transport components were precipitated from the supernatant of the sonicated extract by ammonium sulfate (25-60%) and further purified by chromatography on Sephadex G-100 and DEAE-cellulose. A detailed procedure for the preparation of the iron-sulfur protein (ISP) is described by Kimura (71) and for the ISP reductase the method of Omura et al. (59,72) and modifications of it are used (73,74). The ISP has a molecular weight of 19,000 and shows the typical strong EPR signal a t g = 1.94 in the reduced state. The amino acid sequence of the 114 amino acids has been determined (75). It is assumed that the active site contains two iron atoms bridged by inorganic sulfide and bound to four cysteine residues. Adrenodoxin has been shown to be a one-electron carrier (76) and can be reduced by dithionite or by NADPH via the adrenal flavoprotein. The presence of oxygen readily restores the spectrum of the oxidized pigment, which has absorptions a t 320, 415, and 455 nm. I n the reduced state these bands are bleached and a new peak appears a t 550 nm with an isosbestic point at 306 nm ( 7 6 ) .Pure preparations show a ratio in optical density of 0.8 a t 415 nm relative to 280 nm. The most recent value of the oxidation-reduction potential is -270 mV ( 7 6 ) ) compared to earlier data of -305 mV (69) and -367 mV (77). The NADPH-dependent reductase has a molecular weight of about 54,000 (74) to 60,000 (64) as determined by Sephadex filtration and contains one mole of FAD per mole of enzyme and 2% (w/w) of hexose. The millimolar extinction coefficient is E455 nn, = 11.0 mM-l cm-l ( 6 4 ) . The oxidation-reduction potential is -274 mV a t p H 7.0, and the amino acid composition shows a high content of hydrophobic residues ( 7 4 ) . 69. D. Y. Cooper, H. Schleyer, and 0. Rosenthal, Ann. N . Y . Acad. Sci. 174, 205 (1970). 70. N. Ando and S.Horie, J . Biochem. ( T o k y o ) 70, 557 (1971). 71. T. Kimurtt, Struct. Bonding (Berlin) 5, 1 (1968). 72. T. Omura, E. Sanders, D. Y. Cooper, and R. W. Estabrook, “Methods in Enzymology,” Vol. 10, p. 362, 1967. 73. F. Mitani and S. Horie, J . Biochem. ( T o k y o ) 68,529 (1970). 74. J . W. Chu and T. Kimura, JBC 248, 2089 (1973). 75. M. Tanaka, M. Haniu, K. T. Yasunobu, and T. Kimura, JBC 248, 1141 (19731. 76. J. J. Huang and T. Kimura, Biochemistry 12, 406 (1973). 77. R. W. Estabrook, Int. Meet. Magn. Resonance, 1968 (1968).

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The steroid 11P-monooxygenase system could be reconstituted by appropriate combination of the three components in the presence of NADPH and oxygen (69). A large molar excess of the ISP over the other components was required (69) suggesting the reduction of the cytochrome as the rate-limiting step. An antibody preparation to homogeneous ISP strongly inhibited the overall hydroxylation as well as the reduction of cytochrome P-450 and cytochrome c (78). Spinach ferredoxin and chemical reductants were not able to replace adrenodoxin (79). Chu and Kimura (80) have demonstrated a tight complex formation between adrenodoxin and the reductase, but not between the ISP and the hemoprotein as described for the camphor 5-monooxygenase system (66). An oxygenated complex with absorptions a t 418 and 555 nm and a broad band around 580 nm has been described (68). It could be prepared by anaerobic chemical reduction of the hemoprotein which then was allowed to react with oxygen. This intermediate is only stable in the absence of substrate and reduced ISP because in their presence hydroxylated product will be formed. Huang and Kimura (79) stoichiometrically reduced a cytochrome P-450-substrate complex with dithionite and recorded a difference spectrum in the presence of oxygen. They attributed the absorption peak a t 430 nm to the oxygenated substrate complex in analogy to the reports of oxygenated complexes from the bacterial or liver microsomal monooxygenase systems (51,81). Further experiments showed that hydroxylation can be achieved with this chemically reduced oxygenated intermediate when the second electron is supplied by reduced ISP (68). The first electron can be provided by many reductants, while the second reduction step is absolutely speci,fic for the ISP (68,79). From our present understanding the 1lp-monooxygenase reaction in adrenocortical mitochondria follows similar mechanisms as established for the bacterial camphor monooxygenase system. c. Cholesterol Side Chain Cleaving Monooxygenase. The hemoprotein catalyzing the side chain cleavage of cholesterol in adrenocortical mitochondria seems to be a distinct form of cytochrome P-450 in these organelles, which may be solubiliaed and purified more easily than the llpmonooxygenase. It cleaves off the cholesterol side chain after monooxygenation in 20a and 22R position, yielding isocaproic aldehyde and pregnenolone (82). 78. J. Baron, W. E. Taylor, and B. S. S. Masters, ABB 150, 105 (1972). 79. J. J. Huang and T. Kimura, BBRC 44, 1085 (1971). 80. J. W. Chu and T. Kimura, JBC 248, 6183 (1973). 81. R. W. Estabrook, A. G. Hildebrandt, J. Baron, K. J. Netter, and K. Leibman, BBRC 42, 132 (1971). 82. C. Takemoto, H. Nakmo, H. Sato, and B. I. Tamaoki, BBA 152,749 (1968).

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Several purified hemoprotein preparations from adrenal mitochondria have been reported which are low in or free of llp-monooxygenase activity (66,83,84). A molecular weight of 800,000 was determined for a preparation which showed low llp- and no 18-hydroxylation activity (83), but no information on the specific content of cytochrome P-450 or the size and number of subunits has been reported. The rate of reduction of cytochrome P-450 in the presence of CO, the flavoprotein, and ISP was measured in the preparation free of llp- and 18-hydroxylation, and the biphasic reaction could be stimulated by the addition of cholesterol and decreased by 20a-OH-cholesterol ( 8 4 ) . The biphasic rate of the reduction is thought to result from endogenous substrate in the fast phase, while in the slow phase the rate might be limited by diffusion of added substrate (86). An EPR high-spin signal of this preparation at g = 7.9 a t 100°K has been reported which could not be increased by addition of cholesterol but was eliminated by addition of 20a-OH-cholesterol, suggesting that endogenous cholesterol is bound to the hemoprotein in saturating amounts. On the other hand, Alfano et al. (67) reported that their findings are in agreement with the concept that the g = 7.9 signal corresponds to the llp-monooxygenase cytochrome and the g = 8.2 signal to the cholesterol side chain cleaving hemoprotein. in its high-spin form. Only better purification of the different monooxygenase systems will eliminate the discrepancies between the effects of different substrates on the spectra and spin states of the various preparations. 2. Flavoproteins as Electron Donors

a. Monooxygenase Systems for Xenobiotics. Lipophilic organic compounds can readily enter the organism and would accumulate in membranes and adipose tissue if no monooxygenase system existed to transform these “xenobiotics” (86) into more polar metabolites by the introduction of an oxygen atom. This route of metabolism was early recognized by studying the nature of the excretion products in the urine (87), but only in the mid-1950’s could it be shown that a corresponding drug monooxygenase activity was mainly located in the endoplasmic reticulum of the liver (7). Meanwhile, the occurrence of monooxygenase activities for xenobiotics has also been established in lung (88), small in83. M. Shikita, P. F. Hall, and s. Isaka, BBRC 50, 289 (1973). 84. J. Ramseyer and B. W. Harding, BBA 315,306 (1973). 85. C. R. E. Jefcoate and J. L. Gaylor, Biocheinktry 9, 3816 (1970). 86. H. S. Mason, J. C. North, and M. Vanneste, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 24, 1172 (1965). 87. R. T. Williams “Detoxification Mechanisms,” 2nd ed. Chapman & Hall, London, 1959. 88. H. Uehleke, Xenobioticu 1, 327 (1971).

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testine (80,90), skin (91), and kidney (96).Except for some N-oxidations all monooxygenase reactions in drug metabolism are mediated by cytochrome P-450 species. A vast number of contributions have increased, but also sometimes masked, our understanding of this rather complex system. Only those facts that have been reported on a more biochemical than pharmacological background are presented. Direct evidence for the participation of the hemoprotein cytochrome P-450 in liver microsomal monooxygenations has been presented by the demonstration of the photochemical action spectra of some carbon monoxide-inhibited monooxygenase reactions (93,94),However, this role was already strongly suggested on the basis of the pioneering work of Omura and Sat0 ( l a , l 3 ) on the properties of the liver microsomal CO binding pigment. Starting from the observations of Klingenberg (9) and Garfinkel ( l o ) , Omura and Sat0 solubilized the GO binding pigment by treatment of the microsomal membranes with steapsin (crude pancreas lipase) , heated snake venom, and deoxycholate. Under these conditions, however, the characteristic absorption band a t 450 nm of the reduced CO spectrum was stoichiometrically converted to a new spectrum with a CO absorption peak in the reduced state a t 420 nm ((‘cytochrome P-420”).Cytochrome b, was released from the microsomal membranes by steapsin treatment (0.14%) after 1 hr at 37O and a subsequent incubation with sodium deoxycholate and heat-treated snake venom (both at a concentration of 0.1%) a t Oo for 36-40 hr under nitrogen completely solubilized cytochrome P-420.Ammonium sulfate fractionation (2545%), calcium phosphate gel adsorption and Sephadex G-100 filtration yielded a 5- to 6-fold purification of the hemoprotein. Its prosthetic group consisted of protoporphyrin IX which could be cleaved from the protein in cold acetone acidified with hydrochloric acid. Aggregation was observed above pH 7.5.The spectral properties of cytochrome P-420 are typical of a b-type cytochrome (see Table 111). Assuming a quantitative conversion of cytochrome P-450 into P-420 a molar extinction of 91 mM-l between 450 and 490 nm was derived for 89. L. W. Wattenberg, J. L. Leong, and J. P. Strand, Cancer Res. 22, 1120 (1962). 90. Ch. Lehrmann, V. Ullrich, and W. Rummel, Naunyn-Schmiedeberg’s Arch. Pharmakol. 276, 89 (1973). 91. A. P. Alvares, S. Leigh, A. Kappas, W. Levin, and A. H. Conney, Drug Metab. Disposition 1, 386 (1973). 92. S. Orrenius, A. Ellin, S. V. Jakobsson, H. Thor, D. L. Cinti, J. B. Schenkman, and R. W. Estabrook, Drug Metab. Disposition 1, 350 (1973).

93. D. Y. Cooper, S. S. Levin, S. Narasimhulu, 0. Rosenthal, and R. W. Estabrook, Science 147, 400 (1965). 94. H. Diehl, S. Capalna, and V. Ullrich, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 4, 99 (1969).

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TABLE 111 SPECTRAL CHARACTERISTICS OF ISOLATED LIVERMICROSOMAL CYTOCHROME P-420a

a

Spectrum

Wavelength (nm)

Extinction (cm-1 mM-l)

Oxidized (Soret) Oxidized Reduced (Soret) Reduced (8) Reduced ( a ) GO compound GO compound GO compound CO compound minus reduced

414 535 427 530 559 42 1 538 565 420-490

124 11 149 13 24 213 17 16 111

From Omura and Sat0 (IS).

cytochrome P-450 in the difference spectrum. On the basis of this value the cytochrome P-450 content of liver microsomes was calculated to be of the order of 0.4-1.5 nmole/mg of protein depending on the species or up to 7 nmoles/mg when the animals had been pretreated with phenobarbital (95). However, it should be mentioned that different values of the extinction coefficient have been reported depending on the pretreatment of the animals (96-98). Indeed, if various species of cytochrome P-450 with slightly different maxima of the CO complexes do exist (see below) the absorption difference between 450 and 490 nm can only be an average value. The oxidized cytochrome P-420 obtained by steapsin treatment is in the high-spin state and shows an EPR signal a t g = 6 which is enhanced in the presence of fluoride (99). Its redox potential is about -20 mV at pH 7, and it is readily autoxidizable under partial destruction of the heme group. The reduction by NADPH or NADH in the presence of microsomes is very slow, and no monooxygenase activity of cytochrome P-420 can be detected (1.3). Therefore, efforts have been directed toward the solubiliaation of intact cytochrome P-450, which was achieved by the use of deoxycholate (100) or nonionic detergents (101, 102). Treatment by deoxycholate also caused a conversion of microsomal 95. R. Sato, H. Satake, and Y. Imai, Drug Metab. Disposition 1, 6 (1973). 96. A. Hildebrandt, H. Remmer, and R. W. Estabrook, BBRC 30, 607 (1968). 97. T. Fujita, D. W. Shoeman, and G. J. Mannering, JBC 248, 2192 (1973). 98. J. B. Schenkman, H. Remmer, and R. W. Estabrook, Mol. Pharmacol. 3, 113 ( 1967). 99. K. Murakami and H. S. Mason, JBC 242, 1102 (1967). 100. A. Y. H. Lu, K. W. Junk, and M. J. Coon, JBC 244, 3714 (1969). 101. Y. Hashimoto, T. Yamano, and H. S. Mason, JBC 237, PC3843 (1962). 102. T. Fujita and G. J. Mannering, JBC 248, 8150 (1973).

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V. ULLRICH AND W. DUPPEL P-450 soluble +

P-450 particles

glycerol

E x r e s e n t P i p a s e detergents

P- 450 microsomal + nonionic hterg/

P-420 ‘‘ Fe

k 0 t - s

P-420 soluble, hgh spin

FIG.4. Effect of various treatments on microsomal cytochrome P-450.

cytochrome P-450 to the P-420 form; however, this preparation still contained the low-spin EPR signal which had been attributed to the intact cytochrome (“Fe,”) (101,103). The effects of various treatments of membrane-bound cytochrome P-450 are summarized in Fig. 4. Glycerol has been reported to stabilize cytochrome P-450 a t a concentration of 20-30% (104), and a combination of nonionic detergents and glycerol buffer medium allowed a resolution of the microsomal monooxygenase system and its reconstitution with retention of activity (100). The cytochrome was soluble by the criterion that it remained in the supernatant fraction upon centrifugation for 2 hr a t 105,000 g. The preparation was essentially free of cytochrome P-420 and contained only small amounts of cytochrome b, (105). The g values in the EPR spectrum of the oxidized cytochrome were a t g, = 1.92, gu = 2.25, and y, = 2.42 and corresponded to those of microsomal Fe, (100). The molecular weight has been found to be around 350,000 (106) ; however, gel electrophoresis in sodium dodecyl sulfate and mercaptoethanol suggests that this represents a complex of several polypeptide chains, each with a molecular weight of about 53,000-55,000 (107). Whereas the cytochrome P-450 concentration of the solubilized fraction was about the same as in microsomes, the NADPH-cytochrome P-450 reductase was about 100-fold purified when tested by its ability to catalyze cytochrome c reduction. Only a 10-fold increase, however, was observed for the reduction of solu103. Y. Miyake, J. L. Gaylor, and H. S. Mason, JBC 243, 5788 (1968). 104. Y. Ichikawa and T. Yamano, BBA 131, 490 (1967). 105. A. P. Autor, R. M. Kaschnitz, J. K. Heidema, T. A. van der Hoeven, W. Duppel, and M. J. Coon, Drug Metab. Disposition 1, 156 (1973). 106. A. P. Autor, R. M. Kaschnitz, J. K. Heidema, and M. J. Coon, Mol. Pharmacol. 9, 93 (1973). 107. M. J. Coon, T. A. van der Hoeven, R. M. Kaschnitz, and H. W. Strobel, Ann. N. Y. Acad. Sci. 212,449 (1973).

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ble cytochrome P-450, and this activity was very labile during purification and storage (100). Interestingly, the homogeneous NADPH-cytochrome c reductase prepared by lipase or trypsin treatment of microsomes (108) could not replace the detergent-solubilized form, although antibody experiments clearly showed that both proteins can cross-react (109). There is now evidence that the lipase- or trypsin-solubilized reductase lacks a hydrophobic part of the protein which may be responsible for the attachment of the reductase to the endoplasmic membranes (110). With regard to its prosthetic group the reductase is unusual since it contains FAD together with F M N a t the catalytic site (111). The K, for NADPH of M the detergent-solubilized reductase has been determined as 2 x (100).

With the purified NADPH-cytochrome P-450 reductase and the soluble hemoprotein no activity for the demethylation of benzphetamine was observed unless a lipid fraction also eluted from the column was added. Analysis of the components of this fraction revealed a remarkable specificity of the monooxygenase system for phosphatidylcholine (112). The effect of this phospholipid could be attributed to an enhancement of cytochrome P-450 reduction which proceeded with about a 16-fold higher first-order rate constant in the presence of phosphatidylcholine. The addition of about 0.1 mg of lipid extract to 1.0 ml of a reaction mixture containing the reconstituted system was necessary in order to support the monooxygenation of various drug substrates a t an optimal rate. Although the reconstitution of the microsomal monooxygenase system has very much clarified the nature of the system, there are more aspects with regard t o the function and mechanism that can only be studied in intact liver microsomes. According to isoelectric focusing the cytochrome P-450 fraction could be separated into a number of hemoproteins (113). The existence of multiple forms of cytochrome P-450 had already become apparent from studies on the substrate interactions of intact microsomes which are described next. Spectral investigations on membrane-bound cytochrome P-450 have been facilitated by special spectroscopic techniques for turbid solutions as developed by B. Chance. With the aid of difference spectroscopy most 108. C. H. Williams and H. Kamin, JBC 237, 587 (1962). 109. B. S. S. Masters, E. B. Nelson, B. A. Schacter, J. Baron, and E. L. Isaacson, Drug Metab. Disposition 1, 121 (1973). 110. T. Omura, Abstr., Znt. Congr. Biochem., Bth, 1973 p. 249 (1973). 111. T. Iyanagi and H. S. Mason, Biochemistry 12,2297 (1973). 112. H. W. Strobel, A. Y. H. Lu, J. Heidema, and M. J. Coon, JBC 245, 4851 (1970). 113. A. H. Conney, A. Y. H. Lu, W. Levin, A. Somogyi, S. West, M. Jacobson, D. Ryan, and R. Kuntzman, Drug Metab. Disposition 1, 199 (1973).

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A

Fm. 5. Difference spectrum of liver microsomes from phenobarbital-treated rats after addition of increasing amounts of cyclohexane (118).

of the characteristic spectral features of microsomal cytochrome P-450 had been described before absolute spectra of the solubilized cytochrome P-450 preparations of liver or other tissues had been available. After the observation by Narasimhulu (114) of a spectral change induced by substrates of the steroid 21-monooxygenase of adrenal microsomes, a similar difference spectrum was also obtained when drug substrates were added to oxidized microsomal suspensions (115,116). These substrate binding spectra were especially pronounced when the animals had been pretreated with phenobarbital to increase the content of cytochrome P-450 (117). In the difference spectrum one obtains a peak at about 388 nm and a trough a t 420 nm upon titration with increasing amounts of substrate to the microsomal suspension until saturation is obtained (115) (Fig. 5 ) . Measurements of the effect of substrate concentrations on the spectral change give a spectral dissociation constant which is usually identical to the K,,, value of the monooxygenase reaction (118).

If the substrate is removed by either washing (119) or evaporation 114. S. Narasimhulu, Fed. Proc., F e d . Amer. SOC. Exp. Biol. 22, 530 (1963). 115. H. Remmer, J. Schenkman, R. W. Estabrook, H. Sesame, J. Gillette, S. Narasimhulu, D. Y. Cooper, and 0. Rosenthal, Mol. Pharmacol. 2, 187 (1966). 116. Y. Imai and R. Sato, J. Biochem. (Tokyo) 62,239 (1967). 117. S. Orrenius, J. L. E. Ericsson, and L. Ernster, J . Cell Biol. 25, 627 (1965). 118. V. Ullrich, Hoppe-SeyZer's 2. Physiol. Chem. 350, 357 (1969). 119. J. B. Schenkman, H. Remmer, and R. W. Estabrook, MoZ. Pharmacol. 3, 113 (1967).

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(1.20) the substrate binding spectrum disappears and can be formed again by readmission of substrate. From these experiments the formation of an enzyme-substrate complex of microsomal cytochrome P-450 was suggested which is now generally accepted. It was also confirmed that the conversion of oxidized cytochrome P-450 with its Soret absorption band at about 418 nm to the substrate bound form with the Soret band a t 390 nm occurred concomitant with a transition from a low-spin state to a high-spin cytochrome (121). This was difficult to establish since concentrated microsomal suspensions as used for EPR spectroscopy require a much higher concentration for substrate saturation, and also only a part of the microsomal cytochrome will react with a given substrate. This again has led to the question whether cytochrome P-450 species with different substrate specificities exist. A clear proof for the presence of various species with different spectral properties was obtained by gel electrophoresis (122) and from substrate binding studies with 7-ethoxycoumarin (123). I n rat liver microsomes from untreated control animals no spectrum (and no activity) with 7-ethoxycoumarin was seen (Fig. 6) ; however, cyclohexane was bound and metabolized. In phenobarbital- and benzpyrene-pretreated animals qualitatively different substrate binding spectra were obtained with K , values that correspond exactly to the K,,, values of the O-dealkylation reaction (123). It could be expected that like other hemoproteins cytochrome P-450 in its reduced state would react not only with its physiological ligand dioxygen and the competitive inhibitor carbon monoxide but also with other compounds that can interact a t the iron. Table I V summarizes compounds that have been assumed to form ligand bonds to microsomal cytochrome P-450 either in the reduced or oxidized state (12,1S,119,124-128). A number of conclusions can be derived from these data. First, it seems to be a general phenomenon that ligands in the reduced state form com120. U. Frommer, V. Ullrich, and Hj. Staudinger, Hoppe-Seyler’s 2.Physiol. Chem. 351, 913 (1970). 121. M. R. Waterman, V. Ullrich, and R. W. Estabrook, ABB 155, 355 (1973). 122. A. F. Welton and S. D. Aust, BBRC 56, 898 (1974). 123. V. Ullrich, U. Frommer, and P. Weber, Hoppe-Seyler’s Z . Physiol. Chem. 354, 514 (1973). 124. H. Nishibayashi, T. Omura, and R. Sato, BBA 118, 651 (1966). 125. V. Ullrich, B. Cohen, D. Y . Cooper, and R. W. Estabrook, in “Structure and Function of Cytochromes” (K. Okunuki, M. D. Kamen, and I. Sekuzu, eds.), p. 649. Univ. of Tokyo Press, Tokyo, 1967. 126. V. Ullrich and K. H. Schnabel, ABB 159,240 (1973). 127. A. G. Hildebrandt, K. C. Leibman, and R. W. Estabrook, BBRC 37, 477 (1969). 128. C. R. E. Jefcoate, R. L. Calabrese, and J. L. Gaylor, Mol. Pharmocol. 6, 391 (1970).

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- 0.02

i

1

I '

i

'J

228

- 0.01 FIG.6. Substrate binding difference spectra of 7-ethoxycoumarin with rat liver microsomes (123).

TABLE I V LIQAND-INDUCED DIFFERENCE SPECTRA OF MICROSOMAL CYTOCHROME P-450 Soret bands Ligand

co C N-CPHI NO Fluorenylcarbanion Metyrapone Octylamine Octylmercaptane Methyloctylsulfide

Oxidized

Reduced

Ref.

-

450 430,455

12,15 119,124 186 186 187

436 423 427-432 378,471 436

446 446 442 449 448

188 0

W. Nastainczyk, H. H. Ruf, and V. Ullrich, unpublished.

plexes which in the Soret region absorb a t 450 nm like carbon monoxide. The affinity constants for these ligand complexes are rather high. Second, the oxidized form of the cytochrome also has access to ligands which cause difference spectra with troughs around 390-400 nm and peaks be-

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IRON- AND COPPER-CONTAINING MONOOXYGENASES

277

tween 420 and 470 nm depending on the ligand. Ligand-forming compounds can compete effectively with the binding of substrates as well as of oxygen and therefore should generally be inhibitory for microsomal monooxygenase reactions. However, because of the suggested heterogeneity of cytochrome P-450 this is not valid for all monooxygenase reactions. Thus, liver microsomes obtained from benxpyrene-induced animals contain a cytochrome P-450 species which has a very low affinity to most ligands and therefore is not inhibited by these compounds (123). Also a compound like aniline may form an enzyme-substrate complex with one species and a ligand complex with another resulting in superimposed spectra ( l a g ) . Another difficulty in the interpretation of difference spectra is the occurrence of endogenous enzyme-substrate complexes which may be converted back to the low-spin state by organic compounds like aliphatic alcohols. The resulting spectral change would then appear as a reversed substrate binding spectrum (130). I n summary difference spectra with microsomal suspensions depend very much on the pattern of cytochrome P-450 species present. This pattern may be dependent on species, sex, age, environmental factors, and pretreatment, or it may even change upon solubilization or storage of the microsomes. This in part answers the question concerning the substrate specificity of microsomal monooxygenases. I n general, lipophilic compounds with acceptor groups for an electrophilic “oxenoid” species (see Section III,A,3,c) can be substrates but their monooxygenation rates very much depend on the pattern of cytochrome P-450 species present. The isolation of the single cytochrome P-450 species in a functional state and the determination of their individual substrate specificities has not yet been achieved. The formation of an enzyme-substrate complex generally represents a first fast step in an enzymic reaction and also occurs very rapidly a t cytochrome P-450. The subsequent reduction of the cytochrome-substrate complex in the presence of NADPH is rather slow compared to electron transfer reactions at cytochromes. The reduction can be followed by rapid mixing of a carbon monoxide gassed microsomal suspension with a n anerobic NADPH solution (131). The presence of substrate considerably changed the kinetics of the reduction (Fig. 7 ) . Since obviously the resulting curve was composed of two reactions, the two different rate constants were obtained by curve fitting (130). In the case of phenobarbital-pre129. J. B. Schenkman, D. L. Cinti, S. Orrenius, P. Moldeus, and R. Kaschnite, Biochemistry 11, 4243 (1972). 130. H. Diehl, J. Schadelin, and V. Ullrich, Hoppe-Seykr’s Z . Physiol. Chem. 351, 1359 (1970). 131. P. L. Gigon, T. E. Gram, and J. R. Gillette, M o l . Pharmacol. 5, 109 (1969).

278

-.-= -.--

V. ULLRICH AND W. DUPPEL

I

-ftnal

level after 5 m h

++-

0

2

1

8

6

tlsed

10

--.-*

!

c-

12

t

Fro. 7. Kinetics of reduction by NADPH of microsomal cytochrome P-450 in the presence of carbon monoxide. Dashed curve : without added substrate ; solid curve : lo-' M cyclohexane added (130).

treated liver microsomes in the presence of cyclohexane the first rapid phase proceeded with a first-order rate constant of 66 min-l and corresponded to about 47% of the cytochrome P-450. Without substrate most of the cytochrome reacted slowly with a second-order rate constant of about 9.6 nmole-* x min-l and only about 17% of the fast phase was present. This small amount of rapid phase possibly resulted from endogenous substrate. From the spectral change obtained with cyclohexane in these liver microsomes it was estimated that 32 2 6% of the cytochrome was converted to the eneyme-substrate complex. The hydroxylation of cyclohexane is easy to follow since only one hydroxylation product can be formed which can be determined by gas-liquid chromatography (118). I n a typical experiment an activity of 16.6 nmoles X min-' per nmole of total cytochrome P-450 was calculated, which, on the basis of 32% eneyme-substrate complex, resulted in a molecular activity of 52 nmoles X min-l. This corresponds very closely to the first-order rate con-

5.

279

IRON- AND COPPER-CONTAINING MONOOXYGENASES

NADH

1

NAOPH

4

A

Uncoupling

I

Monooxygenation

FIG.8. Electron transport during microsomal monooxygenation and uncoupling reactions (132).

stant of 66 min-l for the reduction of the enzyme-substrate complex and allows the conclusion that the reduction of the cytochrome P-450 substrate complex is the rate-limiting step in the overall sequence of microsoma1 monooxygenases. Cyclohexane also proved to be a suitable substrate for establishing the postulated 1 :1 : 1 stoichiometry for NADPH uptake, O2consumption, and product formation (132).During the course of these investigations on the stoichiometry it was found that lipophilic substrates without hydroxylatable groups like perfluorinated hydrocarbons stimulate NADPH oxidation ( 1 3 3 ) ,but only 0.5 equivalent of oxygen is consumed and water formed. This “uncoupling” phenomenon is observed to some extent even with normal substrates like n-hexane leading to a less than stoichiometric oxygen consumption and product formation compared to NADPH oxidation (132).Interestingly, NADH via cytochrome b, can provide the third and fourth electron needed to reduce the “active oxygen” species which is not incorporated into the substrate (Fig. 8 ) . This electron sparing effect can account for most of the synergistic action of NADH on microsomal monooxygenations (134-136). Again, the kinetic data reported here only apply for liver microsomes from phenobarbital-pretreated rats or rabbits and for cyclohexane as a substrate. The kinetics for other substrates may differ, and certainly they are different for microsomes from animals pretreated with polycyclic hydrocarbons or other inducers. b. Other Monooxygenase Systems. A second bacterial cytochrome 132. H. Staudt, F. Lichtenberger, and V. Ullrich, Eur. J . Biochem. 46, 99 (1974). 133. V. Ullrich and H. Diehl, Eur. J. Biochem. 20, 509 (1971). 134. B. S. Cohen and R. W. Estabrook, ABB 143, 54 (1971). 135. M. A. Correia and G. J. Mannering, Mol. Pharmacol. 9, 445 (1973). 136. H. A. Sasame, J. R. Mitchell, S. Thorgeirsson, and J. R. Gillette, Drug Metab. Disposition 1, 150 (1973).

280

V. ULLRICH AND W. DUPPEL

P-450-containing monooxygenase has been characterized from Corynebacterium sp. (137). This system is induced by growth of the organism on n-octane and converts this substrate to n-octanol. The cytochrome P-450 is membrane bound and needs a soluble flavoprotein fraction for the reduction by NADH. Tetradecane-related n-alkanes and higher fatty acids are substrates for a monooxygenase system of the yeast Candida tropicalis (138).The enzyme system is located in the microsomal fraction (139) and closely resembles the mammalian drug monooxygenase system since it could be resolved into a cytochrome P-450 containing terminal monooxygenase, a flavin-dependent reductase, lysophosphatidylethanolamine as a lipid cofactor, and NADPH as a reductant. In endosperm preparations of immature seeds of Echinocystis macrocarpa a particulate cytochrome P-450 type of monooxygenase could be found that converts kaur-16-ene to kaur-16-en-19-01 in the biosynthetic pathway of the plant growth hormone gibberellin with NADPH as an electron donor (140). In the fungus, Claviceps purpurea, a hemoprotein of the cytochrome P-450 type is believed to be involved in the biosynthesis of alkaloids, but direct participation in a certain monooxygenation step has not yet been shown (141). Also unknown is the function of a cytochrome P-450 species in the nitrogen-fixing bacteroids, Rhizobium japonicum (1.42). 3. General Mechanisms of Heme-Containing Monooxygenmes

a. Structure of Cytochrome P-450. The correlation of the unusual spectral data of the various cytochrome P-450 species and their functional states with the ligand field of the iron is one of the most challenging problems in heme chemistry. It is obvious that some fundamental differences exist to other hemoproteins. First, the unusual EPR spectra of the P-450 hemoproteins have to be explained on the molecular basis of the ligand field around the iron. The narrow splitting of the EPR signal of oxidized low-spin cytochrome P-450 suggests a high degree of rhombicity (37) which is thought to result from the presence of a soft ligand, probably a mercaptide. There is not yet direct evidence for this assumption, but models obtained from metmyoglobin or methemoglobin and mer137. G. Cardini and P. Jurtshuk, JBC 245,2789 (1970). 138. W. Duppel, J. M. Lebeault, and M. J. Coon, Eur. J . Bzochem. 36, 583 (1973). 139. M. Gallo, B. Roche, L. Aubert, and E. Azoulay, Biochimie 55, 195 (1973). 140. P. J. Murphy and C. A. West, ABB 133, 395 (1969). 141. S. H. Ambike, R. M. Baxter, and N . I3 Zahid, Phytochemktry 9, 1953 (1970). 142. C. A. Appleby, BBA 172, 71 (1969).

5.

281

IRON- AND COPPER-CONTAINING MONOOXYGENASES

Metmyoglobin + CIHpSH

Rat Liver Microsomes

2.5 T = 110.K

3.0 Magnetic Field

3.5 kGauos B

b -.

FIG.9. EPR spectra of microsomal cytochrome P-450 in comparison to metmyoglobin in the presence of butylmercaptane (145).

282

V. ULLRICH AND W. DUPPEL

captanes have led to exactly the same EPR signals (143,144) [see Fig. 9 (146)I. Also in agreement with a mercaptide sulfur in the fifth position would be the strongly negative redox potential of about -400 mV of the different cytochrome P-450 species (.!l8,69,146)which matches the potentials of most iron-sulfur proteins. The reduced form of the hemoprotein has been characterized high-spin by Mossbauer spectroscopy (54). A second special feature common to all cytochrome P-450 species is the strong red shift of the carbon monoxide complex in the reduced state. Other ligands with free electron pairs like carbanions, amines, mercaptides, or isocyanides (see Table IV) produce the same type of ligand spectra. By denaturation this absorption band is rapidly shifted to about 420 nm concomitant with a loss in monooxygenase activity. Interestingly, the 450-nm band of the reduced ligand spectrum seems to be unrelated to the unusual EPR spectrum since a low-spin P-420 (Fe,) can be obtained under mild denaturating conditions. The final degradation state of the cytochrome (“cytochrome P-420”) is high spin, and its spectral characteristics are no different from those of other hemoproteins. A tentative explanation for the unusual properties of cytochrome P-450 would be that the heme group is attached to the apoprotein by protein-porphyrin interaction as well as by a mercaptide-iron linkage. A stepwise inactivation of the monooxygenases could affect the heme binding sequentially, although the conditions under which this will occur are dependent on the individual cytochrome P-450 species. b. Substrate Binding. A third characteristic feature of all heme-containing monooxygenases is the dramatic change of the optical absorption spectrum and of the magnetic properties upon binding of the corresponding substrates. The attachment of the substrate a t the active site probably causes a conformational change which then triggers the conversion to a high-spin enzymesubstrate complex showing g values a t about 8, 4, and 2. The Soret band shifts from about 418 to 390 nm, and also a typical 645-nm absorption band appears. There is a remarkable variation in the degree of specificity among the various cytochrome P-450 species. In the case of the steroid llp- and 21-monooxygenase only 143. C. R. E. Jefcoate and J. L. Gaylor, Biochemistry 8, 3464 (1969). 144. H. A. 0. Hill, A. Roder, and R. J. P. Williams, Struct. Bonding (Berlin) 8, 123 (1970). 145. V. Ullrich and Hj. Staudinger, in “Handbuch der experimentellen Pharmakologie” (B. B. Brodie and J. R. Gillette, eds.), New Ser., Vol. 28, Part 2, p. 251. Springer-Verlag, Berlin and New York, 1971. 146. M. R. Waterman and H. S. Mason, BBRC 39, 450 (1970).

5.

IRON- AND COPPER-CONTAINING MONOOXYGENASES

283

structurally closely related compounds will form the ensyme-substrate complex and only one product is formed with high stereospecificity. On the other hand, the drug monooxygenases in liver, especially after phenobarbital or polycyclic hydrocarbon pretreatment, interact with many lipophilic compounds and usually lead to a pattern of products. Even if one considers several cytochrome P-450 species with overlapping substrate specificities, the spectrum of compounds that act as substrates is extremely broad. Compounds with basic nitrogen atoms can also bind effectively to the active site competing with the substrates, I n this case the low-spin state is retained and the Soret absorption band shifts to slightly higher wavelengths. This may be indirect proof for the addition of substrates near the sixth position and not at the mercaptide site. One may further speculate that the low-spin ferric cytochrome has a nitrogen ligand in the sixth coordination position which is removed after binding of a substrate molecule. Indeed, the isoelectric point of the camphor-bound cytochrome P-450 complex is higher than that of the low-spin cytochrome suggesting the release of a weakly basic group (39). The increase in redox potential after binding of substrates would explain why the cytochrome is reducible only in the presence of substrate. It also points to a simple but effective regulation mechanism of coupling electron transport and monooxygenase activity. c. Oxygen Activation. After reduction of the cytochrome-substrate complex the ferrous ion rapidly binds a dioxygen molecule and forms an oxy complex. This reaction is analogous to the oxygenation of hemoglobin or myoglobin. The addition is reversible since carbon monoxide can displace the oxygen molecule under formation of the characteristic CO complex (53).The actual “active oxygen” complex is formed by the transfer of a further electron to the oxygenated cytochrome. This follows from the stoichiometry of the monooxygenation reaction and from an experiment with the cytochrome of the 1 lp-monooxygenase system. It has been shown that no hydroxylation of the steroid substrate occurs when stoichiometric amounts of oxygen are added to the dithionite-reduced cytochrome, but product is formed upon subsequent addition of the reduced iron-sulfur protein (adrenodoxin) (68).Moreover, the chemical features of the oxy complexes of hemoproteins are inconsistent with an [FeO2I2+form being the active species. In addition, model systems for the oxygen activation strongly suggest that only after a two-electron transfer to the oxygen molecule is a highly reactive hydroxylating species formed. This is obvious in a system consisting of a stannous phosphate complex and oxygen in an aqueous solution in which hydroxylations of

+8

Fe3@

[Fe -OCO]@

+ e0 t-

V. ULLRICH AND W. DUPPEL Fez@

[Fe3Q0p]2@

FIQ.10. Proposed scheme for the iron-catalyzed monooxygenation reaction (148).

organic compounds (RH) proceed by the following equation under incorporation of molecular oxygen (147,148). Sna+

+ RH +

H+ Sn‘+

+ ROH + OH-

(3) With regard to reactivity and products formed the monooxygenase reactions are best mimicked by oxidations with trifluoroperacetic acid (149-161). It is assumed that these oxidations occur in a concerted mechanism with an electrophilic attack of the OH group (152). In analogy with the carbenoid reactions one could characterize the peracid reactions and hence also the monooxygenations as “oxenoid” reactions (148,165). Since a strong polarization of the 0-0 bond in a peroxidic complex seems to be sufficient to create an ‘Loxenoid”species the transition state for the monooxygenase reactions given in Fig. 10 was proposed (148). After the oxygen atom is transferred to the substrate, the product is released and the second oxygen atom after protonation is split off as water. During this process the low-spin state of the cytochrome is re-formed and can enter the monooxygenation cycle again. The complete sequence of the reaction steps is schematically presented in Fig. 11 (130,164). d. Inhibitors. Most commonly known inhibitors interact a t the heme moiety of the cytochromes. They can either prevent the formation of the enzyme-substrate complex by blocking the active site or they can interfere with the reaction of oxygen a t the ferrous ion. Substrate analogs belong to the first type and carbon monoxide is characteristic for the latter. Very often both effects come together when the inhibitor is a lipophilic organic molecule that fits into the active site and in addition has 0 2

+

147. V. Ullrich and Hj. Staudinger, Z . Naturforsch. B 24, 583 (1969). 148. V. Ullrich and Hj. Staudinger, in “Biochemie des Sauerstoffs” (B. Hem and Hj. Staudinger eds.), p. 229. Springer-Verlag, Berlin and New York, 1968. 149. D. Jerina, J. W. Daly, W. Landis, B. Witkop, and S. Udenfriend, JACS 89, 3347 (1967). 150. V. Ullrich, J. Wolf, E. Amadori, and Hj. Staudinger, Hoppe-Seyler’s Z . Physiol. Chem. 349, 85 (1968). 151. U. Frommer and V. Ullrich, 2.Naturforsch. B 26, 322 (1971). 152. P. 0. Bartlett, Rec. Chem. Progr. 11, 47 (1950). 153. G . A. Hamilton, JACS 86, 3391 (1964). 154. V. Ullrich, Angew. Chem., Znt. Ed. Engl. 11, 701 (1972).

5.

IRON- A N D COPPER-CONTAINING MONOOXYGENASES

T

pyp

285

p* [!;14]1@

FIG.11. Proposed reaction cycle for cytochrome P450-dependent monooxygenases (164).

FIG. 12. Inhibitor interaction with the active site of cytochrome P-450 (166).

a free electron pair a t one atom to share with the iron (Fig. 12). These compounds bind with a rather high affinity as a result of the ligand interaction with the ferric ion and create a low-spin state of the cytochrome. With the reduced cytochrome the affinity is even higher and the spectra show the characteristic Soret band around 450 nm (155). Generally, inhibitors interact most effectively with the rate-limiting steps of an enzymic reaction. In case of the heme-containing monooxygenase this is usually the electron transport system for the reduction of the heme. No specific inhibitors for this part of the monooxygenase system have been found except antibodies for the corresponding flavoproteins. It seems, however, that under physiological conditions a regulation of monooxygenase activity makes use of the rate-limiting role of the reducing equivalents ( 1 5 6 ) .

B. NONHEME IRON-CONTAINING MONOOXYGENASES 1. 4-Methoxybenzoate O-Demethyl-Monooqgenase from P. putida

So far only a few iron-dependent monooxygenases are known which activate oxygen by nonheme iron proteins. One example is the 4-methoxy155. V. Ullrich and K. H. Schnabel, Drug Metab. Disposition 1, 176 (1973). 156. R. Scharf and V. Ullrich, Biochem. Pharmacol. 23, 2127 (1974).

286

V. ULLRICH AND W. DUPPEL

benzoate 0-demethyl-monooxygenase isolated from a P. putida strain grown on this substrate (167). The cofactor requirement and the following stoichiometry identifies the enzyme system as a monooxygenase : 4-Methoxybeneoate

+Oa+NADH+ H+ > 4-hydroxybeneoate

+ formaldehyde + NAD+ + Ha0

(4)

Two fractions were obtained after sonication of the bacterial suspension, ammonium sulfate precipitation of the supernatant, and subsequent chromatography on Bio-Gel P-150 and DEAE-cellulose (168). One fraction contained an iron-sulfur protein with an estimated molecular weight of 120,000 (169). Each of the two presumably identical subunits with a molecular weight of about 52,000 contains two iron atoms and two-acid-labile sulfur groups. The EPR spectrum of the reduced enzyme shows signals with g values a t 2.01, 1.91, and 1.78 (160), and the optical spectrum with a peak a t 453 nm in the oxidized state is also different from other iron-sulfur proteins (169). The redox potential was determined to be about +5 mV a t p H 7.8 (161). Substrate binding difference spectra in the visible region and CD spectra show two types of interaction of the protein with different classes of substrates depending on the substituent in para position of the carboxyl group (161) (Fig. 13). Since oxygen consumption is also affected by substrate binding (162),this iron-sulfur protein is believed to be the terminal oxidase in this system. The second fraction contains the reducing system which consists of a FMN flavoprotein associated with an iron-sulfur chromophore with a g value a t 1.94 (163) in the reduced state. After removal of the iron-sulfur protein by Sephadex A-50 chromatography the reductase can still reduce ferricyanide and 2,6dichlorophenolindophenol, but the activity toward cytochrome c as well as the 0-demethylation activity with the terminal oxidase are lost (163). The molecular weight of the reductase is 38,000 as estimated by gel filtra157. F. H. Bernhardt, Hj. Staudinger, and Y. Ullrich, Hoppe-Seyler’s Z . Physiol. Chem. 351, 467 (1970). 158. F. H. Bernhardt, H. H. Ruf, Hj. Staudinger, and V. Ullrich, Hoppe-Seyler’s Z. Physiol. Chem. 352, 1091 (1970). 159. F. H. Bernhardt and W. Nastainceyk, ZRCS (Biochem. Microbial.) 2, 1268 (1974). 160. H. H. Ruf, F. H. Bernhardt, V. Ullrich, and Hj. Staudinger, Abstr. Commun., 8th Meet. Fed. Eur. Bwchem. Soc., 197B No. 447 (1972). 161. F. H. Bernhardt, H. H. Ruf, and H. Ehrig, FEBS (Fed. Eur. Biochem. Soc.) Lett. 43, 53 (1974). 162. F . H. Bernhardt, N. Erdin, Hj. Staudinger, and V. Ullrich, Eur. J . Biochem. 35, 126 (1973). 163. F. H. Bernhardt and Hj. Staudinger, Hoppe-Seyler’s Z . Physiol. Chem. 354, 217 (1973).

5.

287

IRON- AND COPPER-CONTAINING MONOOXYGENASES

ID-

10 .

t

3

1 0.5 -

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

.

.;

,,'

\

.

n

0

I

n

P

O

x

E

;. -0.5 .5 D

4 -1.0 u

is -1.5

. . ::

400

500 A[nm]

-

600

(A)

LOO

500

x [nm]

-

600

iBi

FIO.13. Circular dichroism spectra of the terminal iron-sulfur protein of 4methoxybenzoate O-demethyl-monooxygenase with various substrates. (A) Curve 1, oxidized enzyme; curve 2, addition of 4-methoxybenzoate; and curve 3, dithionite added. (B) Curves correspond to ( A ) but substrate analog 4-trifluoromethylbenzoate added. For details, see Bernhardt et al. (161). tion and ultracentrifugation. The optical spectrum of the reduced iron-sulfur chromophore is similar to those of other iron-sulfur proteins of the ferredoxin type (164). The reducing system can be replaced by artificial electron donors like dithionite or indigodisulfonate (159). There is no pronounced substrate specificity, but the carboxyl group seems to be required for substrate binding. Substrate analogs like 4-trifluoromethylbenzoate which are bound to the enzyme but do not have a hydroxylatable CH bond uncouple the electron transport under formation of hydrogen peroxide (162). Nothing is known so far about. the molecular structure and the ligand field of the terminal iron-sulfur protein. It seems likely, however, that more monooxygenases of this type will. be found in the future. 2. Phenylalanine 4-Monooxygenase

Phenylalanine monooxygenase is found in mammalian liver and catalyzes the conversion of phenylalanine to tyrosine. The first in vitro SYStem was reported in 1913 (165),but the reaction mechanism remained 164. F. H. Bernhardt and H. Pachowsky, IRCS (Biochem. Microbiol.) 2, 1267 (1973). 165. G. Embden and K. Baldes, Biochem. 2 . 5 5 , 301 (1913).

288

V. ULLRICH AND W. DUPPEL

unclear for a long time. Kaufman and co-workers (166-168) found that the monooxygenase system requires a unique reductant, tetrahydropterin, which can only be replaced by a few structural similar pterins. The reduction of this cofactor is mediated by a NADPH-dependent reductase. The ultimate acceptor is a nonheme iron protein which catalyzes tyrosine formation according to the following scheme

Nyx:cDihydropterin

X;:Tine+%O

NADP+

Tetrahydropterin

(5)

Phenylalanine + 0,

In earlier studies the natural cofactor tetrahydropterin has been isolated from rat liver by organic solvent extraction with subsequent chromatography on Dowex 1-C1- and Dowex 50-Na’ columns (169). This natural cofactor can be substituted by 6,7-dimethyltetrahydropterin in the presence of mercaptoethanol. The reductase has been purified from sheep liver (169). The liver was homogenized in 0.03 M acetic acid and the 4000 g supernatant was subjected to ammonium sulfate fractionation. Further purification was achieved by a zinc-ethanol fractionation, a second ammonium sulfate precipitation, and chromatography on calcium phosphate gel, yielding a 80- to 100-fold purification. The terminal enzyme is prepared from rat liver (170).Its purification involves homogenization in 0.01 M acetic acid, ethanol and ammonium sulfate fractionation, chromatography on calcium phosphate gel, a second ammonium sulfate precipitation followed by chromatography on DEAEcellulose and Sephadex G-200. Two major forms of the enzyme with molecular weights of 110,000 and 210,000 could be identified, which on SDS polyacrylamide electrophoresis yielded subunits with a molecular weight of 51,000-55,000. Each of the subunits contained one mole of iron. Electron paramagnetic resonance measurements showed a high-spin ferric iron signal at g = 4.23, which is attributed to a rhombic symmetry of the iron environment (17’f). This signal disappeared on addition of substrate. The apoenzyme 166. S. Kaufman, Advan. Enzymol. 35,24 (1971). 167. S. Kaufman, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 261. Maruzen, Tokyo, 1966. 168. S. Kaufman in “Oxygenases” (0. Hayaishi ed.), p. 129. Academic Press,New York, 1962. 169. S. Kaufman and B. Levenberg, JBC 234, 2683 (1959). 170. S. Kaufman and D. B. Fisher, JBC 245, 4745 (1970). 171. D. B. Fisher, R. Kirkwood, and S. Kaufman, JBC 247, 5161 (1972).

5.

IRON- AND COPPER-CONTAINING MONOOXYGENASES

289

could be prepared by treatment with o-phenanthroline and cysteine. Monooxygenase activity was lost by this treatment but could be fully restored by the addition of Fez+, but other metals were ineffective. The activity of the phenylalanine monooxygenase system can be stimulated 20-fold by the addition of lysolecithin or a-chymotrypsin (172). The first compound induces conformational changes of the enzyme exposing a sulfhydryl group, while the latter partially hydrolyzes the protein and reduces its size to a dimer with a molecular weight of 67,000. The K, for phenylalanine is decreased by about 50% and the saturation curve for the substrate is converted from a sigmoidal to a hyperbolic form. I n contrast to the action of chymotrypsin the lysolecithin stimulation is reversible. Under certain conditions the specific activity of the monooxygenase decreases with increasing enzyme concentration (173). This effect can be overcome by the addition of a stimulatory protein, which has been purified 1000-fold. It has a molecular weight of 51,500 consisting of four identical subunits. This stimulatory protein greatly increases the affinity of the monooxygenase for tetrahydropterin, the natural reductant, without changing the V,,,. From kinetic experiments it is concluded that during the reaction an intermediary product is released which binds to the free enzyme, thus inhibiting the reaction. The stimulatory protein catalyzes the breakdown of this inhibitory intermediate to product and reverses the inhibition (1’74). The possible function of lysolecithin and the stimulatory protein in the in vivo regulation of the phenylalanine monooxygenation activity is discussed by Fisher and Kaufman (172). When the monooxygenase protein is partially altered by the addition of lysolecithin or a-chymotrypsin, the product of phenylalanine monooxygenation, tyrosine, uncouples the enzymic reaction (175). Under these conditions the enzyme oxidizes tetrahydropterin under formation of hydrogen peroxide, but no product is formed. The role of iron in the enzymic monooxygenation of phenylalanine is not fully established, but there are indications that it is directly involved at the catalytic site since the EPR signal of the oxidized iron a t g = 4.23 disappears on addition of substrate and reduced pterin (17’1). This could either result from reduction or from a change of the spin state of the iron. An oxoiron complex is believed to be the possible active oxidant in the hydroxylation reaction (171,176). 172. 173. 174. 175. 176.

D. B. Fisher and S. Kaufman, JBC 248,4345 (1973). C. Y. Huang, E. E. Max, and S. Kaufman, JBC 248, 4235 (1973). C: Y. Huang and S. Kaufman, JBC 248,4242 (1973). D. B. Fisher and S. Kaufman, JBC 248, 4300 (1973). K. B. Sharpless and T. C. Flood, JACS 93, 2316 (1971).

290

V. ULLRICH AND W. DUPPEL

3. Tyrosine 3-Monooxygenase A monooxygenase system similar to the phenylalanine system is found in the adrenal medulla and sympathetically innervated tissues catalyzing the hydroxylation of tyrosine to 3,4-dihydroxyphenylalanine(DOPA).

6

I

t

tetrahydropterin, NADPHtH+,O,_

@+

reductase

yH2 HCNH:

HCNH:

co;

co;

I

Qrosine

H,O

-

I I I

I

Norepinephrine

(6)

yH2 I

3,4-Dihydroxyphenylalanine

The enzyme represents the rate-limiting step in the biosynthesis of norepinephrine, and its activity is regulated by competitive feedback inhibition by norepinephrine (177’). The dihydropterin reductase regenerating the tetrahydropterin in the presence of NADPH can be purified from sheep liver and is similar to or even identical with that of the phenylalanine monooxygenase system (178). The terminal protein is obtained from adrenal medulla homogenates giving a crude granule fraction after centrifugation a t 24,000 g (179). From this fraction the enzyme is solubilized by digestion with chymotrypsin. After centrifugation the enzyme is further purified by ammonium sulfate precipitation, chromatography on Sephadex G-150, and a second ammonium sulfate fractionation. The molecular weight of the partially purified enzyme is reported to be about 40,000. The enzyme is specific for L-tyrosine, with a K,,, of 1 X 10-6 M and has a pH optimum a t 6.Cb6.2 (180).

A soluble preparation of the membrane-bound enzyme has been obtained from rat brain (181). Its molecular weight was estimated to be about 200,000. After limited tryptic digestion this preparation showed a marked alteration of the molecular weight (about 50,000) and a sub-

s.

177. M. Levitt, J. W. Gibb, J. W. Daly, M. Lipton, and Udenfriend, Biochem. Pharmacol. 16, 1313 (1967). 178. S. Kaufman, “Methods in Enzymology,” Vol. 5, p. 809, 1962. 179. R. Shiman, M. Akino, and S. Kaufman, JBC 246, 1330 (1971). 180. T. Nagatsu, M. Levitt, and S. Udenfriend, JBC 239, 2910 (1964). 181. R. T. Kuceenski and A. J. Mandell, JBC 247,3114 (1972).

5.

29 1

IRON- AND COPPER-CONTAINING MONOOXYGENASES

stantial increase in enzymic activity which may be caused by a conformational change after cleavage of certain peptide bonds (182). The previously reported requirement for Fez+ has been explained by the ability of Fez+to decompose H,O, which is generated by reoxidation of the tetrahydropterin and which inactivates the enzyme (179,185). Catalase or peroxidase act similarly ; however, in the trypsin-solubilized preparation of Petrack et al. (183) mercaptoethanol and Fez+ are required in addition to catalase to activate the enzyme during preincubation. This suggests an additional role of iron in tyrosine monooxygenase, and it can be assumed that the system and its mechanism are very similar to the phenylalanine monooxygenase system. 4. Tryptophan 5-Monooxygenase

Tryptophan 5-monooxygenase catalyzes the hydroxylation of tryptophan in the 5 position which represents the first step in the biosynthesis of serotonin. Ht* <&OH

H,C-CH-CO,-

+

L

H Tryptophan

tetrahydropterin, reductase

H 5 - Hydroxy tryptophan

H,O

(7)

\ Serotonin

The first report of this hydroxylation activity in homogenates of dog and rabbit brainstems was published by Grahame-Smith in 1964 (184). Mammalian brain is the main source for this enzyme, but it has also been found in Chromobacterium violaceurn (185), tumors (186), and neoplastic mast cells (187). The extremely low activity in vitro and the lack of a sensitive assay hindered the study of this enzyme for a long time. With the development of sensitive fluorometric (188) and radioactive tests (189) these difficulties have been overcome. The terminal protein of the monooxygenase system has been purified from brain after homogenization, centrifugation 182. R. Kucsenski, JBC 248, 2261 (1973). 183. B. Petrack, F. Sheppy, V. Fetser, T. Manning, H. Chertock, and D. Ma, JBC 247, 4872 (1972). 184. D. G. Grahame-Smith, BBRC 16, 586 (1964). 185. C. Mitoma, H. Weissbach, and S. Udenfriend, ABB 63, 122 (1956). 186. W. Lovenberg, E. Jequier, and A. Sjoerdsma, Science 155, 217 (1967). 187. S. Hosoda and D. Click, JBC 241, 192 (1966). 188. P. A. Friedman, A. H. Kappelman, and S. Kaufman, JBC 247, 4165 (1972). 189. A. Ichiyama, S. Nakamura, Y. Nishizuka, and 0. Hayaishi, JBC 245, 1699 (1970).

292

V. ULLRICH AND W. DUPPEL

at 39,000 g, adsorption of the supernatant on calcium phosphate gel with subsequent elution, and ammonium sulfate fractionation in the presence of 2 mM dithiothreitol (188). Tetrahydropterin is required as a cofactor, together with NADPH and the sheep liver dihydropterin reductase, as in the phenylalanine and tyrosine monooxygenase system. As in these systems the natural reductant can be replaced by dimethylpterin derivatives in the presence of sulfhydry1 groups. Kinetic experiments show that the tryptophan B-monooxygenase system and its mechanism closely resembles that of the phenylalanine system. The K,,, for substrate and oxygen varies with different pterin cofactors and Fez+ions are stimulatory as is catalase. These findings together with an inhibition by substrate a t higher concentrations in the presence of the natural cofactor tetrahydropterin indicate a possible common mechanism and similar enzyme systems for the monooxygenation of phenylalanine, tyrosine, and tryptophan in mammalian tissues. 5. Proline 4-Monooxygenase

Proline 4-monooxygenase is found in many tissues of active collagen synthesis as in chick embryo homogenates or in skin of newborn rats. It catalyzes the hydroxylation of proline in the 4 position, but only when it is incorporated in protocollagen or in synthetic copolymers with repeating units (X-Pro-Gly) (190,191). For reviews see (192-194). The enzyme has been purified extensively (190,195,196), and the best preparation has recently been obtained by affinity chromatography on agarose columns (191). It has been shown to be a tetramer with a molecular weight of 230,000, composed of two different types of subunits, which are maintained in the active conformation by disulfide bonds. This preparation showed no lysine and proline 3-monooxygenase activity, usually associated with other preparations. Electron micrographs have been taken from this pure preparation showing tetramers in which two V-shaped dimers are interlocked (197). Antibodies have been prepared, and with a specific staining for peroxidase labeled y-globulins it has been shown 190. K. I. Kivirikko and D. J. Prockop, ABB 118, 611 (1967). 191. R. A. Berg and D. J. Prockop, JBC 248, 1175 (1973). 192. M. E. Grant and D. J. Prockop, N . Engl. J . M e d . 286, 194 (1972). 193. J. M. Manning and A. Meister, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 27. Maruzen, Tokyo, 1966. 194. D. Fujimoto and N. Tamiya, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 58. Maruzen, Tokyo, 1966. 195. R. E. Rhoads and S. Udenfriend, ABB 139, 329 (1970). 196. J. Halme, K. I. Kivirikko, and K. Simons, BBA 198, 460 (1970). 197. B. R. Olsen, R. A. Berg, K. I. Kivirikko, and D. J. Prockop, Eur. J. Biochem. 35, 135 (1972).

5.

293

IRON- AND COPPER-CONTAINING MONOOXYGENASES

by light microscopy that the enzyme was distributed in granular structures in the cytoplasma of the cells (198). The isoelectric point was determined to be a t pH 4.4, indicating that the enzyme is a very acidic protein (199). For full activity of the purified monooxygenase Fez+,a-ketoglutarate and ascorbate are required (195). A reaction mechanism involving the decarboxylation of a-ketoglutarate has been proposed (600). COO@

+ (Y

-Ketoglutarate

(8) 0kf)j

‘0, C 50

€I0

I 2

HO C -~O Q *’,

I

7%

7H2 y

-CO,

Qcow-R

COO-

FH2

COR’ R

coo-

According to this mechanism this enzyme is not a typical monooxygenase since the second oxygen atom is not reduced to water but incorporated into the succinate molecule (201).

6. Lysine 5-Monooxygenase Besides proline 4-monooxygenation lysine monooxygenation has always been found during the formation of collagen from protocollagen, and it has been assumed that one enzyme catalyzes both reactions (20.9). This, however, was disproved by the discovery of a separate lysine 5-monooxygenase (,203).A partially purified preparation has been obtained from chick embryos after homogenization and ammonium sulfate precipitation, adsorption on calcium phosphate gel, two chromatographic elutions from DEAE-cellulose with different ionic strength, and subsequent gel filtration on Bio-Gel A-1.5 m (204). This treatment resulted in a 600-fold 198. R. A . Berg. B. R. Olsen, and D. J. Prockop, BBA 285, 167 (1972). 199. M. Pankalainm, H. Aro, K. Simons, and K. I. Kivirikko, BBA 221, 559 (1970). 200. R. E. Rhoads and S. Udenfriend, Proc. Nat. h a d . Sci. U . S. 60, 1473 (1968). 201. B. Lindblad, G. Lindstedt, M. Tofft, and S. Lindstedt, JACS 91, 4604 (1969). 202. K. I. Kivirikko and D. J. Prockop, Proc. Nut. Acnd. Sci. U . S. 57, 782 (1967). 203. R. L. Miller, ABB 147, 339 (1971). 204. K. I. Kivirikko and D. J. Prockop, BBA 258, 366 (1972).

294

V. ULLRICH AND W. DUPPEL

purification. Two forms of the enzyme have been obtained with apparent molecular weights of about 550,000 and 200,000. The substrate of the enzyme is either protocollagen or a synthetic peptide with the structure (X-Lys-Gly), (205). The present data indicate that the peptide chain length and the amino acid sequence in the vicinity of lysine are critical factors for substrate specificity. The cofactors required are the same as fof the proline monooxygenase and the K , values are 1 X M, M for Fez+,a-ketoglutarate, and ascorbate, reM , and 5 X 5X spectively. The pH optimum is a t 7.4, and the activity is increased by the addition of bovine serum albumin, catalase, and dithiotreitol. These results closely correlate with those of the proline 4-monooxygenase and although two separate enzymes have been isolated it can be assumed that the reaction mechanism, which is not defined exactly in either case, is very similar in both enzyme systems.

IV. Copper-Containing Monooxygenases

A. DOPAMINE p-MONOOXYGENASE Dopamine /?-monooxygenase catalyzes the hydroxylation of 3,4-dihydroxyphenylethylamine (dopamine) to norepinephrine, which is the last step in its biosynthesis. Other compounds like phenylethylamine, tyramine, and epinine can also serve as substrates (206'). Studies on the stoichiometry showed that 1 mole of O2 and 1 mole of ascorbate are used for each mole of norepinephrine formed, according to the reaction (207) : Dopamine

+ ascorbate + O2+ norepinephrine + dehydroascorbate + HzO

(9)

Fumarate is also required for maximum activity, but the requirement is not very specific. Ascorbate can be partially substituted for by catechol (208). Reduced pyridine nucleotides have no effect on the reaction. The enzyme has been purified from bovine adrenal medulla. It was solubilized with Cutscum (isooctylphenoxypolyethoxyethanol)and further purified by ammonium sulfate precipitation and chromatography on DEAE-cellulose and Sephadex G-200 (208). The purified enzyme showed only one protein band on disc electrophoresis. The molecular 205. K. I. Kivirikko, K. Shudo, S. Sakakibara, and D. J. Prockop, Biochemistry

11, 1n (1972). 206. S. Friedman and S. Kaufman, JBC 240, PC562 (1966). 207. E. Y. Levin, B. Levenberg, and S. Kaufman, JBC 235, 2080 (1960). 208. 5. Friedman and S. Kaufman, JBC 240,4763 (1965).

5.

295

IRON- AND COPPER-CONTAINING MONOOXYGENASES

weight was shown to be 290,000, consisting of four identical subunits. Two pairs are linked by disulfide bridges, and these dimers are held together by noncovalent bonds (209). The activation of the enzyme by dicarboxylic acids or salts is not mediated by a major change in the protein conformation, but rather by nonspecific anion activation, which occurs near the active center. Kinetic studies showed that fumarate facilitates the interaction of the reduced enzyme with oxygen and changes the K, for the substrate (210). Copper has been shown to be the effective metal in dopamine p-monooxygenase (206,208) and to undergo oxidation and reduction during the reaction. One monomer of the enzyme contains two copper ions. Electron paramagnetic resonance data showed that the copper is reduced by ascorbate, then reacts with oxygen, and the postulated active oxygen intermediate reacts with the substrate to yield the Cuz+protein, hydroxylated substrate and water, according to the equation (211): , cuz+ E,

ascorbate

\

*

cu=+

Norepinephrine +%O

/-Kine

CU2+

/ E

,cu+

,E k u +

(10)

\

,or

‘cuz+

The oxygen in the hydroxylated product has been shown t o be derived from molecular oxygen (212). A part of the copper ions is present in the cuprous state ; further reduction occurs with ascorbate, but addition of substrate causes reoxidation (208). The reaction is inhibited by 6X M cyanide as well as by carbon monoxide, both of which form complexes with Cul+, showing that the reduced copper ion plays an essential role in the reaction (208). By means of stereospecifically tritiated substrate it has been shown that the hydroxylation reaction proceeds with overall retention of configuration (21.3). The properties of copper in biological systems have been reviewed by Malkin and Malmstrom (5’14) and Peisach et al. (215). 209. J. E. Craine, G. H. Daniels, and S. Kaufman, JBC 248, 7838 (1973). 210. M. Goldstein, T. H. Joh, and T. Q. Gravey, 111, Biochemistry 7 , 2724 (1968). 211. S. Friedman and S. Kaufman, JBC 241, 2256 (1966). 212. S. Kaufman, W. F. Bridgers, F. Eisenberg, and S. Friedman, BBRC 9, 497 (1962). 213. K. B. Taylor, JBC 249, 454 (1974). 214. R. Malkin and B. G. Malmstrom, Advan. Enzymol. 33, 177 (1970). 215. J. Peisach, P. Aisen, and W. E. Blumberg, eds., “Biochemistry of Copper.’’ Academic Press, New York, 1966.

296

V. ULLRICH AND W. DUPPEL

B. PHENOL 0-MONOOXYGENASE Phenol o-monooxygenase is the name of an enzyme system that hydroxylates monophenols to orthodiphenols which are further oxidized to the corresponding o-quinones.

=&

OH

+

H,O

(11)

0,

R

R

Other names for this system are phenolase, tyrosinase, o-diphenol: 0, oxidoreductase, phenol-oxidase, or when the two activities are separated, cresolase and catecholase. Many different types of phenol monooxygenases are found in bacteria, plants, insects, marine animals, and mammals, where they participate in the biosynthesis of lignins, flavonoids, tannins, adrenaline, and melanine. For reviews see (216-224). It has long been known that phenolases are copper enzymes, and Mason et al. (2) could show by means of lRO-labeledoxygen that they belong to the class of monooxygenases. Several preparation methods have been worked out for the different phenol monooxygenases. Often acetone powders are prepared which are then further purified by chromatography (226-229). Multiple forms of the mushroom (226,227,230) and mammalian melanoma ( 2 S 1 ) phenol 216. C. R. Dawson and W. B. Tarpley, “The Enzymes,” 1st ed., Vol. 2, p. 454, 1951. 217. H. S.Mason, Advan. Enzymol. 16, 105 (1955). 218. A. B. Lerner, Advan. Enzymol. 14,73 (1953). 219. I. W. Sizer, Advan. Enzymol. 14, 129 (1953). 220. J. M. Nelson and C. R. Dawson, Advan. Enzymol. 4,99 (1944). 221. A. Mehler, in “Oxygenases” (0. Hayaishi, ed.), p. 87. Academic Press, New York, 1962. 222. D. Kertesa, in “Oxygenases” (0. Hayaishi, ed.), p. 307. Academic Press, New York, 1962. 223. H. S. Mason, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 287. Maruzen, Tokyo, 1966. 224. R. Zit0 and D. Kertesa, in “Biological and Chemical Aspects of Oxygenases” (K. Bloch and 0. Hayaishi, eds.), p. 290. Maruaen, Tokyo, 1966. 225. S. Bouchilloux, P. McMahill, and H. S. Mason, JBC 238, 1699 (1963). 226. R. L. Jolley, D. A. Robb, and H. S. Mason, JBC 244, 1593 (1969). 227. R. L. Jolley, R. M. Nelson, and D. A. Robb, JBC 244, 3251 (1969). 228. S.S. Patil and M. Zucker, JBC 240, 3938 (1965). 229. S.H. Pomeranta, JBC 238, 2351 (1963). 230. D. Kertesz and R. Zito, BBA 96, 447 (1965). 231. J. D. Burnett, JBC 246, 3079 (1971).

5.

IRON- AND COPPER-CONTAINING MONOOXYGENASES

297

monooxygenase have been described. The molecular weight of the mushroom protein is about 124,000-128,000 (227,230) and four isozymes have been identified, each of which contains one copper ion. After purification the cupric copper signal can be detected by EPR, but Cul+is also present in these preparations (225,230). Amino acid determinations of mouse phenol monooxygenase showed high content of hydrophobic residues, and the a-helical portion of the enzyme was determined to be about 2574 by C D and ORD measurements (231). The first phenol monooxygenase of a prokaryotic strain of Streptornyces glaucescens has been purified by Lerch and Ettlinger (232). Per molecular weight of 29,100 the enzyme contained one atom of copper in the cuprous form. No isozymes have been found in contrast to most eukaryotic organisms. The enzyme is very thermolabile compared to other fungal phenolases and has a half-life of only 52 sec a t 60°. Studies with KCN, a noncompetitive inhibitor, and 4-nitrocatechol, a competitive inhibitor, showing the same inhibitor effects on the monooxygenation of tyrosine and the oxidation of DOPA, indicate a tight coupling of these two reaction steps. On the other hand, the oxidation of the o-diphenol is more strongly dependent on .the oxygen concentration than the monooxygenation step. Two phenol monooxygenase isozymes of the eubacterium, Pibrio tyrosinaticus, with molecular weights of 41,000 and 38,500 have been purified (233).They do not cross-react with antiserum against hamster melanoma tyrosinase. The apoenzymes have been prepared by treatment with diethyldithiocarbamate and the activity could be fully restored by addition of Cu2+. The mechanism of the reaction is believed to be similar to the dopamine p-monooxygenation where oxygen reacts with two Cu+to an activated oxygenated complex which yields the Cu2+ enzyme, the diphenol, and water. The unique property of this enzyme is that the product of the first step (monooxygenation) , o-diphenol, serves as the electron donor for the reduction of the cupric ions under formation of the corresponding o-quinone.

232. K. Lerch and L. Ettlinger, Eur. J. Biochem. 31,427 (1972). 233. S. H. Pomerantz and V. V. Murthy, ABB 160,73 (1974).

This Page Intentionally Left Blank

Mobbdenum Iron-Sulfur

Fluvin Hydroxylases and Related Enzymes R . C. BRAY I . General Introduction .

. . . A . Enzymes to Be Considered .

. . . . . . . . . . 300 . . . . . . . . . . 300 B. Distribution and Biological Importance of the Enzymes . . . 301

I1. Milk Xanthine Oxidase . A . Introduction . . . . B . Molecular Properties . C . Catalytic Properties

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

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

I11. Other Molybdenum Hydroxylases

. .

. . . . . . . . . .

A Introduction . . . . . . . . . . . . . . B Molecular Properties . . . . . . . . . . . C . Catalytic Properties . . . . . . . . . . . I V . Genetic Studies and the Molybdenum Hydroxylases . . . A Introduction . . . . . . . . . . . . . . B . Xanthinuria and Gout in Man . . . . . . . . C The “Common Cofactor” of Nitrate Reductase and the Molybdenum Hydroxylases . . . . . . . . . D . Molybdenum Hydroxylases in Drosophile ~ e l a ~ o g a s ~ e.r E . Molybdenum Hydroxylases in Aspergillus nidulans . . V . Sulfite Oxidase of Liver . . . . . . . . . . . A Introduction . . . . . . . . . . . . . . B. Molecular Properties . . . . . . . . . . . C . Catalytic Properties . . . . . . . . . . .

. .

.

299

. . . .

. . . . . . . .

303 303 304 344 388 388 389 394 400 400 400

. . 402 .

. 406

. . 412

. . 414

. . 414 . . 414

. .

417

300

R. C. BRAY

I. General Introduction

A. ENZYMES TO BE CONSIDERED This chapter is concerned with the enzymes, xanthine oxidase (EC 1.2.3.2), xanthine dehydrogenase (EC 1.2.1.37), aldehyde oxidase (EC 1.2.3.1), and sulfite oxidase (EC 1.8.3.1). The first three of these form a very closely related group. Thus, their wide specificities (discussed in detail in Section II,C,l,a) overlap to a large extent and all of them, from all sources so far as is known, contain FAD, together with molybdenum and iron-sulfur centers. The reactions they catalyze are generally of the form: RH-

-2e

-H+

+OH-

ROH

where R H is the reducing substrate. The oxygen introduced into R H is derived from water, while the electrons from it_are accepted by the oxidizing substrate. The enzymes operate by undergoing a cycle of reduction by the reducing substrate then reoxidation by the oxidizing sustrate. Massey ( 1 ) referred to these enzymes as the “iron-sulfur flavoprotein hydroxylases.” However, from a mechanistic point of view, the presence of molybdenum is more noteworthy than is that of iron or flavin, and it is therefore proposed that they be called the “molybdenum hydroxylases.” For some of the molybdenum hydroxylases, oxygen is an efficient oxidizing substrate, whereas for others, it is not. However, in later sections it will become apparent that reactivity toward oxygen should probably be regarded as a relatively trivial property among these enzymes. Thus, there have been reports that specific chemical modification may lead to interconversions between “oxidase” and “dehydrogenase” forms of a given enzyme and indications, furthermore, that the dehydrogenase forms may be biologically the more significant. For convenience, the traditional names of the various enzymes will be used, e.g., rabbit liver aldehyde oxidase, Drosophila xanthine dehydrogenase, and milk xanthine oxidase. However, it must be emphasized that this nomenclature is really rather unsatisfactory, both with regard to the designations “oxidase” and “dehydrogenase” and in relation to the implied specificity to the reducing substrates. In addition to the molybdenum hydroxylases, sulfite oxidase is discussed in this chapter, though it contains heme rather than flavin and has no iron-sulfur. This is because i t contains molybdenum, apparently 1. V. Massey, in “Iron-Sulfur Proteins” (W. Lovenberg, ed.), Vol. 1, p. 301. Aca-

demic Press, New York, 1973.

6.

MOLYBDENUM HYDROXYLASES

301

in a chemical environment within the active center, which is quite like that of the metal in the other enzymes. A further somewhat related enzyme was considered for inclusion but was finally omitted. This was dihydro-orotate dehydrogenase (EC 1.3.3.1), an enzyme which contains flavin and iron-sulfur centers but has no molybdenum or heme. It was not included, mainly, since little work seems to have been done on it after the appearance of earlier reviews (2,3,Sa). Xanthine oxidase from cow’s milk has been studied far more extensively than have any of the other enzymes with which we are concerned. Therefore the chapter will of necessity be primarily devoted to it. Unless otherwise stated, the term “xanthine oxidase” is to be taken to refer to the enzyme from milk. Occasionally, in the sections on this enzyme, comparison with the properties of other molybdenum hydroxylases will be appropriate. However, a separate section specifically on these related enzymes is included (Section 111).Xanthine oxidase and related enzymes have been extensively reviewed (e.g., 1-7). Of these reviews, Bray ( 4 ) and D e Renzo (7) covered earlier work fairly fully, while Bray ( 5 ) , Bray and Swann (6),and Massey ( 1 ) remained reasonably up to date a t the time the present chapter was prepared. B. DISTRIBUTION AND BIOLOGICAL IMPORTANCE OF THE ENZYMES The enzymes to be considered are widely distributed. Thus, one or more molybdenum hydroxylases have been found in organisms as different in complexity as man and bacteria ( 4 ) . There has been less work on sulfite oxidase but i t appears (8) that this is as widely distributed, or even more so, than are the xanthine-oxidizing enzymes. It seems clear that enzymes metabolizing xanthine must have differing roles in different species. I n some lower organisms xanthine can serve as sole nitrogen source and when this is the case, conversion of xanthine 2. G. Palmer and H. Brintzinger, in “Electron and Coupled Energy Transfer in

Biological Systems” (T. E. King and M. Klingenberg, eds.), Vol. 1, Part B, p. 379. Dekker, New York, 1972. 3. K . V. Rajagopalan and P. Handler, in “Biological Oxidations” (T. P. Singer, ed.), p. 301. Wiley (Interscience), New York, 1968. 3a. T. P. Singer, M. Gutman, and V. Massey, in “IronSuIfur Proteins” (W. Lovenberg, ed.), Vol. 1, p. 225. Academic Press, New York. 4. R. C. Bray, in “The Enzymes,” 2nd ed., Vol. 7, p. 533, 1963. 5. R. C. Bray, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 216. Climax Molybdenum Co., London, 1973. The same paper is also reproduced in J . Less-Common Metals 36, 413 (1974). 6. R. C. Bray and J. C. Swann, Struct. Bonding (Bedin) 11, 107 (1972). 7. E. C. De Renzo, Aduan. Enzymol. 17, 293 (1956). 8. D. L. Kessler and K. V. Rajagopalan, JBC 247, 6566 (1972).

302

R. C. BRAY

to uric acid represents the first step of nitrogen utilization (see, e.g., 9). I n higher organisms the role of the enzymes becomes more obscure. It has widely been assumed for all molybdenum hydroxylases that the natural reducing substrates are normally purines or other heterocyclic compounds rather than aldehydes (10). The question of the natural oxidizing substrates wiIl be considered in Section II,B,3,d. In mammals oxidation of hypoxanthine to xanthine and of xanthine to uric acid are steps in purine catabolism prior to excretion, uric acid being the end product in primates. How important such reactions are to the organism is difficult to ascertain with certainty. On the one hand, individual humans, genetically deficient in xanthine oxidase, are apparently little the worse for the deficiency (11). Furthermore, inhibition of human xanthine oxidase activity by administration of the highly potent and specific inhibitor, allopurinol [ 4-hydroxypyrazolo (3,4-d)-pyrimidine], is beneficial in gout and in other conditions which result in overproduction of uric acid but is relatively free from side effects (see, e.g., 12), Similarly, young chicks treated with allopurinol grow normally despite inhibition of xanthine dehydrogenase activity resulting in hypoxanthine and xanthine replacing uric acid as the major end product of their nitrogen metabolism ( 1 3 ) .These observations might imply that oxidation of xanthine is superfluous in higher organisms. On the other hand, allopurinol administration to mice has been reported to decrease incorporation of hypoxanthine into nucleic acids (14) ; thus, the enzyme is not entirely without effect on important anabolic processes. However, there seems to have been very little foundation indeed for earlier ideas (cf. 4 ) of an association between low xanthine oxidase activities and cancer (16).

A novel biological role for xanthine oxidase has been put forward by Fried and co-workers (16). They suggested that xanthine oxidase exists primarily to produce hydrogen peroxide and superoxide free radicals from 9. A. J. Darlington and C. Scazzocchio, BBA 166, 569 (1968). 10. T. A. Krenitsky, S. M. Niel, G. B. Elion, and G. H. Hitchings, ABB 150, 585 (1972). 11. R.W. E. Watts, K. Engelman, J. R. Klinenberg, J. Seegmiller, and A . Sjoerdsma, Nature (London) 201, 395 (1964). 12. T. F. Yii and A. B. Gutman, Amer. J . M e d . 37, 885 (1964). 13. E. Weir and J. R. Fisher, BBA 222, 556 (1970). 14. R.Pomalea, S. Bieber, R. Friedman, and G. H. Hitchings, BBA 72, 119 (1963). 15. Milk xanthine oxidase was reported to have antitumor properties by A. Haddow, G. de Lamirande, F. Bergel, R. C. Bray, and D. A. Gilbert [Nature (London) 182, 1144 (1958)l. However, an unpublished reevaluation of the original data by R. C. Bray [quoted in Bray and Swam (611 showed that the result was not statistically significant. 16. R. Fried, L. W. Fried, and D. R. Babin, Eur. J. Biochem. 33, 439 (1973).

6.

MOLYBDENUM HYDROXYLASES

303

oxygen and that oxidation of the reducing substrate is purely incidental to this. The reduced oxygen derivatives are then supposed t o be available for coupled oxidation reactions, e.g., in drug catabolism. The most obvious objection to this hypothesis is the conclusion, discussed in Section I11B,3,d, that the dehydrogenase form of the xanthine-oxidizing enzymes is more “native” than is the oxidase form. On the other hand, Krenitsky et al. (10) proposed an entirely different, and perhaps more plausible, role on the basis of an element of complementarity they observed in the substrate specificities of milk xanthine oxidase and liver aldehyde oxidase (see also Section II,C,l,a). They suggested enzymes of this type may have evolved to hydroxylate, and so protect the host organism against a wide variety of naturally occurring pyrimidines. However,. it must be conceded that the true role of the enzymes in humans and many other species remains somewhat problematical.

II. Milk Xanthine Oxidare

A. INTRODUCTION Work on milk xanthine oxidase dates back to the very earliest days of biochemistry (4,7). At different times the enzyme has proved an attractive object of study by workers interested in a surprisingly wide range of aspects within the general area of enzymology. Work has always been facilitated by the comparatively high stability of xanthine oxidase and by the comparative ease with which i t may be obtained from cow’s milk. The latter usually contains around 50 mg/liter of the enzyme, or even more. Much early work on xanthine oxidase centered around the general problems of enzyme kinetics and specificity [see Dixon and Webb (17) for original references]. It was one of the first flavoproteins t o be purified (in 1939, see 18J.9) and crystallized (in 1954, see d0,21). It was also the first mammalian enzyme found to contain molybdenum (in 1953, see d d ) , while iron was found in it in 1954 (23). From this time onward, 17. M. Dixon and E. C. Webb, “Enzymes.” 2nd ed., pp. 78 and 203. Longmans, Green, London, 1964. 18. H.S. Corran, J. G. Dewan, A. H. Gordon, and D. E. Green, BJ 33, 1694 (1939). 19. E.G. Ball, JBC 128, 51 (1939). 20. P. G. Avis, F. Bergel, R. C. Bray, and K. V. Shooter, Nature (London) 173, 1230.( 19541. 21. P.G. Avis, F. Bergel, and R. C. Bray, JCS p. 1100 (1955). 22. D. E.Green and H. Beinert, BBA 11, 599 (1953). 23. D. A. Richert and W. W. Westerfeld, JBC 209, 179 (1954).

304

R. C. BRAY

most mechanistic work on the enzyme has been aimed at explaining the respective roles of these metal centers and FAD in the various catalytic reactions. Xanthine oxidase has a turnover number which is low enough to permit pre-steady-state kinetic studies of the reaction mechanism. Thus, the enzyme, containing several redox-active centers and being stable, water-soluble, readily available, and readily studied, may be regarded as a unique model for investigations of biological electron transfer processes generally. Since all of the centers of the enzyme can exist in both paramagnetic and diamagnetic oxidation states, it is not surprising that magnetic methods have played a large part in many of the more recent studies. Indeed, xanthine oxidase may be said to have acted as a spur to the application of such methods in the biochemical field. It was one of the first enzymes to be studied by electron paramagnetic resonance spectroscopy (EPR) (24,SS). It was also one of the first purified iron-sulfur proteins to be studied by EPR (26'), by magnetic susceptibility measurements (26,27), and by Mossbauer spectroscopy (28). More specifically, the rapid freezing method for following the time-course of fast reactions by E P R (29-32) was originally evolved for work on xanthine oxidase. This method provided the initial evidence for the functional role of molybdenum in the catalytic processes of the enzyme ( 3 3 ) .Electron paramagnetic resonance, alone and in conjunction with rapid freezing, has continued to provide important information on the catalytic mechanism of xanthine oxidase. However, much recent work of importance has depended on the use of more standard chemical and biochemical methods.

B. MOLECULAR PROPERTIES 1. Purification

a. Alternative Methods and Homogeneity of the Product. Apart from earlier work ( 4 ) ,a number of comparatively simple methods of obtaining 24. R. C. Bray, B. G. Malmstrom, and T. Vanngird, BJ 71,24P (1959). 25. R. C. Bray, B. G. Malmstrom, and T. Vanngird, BJ 73, 193 (1959). 26. R. C. Bray, R. Pettenson, and A. Ehrenberg, BJ 81, 178 (1961). 27. A. Ehrenberg and R. C. Bray, ABB 109, 199 (1965). 28. C. E. Johnson, P. F. Knowles, and R. C. Bray, BJ 103,lOc (1967). 29. R. C. Bray, BJ 81, 189 (1961). 30. R. C. Bray and R. Pettersson, BJ 81, 194 (1961). 31. D. P. Ballou and G. A. Palmer, Anal. Chem. 46, 1248 (1974). 32. R, C. Bray, D. J. Lowe, C. CapeillBre-Blandin, and E. M. Fielden, Bwchem. SOC.Trans. 1, 1067 (1973). 33. R. C. Bray, BJ 81, 196 (1961).

6.

MOLYBDENUM HYDROXYLASES

305

highly purified xanthine oxidase in good yield have been reported in detail in recent years (34-36). It may be well to emphasize some of the differences among these methods. Handler and co-workers (34) started from cream and avoided treatment with proteolytic enzymes. Massey and co-workers (35) used buttermilk (37) from pasteurized cream and included a proteolytic digestion step [as originally introduced by Ball ( 1 9 ) ] to improve their yields. Bray and co-workers (36) used buttermilk, in this case from unpasteurized cream, and they, too, used the proteolytic step. They described alternative susequent procedures for isolating the enzyme and some of these included a selective denaturation step, employing 0.1 to 0.6 M sodium salicylate as denaturant (38).Though the methods of all three groups of workers yield xanthine oxidase preparations which are grossly similar to one another, there are nevertheless differences, particularly in activity and molybdenum content, which will be considered fully in subsequent sections. Thus, the effect of proteolysis in relation to oxidase-dehydrogenase conversions is considered in Section 11,BJ3,d,and its effect on the molecular weight in Section II,B,2,b. Denaturation of demolybdo xanthine oxidase by salicylate (and perhaps in pasteurization) is discussed in the next subsection. Xanthine oxidase was ‘obtained about 80% homogeneous in the ultracentrifuge in 1939 (18) and crystalline and approaching 100% apparent homogeneity by various criteria in 1956 (40). It is, then, not surprising to note that the later, simpler, purification methods (34-36) also yield enzyme of high apparent purity. Thus, the preparations of Massey and co-workers (35)were stated (in a footnote) to show a single band in polyacrylamide gel electrophoresis, while those of Bray and co-workers 34. C.A. Nelson and P. Handler, JBC 243,5368 (1968). 35. V. Massey, P.E. Brumby, H. Komai, and G. Palmer, JBC 244, 1682 (1969). 36. L. I. Hart, M. A. McGartoll, H. R. Chapman, and R. C. Bray, BJ 116, 851 (1970). 37. Buttermilk is the aqueous phase remaining from cream, after its separation from butter, The separation is normally achieved by cooling, agitating, and centrifuging a t low speed. 38. Several different actiov of salicylate on xanthine oxidase must be distinguished. A t high concentrations it is a selective denaturing agent, useful in the preparation of enzyme free from the demolybdo form (see Section II,B,l,b). At lower concentrations (e.g., 1 mM) it is frequently used as a stabilizer of xanthine oxidize preparations. It presumably does this by chelating CU*+,etc., as well as by virtue of being a competitive inhibitor (39). A third action of salicylate (no doubt related to its denaturing properties) is implied by indications that the reagent may be used in preparing deflavo xanthine oxidase (Section II,B,3,b). 39. F.Bergel and R. C. Bray, BJ 73, 182 (1958). 40. P. G.Avis, F. Bergel, R. C. Bray, D. W. F. James, and K. V. Shooter, JCS p. 1212 (1956).

306

R. C. BRAY

(36) gave only single symmetrical peaks in the ultracentrifuge, with those of Handler and co-workers (34) showing only relatively minor impurities is also a good measure of the purity by this method. The ratio EZ80/E460 of xanthine oxidase preparations (see Table 11).On this basis the samples of Avis et al. (21) and Hart et al. (36) are slightly purer than those of other workers, although the differences are fairly small. An impurity, which is frequently difficult to remove from xanthine oxidase preparations, shows in the ultracentrifuge as a faster sedimenting peak with S about 17 (18). Hart et al. (36) used recycling gel filtration on Sephadex G-200 to eliminate this material, while Bray et al. (41) showed that the impurity so separated was of low activity, consisting largely of partly polymerized apo-xanthine oxidase protein (42). For much work on the enzyme (e.g., by E P R ) it has not been necessary to u s e t h e most highly purified xanthine oxidase samples. Thus, in the author’s laboratory the final purification step of Hart et al. (36) is frequently omitted, making the task of preparing the enzyme in quantity a very simple one. Purification of xanthine oxidase by means of affinity chromatography (44) will be considered in the next subsection. Despite the extreme value of the information derived from affinity chromatography experiments (&), it seems that preparation of the enzyme on a large scale by this method cannot be easy. Later work on xanthine oxidase (46,46), from the laboratory where the affinity chromatography studies were initiated, employed enzyme prepared by conventional methods. Thus, the use of affinity chromatography cannot yet be regarded as routine in xanthine oxidase purification. b. Active and Inactive Forms: Demolybdo Xanthine Oxidase. I n 1952, 41. R. C. Bray, A. J. Chisholm, L. I Hart, L. S. Meirwether, and D. C. Watts, in L‘Flavinsand Flavoproteins’’ (E. C. Slater, ed.), p. 117. Elsevier, Amsterdam, 1966. 42. A typographical error in Andrews et al. (43) has caused some confusion. In

the legend to Fig. 3 of this reference the symbols for extinction and activity are interchanged. In fact, the high molecular weight fraction removed in gel filtration has only traces of xanthine oxidase activity, as is clear from the text of the paper and from later work (41). Nevertheless, the error has given rise to the quite unjustifiable idea (34) that the 17 S impurity is an enzymically active dimer of the normal xanthine oxidase molecule. [Note that the latter is itself a dimer (see Section II,B,2,c); thus, the 17 S material must in reality be a tetramer.1 43. P. Andrews, R. C. Bray, P. Edwards, and K. V . Shooter, BJ 93,627 (1964). 44. D. Edmondson, V. Massey, G. Palmer, L. M. Beacham, and G. B. Elion, JBC 247, 1597 (1972). 45. J . 9. Olson, D. P. Ballou, G. Palmer, and V . Massey, JBC 249,4350 (1974). 46. J. S. Olson, D. P. Ballou, G. Palmer, and V . Massey, JBC 249,4363 (1974).

6.

MOLYBDENUM HYDROXYLASES

307

Morell ( 4 7 ) observed that anaerobic bleaching of the visible absorption spectrum of xanthine oxidase takes place in two distinct phases and attributed this phenomenon to the presence of an inactivated form of the enzyme in his preparations. Many subsequent workers apparently were loathe to admit that their own preparations might be contaminated with inactive xanthine oxidase, or allowed themselves, in various ways, to be confused by the presence of this material (e.g., 94,95,48). However, final proof of the correctness of Morell’s hypothesis came in 1972 with the resolution of the active and inactive forms of the enzyme by affinity chromatography by Edmondson et al. ( 4 4 ) . These workers used an analog of the inhibitor, allopurinol, bound covalently to a gel column. Active enzyme only, in a reduced form, was bound and could be eluted after reoxidation, thereby yielding the most active xanthine oxidase preparations which have so far been reported (see next subsection and Table I for specific activity data). Prior to the work of Edmondson e t al. ( 4 4 ) , there had been gradual improvements in xanthine oxidase preparation methods, resulting in progressively lower proportions of the inactive enzyme. Thus, we have the unusual phenomenon (5) that during the period 1953-1972, the specific activity of the best, essentially “homogeneous,” xanthine oxidase samples in the literature, increased more than threefold, a plot of activity against the year of the observation showing a roughly linear rise of activity with time. The properties of xanthine oxidase prepared by affinity chromatography (discussed in subsequent sections) provide some assurance, however, that limiting activity has now been reached. In 1956, Avis et al. (49) studied the molybdenum content of their crystalline and other xanthine oxidase preparations and found this to be nonstoichiometric and somewhat variable. The activity of their samples was also variable but did not correlate fully with molybdenum content. This led them to the hypothesis [more explicitly stated in Bray et al. (@)I of not one, but two, inactive forms of xanthine oxidase, one containing, and the other deficient in, molybdenum. Hart et al. (56) referred to these forms respectively as “inactivated” and “demolybdo” xanthine oxidase. Here, the latter term will be retained, but the inactivated form will be referred to as “desulfo” xanthine oxidase. The reason for adopting this name will become apparent in Section II,B13,e, where the nature of this form and its production during cyanide treatment of the enzyme, via loss of an essential sulfur atom, will be considered. As will be discussed in that section, desulfo xanthine oxidase is normally a preparation (or 47. D. B. Morell, BJ 51, 657 (1952). 48. M. A. McGartoll and R. C. Bray, BJ 114, 443 (1969). 49. P. G. Avis, F. Bergel, and R. C. Bray, JCS p. 1219 (1956).

308

R. C. BRAY

storage) artifact and conversion of active enzyme to it may be minimized by working throughout in the presence of salicylate and EDTA (cf. 58). Like other workers up to 1972, Hart et al. (36)had no means of eliminating desulfo xanthine oxidase from their purified preparations (60). The other inactive species, demolybdo xanthine oxidase, will be considered here. In retrospect, the first indication of its existence was the analysis originally reported for the enzyme of 1 Mo per 2 FAD ($2; also see 4 ) , as compared with the now accepted value of 1 : l (Section II,B,2,a). Hart et aZ. (36) succeeded in providing clear evidence from their analytical data for the existence of demolybdo xanthine oxidase, although they were unable to isolate it free from the other forms. They found that the amount of the demolybdo form, relative to the other forms, never changed in the course of conventional purification or handling of the enzyme. From extended studies on enzyme samples from the milk of individual cows i t was concluded that in contrast to desulfo xanthine oxidase, the demolybdo form is a natural product secreted along with the active enzyme and that the relative amounts are nutritionally rather than genetically determined. In some milk samples, there were slightly higher concentrations of the demolybdo form than of the normal forms. The demolybdo enzyme is more susceptible to denaturation (by salicylate) than is the active enzyme. This provided a means of eliminating unwanted demolybdo form from the preparations of Hart et al. (36'). When their enzyme was not so treated, it had a lower activity and a lower molybdenum content but was otherwise indistinguishable from that prepared by the denaturation method. Although affinity chromatography would be expected to separate both demolybdo and desulfo xanthine oxidase from the active enzyme, it seems, nevertheless, that in work in Massey's laboratory only the latter form was encountered and removed (44). The reason for the absence of the demolybdo form in their preparations (36) is not clear. One possibility would be that the nutritional status of the cows of Ann Arbor, Michigan in 1969, was superior to that of those in Reading, Berkshire in 1970. Another would be that the demolybdo form was initially present in the former but was eliminated by denaturation in the pasteurization of the cream which was employed (36).Whatever the reason, it seems clear that Massey and co-workers (36)were more fortunate than other workers (36,49) in having preparations containing no more than traces of the demolybdo form. Thus, prior to the advent of affinity chromatography, 50. Hart and Bray (61) succeeded in preparing partially purified xanthine oxidase apparently free from the desulfo form (see second line of Table I) by rapid

working, using high concentrations of the stabilizing agents, 51. L. I. Hart and R. C. Bray, BBA 146, 611 (1967).

6.

MOLYBDENUM HYDROXYLASES

309

they had to deal with problems caused by the presence of one inactive contaminant of the enzyme only, rather than two. Early work (49) and other unpublished experiments from the author’s laboratory have given no indications of the presence in demolybdo xanthine oxidase of any metal ions other than iron. Furthermore, attempts (unpublished) to increase activity in samples rich in the demolybdo form by incubation with molybdenum compounds have not proved successful. Demolybdo xanthine oxidase may, therefore, not simply have vacant metal binding sites but may also lack those groups which normally supply the ligands of the metal in the active enzyme (see also Section IV) . Many recent workers on milk xanthine oxidase do not appear to have given data adequate to decide whether or not their samples contained the demolybdo form. However, work by Johnson et al. (52) has a n important bearing on the origin of this species. They worked not on milk xanthine oxidase but on the enzyme in rat liver and other organs. They found that administration of tungsten to rats resulted in dramatic decreases in xanthine oxidase (as well as in sulfite oxidase) activity. They were unable to restore the activity of liver homogenates from such animals by addition of molybdate in vitro. However, in vivo restoration was readily achieved, and this could be observed even in the presence of inhibitors of protein synthesis. I n later work by the same group (53),the xanthine oxidase fraction from the livers of tungsten-treated rats was purified and shown t o contain the demolybdo enzyme. This cross-reacted fully, immunologically, with the normal enzyme and in contrast to preparations from milk (36) was obtained almost free from the normal molybdenumcontaining form. The demolybdo rat enzyme appeared (53) inactive toward xanthine as substrate (cf. 26) but active toward NADH (see Section II1C,2,a).It is interesting that, whereas inactive W-containing analogs of nitrate reductase (54, 55) and also apparently of sulfite oxidase (55a) are obtainable on tungsten administration, only demolybdo xanthine oxidase and not a W-analog is obtained ( 5 3 ) . C. Specific Activity. The relative ease with which xanthine oxidase may be largely separated from other proteins, coupled with the very substantial difficulties which are involved in separating active and inactive forms 52. J. L. Johnson, K. V. Rajagopalan, and H. J. Cohen, JRC 249,859 (1974). 53. J. L. Johnson, W. R. Waud, H. J. Cohen, and K. V. Rajagopalan JBC 249, 5056 (1974). 54. A . Notton and E. J. Hewitt, BBRC 44, 702 (1971). 55. K.-Y. Lee, R. Erickson, S.-S. Pan, G. Jones, F. May, and A. Nason, JBC 249, 3953 (1974). 55a. J. L. Johnson, H. J. Cohen, and K. V. Rajagopalan, JBC 249, 5016 (1974).

310

B. C. BRAY

of the enzyme and the considerable variability in the amounts of the latter, has given rise to special ways of expressing the specific activities of xanthine oxidase samples. Clearly, if impurities in a xanthine oxidase preparation include both extraneous proteins and inactive forms of the enzyme, then the latter are less likely to be inert in all types of experiment than are the former. Hence, knowledge of the relative concentration of active and inactive forms is likely to be more useful to an experimenter than is a conventional specific activity measurement expressed relative to total protein. To this end, Avis et al. (Id) introduced the ratio: Activity/E4601which they abbreviated to “AFR,” standing for activity-flavin ratio. This ratio is very readily determined and has been used with only very slight variations by many subsequent workers on the enzyme ( 5 6 ) . Some of the more recent specific activity data on xanthine oxidase are summarized in Table I (36,36,44,68).I n later sections, the rationale of the various methods of extrapolating activities to the theoretical limit for enzyme free from inactive forms, of Activity/Easo ( 2 3 . 5 O ) = 190, will become apparent. The percentage of functional enzyme in a sample is given by Activity/E,,, divided by 1.9. Table I shows that only samples prepared by affinity chromatography (4)come near the theoretical limit. At the time the previous edition of “The Enzymes” was prepared ( 4 ) , even the best xanthine oxidase samples in the literature contained some 50% of inactive forms ( 6 ) . 2. Composition and Physical Properties

a. Analysis. It seems generally agreed that milk xanthine oxidase, free from the demolybdo form, contains FAD, molybdenum, iron, and acidlabile sulfur in the ratio 1:1:4 :4. Detailed analytical data for all of these constituents are given by Massey et al. (35) and for the first three constituents only by Bray and co-workers (36,49). The amino acid analysis of xanthine oxidase was first reported by Bray and Malmstrom (69), and subsequent work has confirmed their results (34,36,41).The analysis shows no striking features other than a rather 56. The precise abbreviation used to denote this ratio in publications by the present

author has sometimes been dictated by the editorial whims of the journal in which the work was published. Thus, the Biochemical Journal did not like the original “AFR,” while in one case Bur. J . Biochem. (67) changed “Activity/Em1’ to the less meaningful “activity/absorbance ratio,” without telling the author. Here, we shall use “Activity/E,.” 57. J. C. Swam and R. C. Bray, Eur. J . Biochem. 26,407 (1972). 58. M. A. McGartoll, F. M. Pick, J. C. Swann, and R. C. Bray, BBA 212, 523 ( 1970)

6.

311

MOLYBDENUM HYDROXYLASES

TABLE I SPECIFICACTIVITYOF PURIFIED XANTHINEOXIDASE Sample Limiting extrapolated valuec Calculated! Prepared by affinity chromatography Prepared by salicylate denaturation Prepared conventiondly Prepared conventionaIly

Activity/El~,o (23.5')"

IU/mg (23.5°)b

Edmondson et al. (44)

188d

5.08

McGartoll et al. (68) Edmondson et al. (44) Hart et al. (36)

192~

5.18

183d

140

3.6

Hart et al. (36) Massey et al. (36)

92 143d

2.4 3.1d

Reference

-

Defined as AE:iY/min divided by E:,"," for the enzyme a t the dilution of the assay. Conditions are: xanthine, 0.1 m M ; pyrophosphate buffer p H 8.2, saturated with air (36). * Defined as micromoles of substrate transformed per minute per milligram of protein, under the conditions of footnote a. Various extrapolations were used, including those to zero slow phase reduction by xanthine (Section II,C,B,f), binding of one alloxanthine per FAD (Section II,C,l,g), and zero Slow molybdenum E P R signal (Section II,B,4,d). All data of Massey and co-workers have been reported a t 25", not 23.5" as originally specified (81). Correction has been made on the basis of a 1.12-fold change in activity between these temperatures (36). If the temperature coefficient of Bray el al. (86) were used, all Massey's values would be slightly lower than those indicated. Calculated from the corresponding a ~ t i v i t y / E ~on ~ othe basis of MW = 283,000 and t E t / 2 F A D = 72 (Table 11),with AeEY (xanthine) = 9.6 (49). Based on the highest activity/Mo ratio found for specially prepared, partially purified enzyme by Hart and Bray @I), divided by the extinction coefficient a t 450 nm for enzyme free from the demolybdo form. 0 The original value of 197 (68) becomes 192 when the €460 value of Table 11 i s employed.

low tryptophan content (59,60). Only total half-cystine has been reported, not thiol and disulfide contents. As summarized earlier ( 4 ) , reaction with mercurials and other reagents has indicated the presence of thiol groups. More recent work by Massey et al. ( 3 5 ) confirmed and amplified evidence for an essential thiol, which is available for fast reaction with mercurials only in the reduced enzyme. However, the role of this thiol in the catalytic reaction is quite uncertain ( 1 ) . Massey et al. 59. R. C. Bray and B. G. Malmstrom, BJ 93, 633 (1964). 60. E. Bayer, A. Bacher, P. Krauss, W. Voelter, G. Barth, E. Bunnenberg, and C. Djerassi, Eur. J. Biochem. 22, 580 (1971).

312

R. C. BRAY

(61) mentioned studies on the thiol content of the enzyme with 5 3 dithio-bis (2-nitrobenzoic acid) but gave no details. Finally, the relationship between thiol groups of the enzyme and the oxidase and dehydrogenase activities is discussed in Section II1B,3,d, while the “persulfide” group of Massey and co-workers (4$,68), which has been lost in the case of the desulfo enzyme, is considered in Section II1B,3,e. Clearly, further work on the thiol content of xanthine oxidase is called for. b. Physical Properties. Only properties of the oxidized (“resting”) enzyme will be considered in this section, while discussion of the reduced enzyme and magnetic properties of the enzyme are reserved for Section II,B,4. Measurements of a number of physical parameters for xanthine oxidase, carried out in different laboratories, are summarized in Table I1 (63). The five sets of molecular weight determinations by physical methods are in tolerable agreement and give an average result of 283,000 -C- 18,000. Nelson and Handler (34) concluded that the molecular weight of their preparations is significantly higher than that of the other workers and attributed this to the fact that they did not add proteolytic enzymes in the preparation. Table I1 shows that such a trend in molecular weight may indeed be indicated, although the difference is probably within the limits of error of the measurements, particularly since proteolyzed and nonproteolyzed enzymes were not compared side by side in the same laboratory. Further data, indicating that any change in molecular weight resulting from the proteolytic treatment must be small, come from agreement between the amino acid analyses of treated and nontreated enzyme (34,63a164). Agreement between the molecular weight of the enzyme from physical measurements and than from chemical determinations, based on FAD content and assuming 2 FAD per mole (Table 11), is good according to the data of Avis et al. (40,49) and Hart et al. (36), but less so from that of Massey et al. (35). Though protein dry weight determinations tend to be imprecise, the 20% discrepancy indicated in Table I1 would seem outside the expected limits of error. Perhaps the best interpretation is that the apparently higher molecular weight of Massey et al. (35) 61. V. Massey, H. Komai, G. Palmer, and G. B. Elion, JBC 245, 2837 (1970). 62. V. Massey and D. Edmondson, JBC 245,6595 (1970). 63. H. Komai, V. Massey, and G. Palmer, JBC 244, 1692 (1969).

63a. For more recent data on effects of proteolysis on the subunit composition, see W. R. Waud, F. 0. Brady, R. D. Wiley, and K. V. Rajagopalan, ABB (in press). 64. Apart from its effect on molecular weight, treatment of milk xanthine oxidase with proteolytic enzymes also modifies the behavior of the enzyme in chromatography on calcium phosphate (4,561. Similarly electrophoretic mobility and other properties of the enzyme from mouse tissues is changed on treatment with trypsin (66,66a,66).

0,

TABLE I1 MOLECUL.4R WEIGHT,EXTINCTION COEFFICIENTS, AND OTHER

Y PHYSICAL DAT.4 ON

MILK X A N T R I N E

(43)

Nelson and Handler. (34)

Massey et al. (35)

Hart et aZ.6 (36)

0 . 74c 11.4

0.737d 11.3

11.7

e

I

290,000

274,000

Avis et al.

Andrews et al.

Physical constant

(40,491

Partial specific volume (P) Sedimentation coefficient 0

s2o.lo

Molecular weight from s, D , and c Molecular weight from approach to equilibrium Molecular weight from gel filtration Molecular weight from FAD content, assuming 2 per mole 4$/2 FAD Em/Em E:Zprotein (280 nm)

265,000

300,000

286,000 362,0000p~

308,000

710 5.0-5.2 11.5

760 5.1-5.3

5.4 11.3

290,000'

OXIDASE

Average or best values

I

68-72 5.0-5.2 12.20-12.5'

No proteolytic treatment was used in the enzyme preparation of Nelson and Handler (34). Identical values were obtained for enzyme prepared with and without salicylate denaturation. Measured value. d Calculated from the amino acid analysis. 6 I n another paper (63), S Z ~ is . given ~ as 11.55 (concentration not stated) with a value for the deflavoenzyme of 11.2. f sio,w not determined; S Z ~ 11.0, , ~ a t 13 mg/ml, for enzyme prepared by salicylate denaturation. 0 Calculated from data in the reference given. A value of about 380,0000 based on amino acid analyses is quoted in Olson et al. (45),though details are not given.

a

283,000

72 5.0 11.7

3 3 3

314

R. C. BRAY

results a t least in part from contamination with the apoenzyme or its polymerization products (see Section II,B,l,a) since their E280/E450 ratio is relatively high (35, Table 11) and details of homogeneity tests are not given in their work. From the above discussion and averaging other data in Table I, it is concluded that pure xanthine oxidase, whatever the method of preparation, has a molecular weight of about 283,000, with the following extinc5.0; E:FFtein(280 nm), tion coefficients and absorption ratios : E280/E460, 11.7; and ey$/2 FAD = 72 (67).It may be added that, on full reduction of the enzyme, absorption a t 450 nm decreases by 70% (46). Earlier work on the isoelectric point of xanthine oxidase is summarized by Bray ( 4 ) . c. Dimeric Nature of the Enzyme (Including Other Molybdenum H y droxylases) . Work on the subunit structure of xanthine oxidase has generally been rather incomplete, Some ultracentrifuge studies were carried out by Andrews et al. (43) and by Nelson and Handler (34). It seemed difficult to break the enzyme down into units of less than half the original molecular weight. The sedimentation coefficient changed little over the pH range 3.5-9.8 ( 4 3 ) . Further, thiols plus urea (6.7 M ) reduced the sedimentation coefficient (cf. Table 11) only to about 8.3 ( 4 3 ) ,while even thiols plus guanidine (8 M ) did not bring the molecular weight (from approach to equilibrium studies) below 100,000 ( 3 4 ) . A stable unit of about half the molecular weight of the native enzyme therefore seemed to be indicated. Exposure to acidic conditions (pH about 1.9) also caused the enzyme to break down to units of about half the original size (34) or somewhat less (43), while alkaline conditions (pH 11.7 or above) caused incomplete breakdown to species with a variety of sizes ( 4 3 ) . Later work on the subunit structure of xanthine oxidase and of other molybdenum hyroxylases, by gel electrophoresis methods, is in general agreement with these conclusions. Thus, indications of subunit molecular weights of 150.000, 130,000-140,000. and 120,000 have been reported, respectively, for the enzymes from milk (67a,67b),Drosophila (67c,68),and 65. P. Joyce and E. J. Duke, BJ 125, lllP (1971). 65a. E. J. Duke, P. Joyce, and J. P. Ryan, BJ 131, 187 (1973). 66. T. J. Hayden, J. P. Ryan, and E. J. Duke, Biochem. SOC.Trans. 1, 247 (1973). 67. The molar extinction coefficient at 450 nm (per 2 moles of FAD) is particularly useful for calculating the concentration of the enzyme in partially purified xanthine oxidase samples. 67a. L. G. Nagler and L. S . Vartanyan, Biokhimiya 38, 561 (1973) ; Chem. Abstr. 79, 143851 (1973). 67b. L. G. Nagler and L. S. Vartanyan, Dokl. Akad. Nauk. SSSR 212, 1461 (1973) ; Chem. Abstr. 80, 105355 (1974). 67c. E. P. Candido, D. L. Baillie, and A. Chovnick, Genetics 77, s9 (1974). 68. W. D. Seybold, BBA 334,266 (1974).

6.

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315

chicken liver (6%) [see also work on aldehyde oxidase in Nelson and Handler ( 3 4 ) ] .There has been no indication, however, that the halfmolecule subunit is enzymically active (but see 68b). The physical and analytical data discussed in earlier subsections are entirely consistent with, but do not prove, that the xanthine oxidase molecule is a functional dimer, with two identical, independent active-site systems, each comprising 1 Mo, 1 FAD, and 4 Fe atoms, as was suggested for example by the present author ( 4 ) . Other possibilities also have to be considered, however. For instance, there might be two active center systems in the molecule, which were either not quite identical to one another or which had some sort of allosteric interaction between them. Alternatively, the whole molecule might be a single, very complicated, catalytic system, for example, with the two molybdenum atoms close together and forming a single unit reminiscent of the two iron-system of spinach ferredoxin. Although, briefly, the present author favored the idea of a single catalytic system in the xanthine oxidase molecule (48,69,70), he has now reverted to his earlier position (4-6‘). The most direct and convincing evidence for two equivalent active sites comes from inhibitor binding studies with alloxanthine. When appropriate corrections are made for the presence of inactivated enzyme, active xanthine oxidase binds precisely 2 moles of alloxanthine per mole, i.e., one inhibitor molecule per Mo atom (44,61,71).Binding is very strong and provides no indication that the second molecule of inhibitor might be bound less firmly than the first. Conversely, evidence for a single catalytic system in the molecule might come either from a demonstration of binuclear molybdenum in the enzyme (72) or from a demonstration that the two FAD molecules were not located equivalently to one another. In fact, evidence from EPR work (see also Section 1I,B14,d) is against binuclear molybdenum in xanthine oxidase, although this situation is of course very common in low molecular weight Mo compounds. Thus, although integration of the Mo (V) EPR signals has never been reported to exceed the equivalent of about or?e unpaired electron per two Mo atoms (673, this may readily be explained in terms of equilibria among paramagnetic and diamagnetic valence states 68a. R. Andres, H. G. Lebherz, and H. Usprung, M o l . Biol. R e p . 1, 81 (1973). 68b. P. E. Brumby and V. Massey, BJ 89, 46P (1963). 69. R. C. Bray, P. F. Knowles, and F. M. Pick, FEBS Sgmp. 16, 267 (1969). 70. R. C. Bray, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 385. Univ. Park Press, Baltimore, Maryland, 1971. 71. Variation of the stoichiometry of alloxanthine binding with the proportion of active enzyme in xanthine oxidase samples was one of the methods used to extrapolate the limiting Activity/E,, ratio of the enzyme (61; see also Table I). 72. The term “binuclear” is used here in the inorganic chemist’s sense to denote 2 Mo atoms, close together and separated only, e.g., by bridging 0 or S atoms.

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of Mo. Furthermore, a single electron interacting with two 05M0nuclei would be expected to show an 11-line EPR spectrum and not the 6-line spectrum which is observed (7s). There have been several claims of nonequivalence of the two FAD molecules. The first, relating to milk xanthine oxidase, was made by the present author ( 4 8 ) . It soon became clear, however, that the nonequivalence was apparent rather than real and mainly resulted from the preparations used containing, by chance, about equal amounts of active and desulfo enzyme, with only FAD of the former responding to the test which was applied. The test for nonequivalence (48) involved alkylation of enzymebound FAD molecules in the reduced state by iodoacetamide (74) (see Section II,B,3,c). I n the system which was used, only active enzyme molecules were reduced and hence had reactive FAD (58,74). It was, however, originally erroneously assumed (48) that all the molecules in the enzyme preparation were identical, having one FAD reactive and the other unreactive. The other claims of nonequivalence of the FAD molecules relate to dissociation of flavin from molybdenum hydroxylases in the presence of 3 M KI, a process which is accelerated by prior reduction of the enzyme (see Section II,B,3,b). Kanda and Rajagopalan (75) worked with chicken liver xanthine dehydrogenase and Coughlan and Johnson (75a) with milk xanthine oxidase. Both groups obtained evidence for only partial dissociation of the flavin, from enzyme which had been prereduced with xanthine, and attributed this to nonequivalence of the two FAD molecules. However, later work by Cleere and Coughlan (75b) has provided strong evidence that these results should instead be explained in a manner analogous to those relating to iodoacetamide treatment. We therefore return to the proposition that xanthine oxidase has two independent active center systems per mole (75c),but there are two further points to be considered. The first concerns the presence of desulfo 73. R. C. Bray and L. 9. Meriwether, Nature (London) 212, 467 (1966). 74. H. Komai and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 399.Univ. Park Press, Baltimore, Maryland, 1971. 75. M. Kanda and K. V. Rajagopalan, JBC 247, 2177 (1972). 75a. M. P. Coughlan and D. B. Johnson, BBA 302, 200 (1973). 75b. W.F.Cleere, C. O’Regan, and M. P. Coughlan, BJ 143, 465 (1974). 75c. There has also been a suggestion (7bd) of nonequivalence of the active center systems (in turkey xanthine dehydrogenase) based on the finding of about 50% inhibition, developing relatively slowly, in the presence of sulfide. Perhaps, however, i t would be desirable for this work to be repeated, both in view of the difficulties of working with sulfide over long periods (except at high pH values) and also in view of the failure of other workers (681, to comment on inhibition of molybdenum hydroxylases by sulfide.

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(or demolybdo) centers in the preparations and the question of whether these occur two-to-a-molecule or sometimes as mixed species, with one half of the molecule in the desulfo, and the other half in the active form. If the two halves of the molecule are truly independent, then a random distribution of double-active, mixed, and double-desulfo molecules would be expected. Such a situation would tend to make effective affinity chromatography difficult and has indeed been put forward to explain low yields of fully active enzyme from such work (44). A final point concerns whether, in the double-active molecule, the two halves are exactly the same or only nearly the same. Although the author (76e) earlier concluded that detailed features of an E P R signal from Mo(V) in the reduced enzyme (the Rapid signal, see Section II,B,4,d) might result from slight differences in the environments of the metal atoms in the two halves of the enzyme molecule, nevertheless, other explanations of the experimental data are quite possible. We are therefore left with no firm evidence against complete equivalence of the two halves of the xanthine oxidase molecule. 3. Chemical Modification

a. General Comments. There seems to have been relatively little work aimed a t the chemical modification of amino acid residues in xanthine oxidase other than cysteine (cf. Sections II,B,2,a and II,B,3,d). However, Greenlee and Handler ( 7 6 ) , in connection with their proposal (77; Section II,C,l,a) that the active site contains a group with a pK of 10.7, studied reaction of the enzyme with some reagents for amino groups, including 2,4-dinitrofluorobenzene. They succeeded in obtaining, e.g., about 50% inhibition of enzymic activity, when some 7 NH, groups had apparently reacted with this compound. On the other hand, much effort has been expended in trying to remove Fe, Mo, and FAD, selectively, and/or reversibly, from xanthine oxidase. Only removal of the last of these has proved successful, and this will be discussed in the next subsection. Early claims that molybdenum could be removed reversibly from the enzyme rested on slender evidence ( 4 ) . Later, similar claims relating to iron (77a,77b) are also to be discounted (36). 7Bd. W. F. Cleere and M. P. Coughlan, BJ 143, 331 (1974). 75e. R. C. Bray and T. Vanngbrd. BJ 114, 725 (1969). 76. L. Greenlee and P. Handler, JBC 239, 1096 (1964). 77. L. Greenlee and P. Handler, JBC 239, 1090 (1961). 77a. E. Beyer and W. Voelter, BBA 113, 632 (1966). 77b. M. Uozumi, R. Hayashikawa, and L. H. Piette, ABB 119, 288 (1967). 77c. U . Branzoli and V. Massey, JBC 249, 4339 (1974).

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b. Deflavo Xanthine Oxidase (Including Other Molybdenum Hydroxylases). It was not until 1969 that FAD was successfully removed in a reversible manner from xanthine oxidase by Komai et al. ( 6 3 ) . Their procedure, based on early work by Morel1 ( 4 7 ) ,involved treatment with high concentrations of calcium chloride (e.g., with 2 M CaCl, a t pH 8.0 and 20° for 90 min). A product which was almost homogeneous in the ultracentrifuge (see Table 11) was obtained in 35% overall yield after dialysis and purification on Sephadex G-200.The deflavo enzyme contained no FAD but had undiminished contents of Mo, Fe, and acid-labile sulfur and exhibited the visible absorption spectrum shown in Fig. 1. The catalytic properties of deflavo xanthine oxidase are considered more fully in later sections; however, we note here the very important finding (63) that it is devoid of oxidase activity but retains dehydrogenase activity. Reconstitution of about 60% of the original oxidase activity was achieved 50,000

40,000

-0

2 30,000

0

0

?

L 2U 2 L

0 a Y

u,

20,000

Dif,ference

A

I0,OOC

Wavelength

(nm)

FIQ.1. Absorption spectra of native and deflavo milk xanthine oxidase, at pH 8.5. The lower curve shows the calculated difference spectrum between the two forms of the enzyme. Reproduced from Komai et al. (63).

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by incubation with FAD (for 5 min a t 2 5 O ) . Reconstitution of the activity could also be achieved with F M N in place of FAD but whereas very tight binding of FAD to the protein was indicated, binding of FMN was weaker, with K,,, about M . Stability of the deflavo enzyme is not discussed by Komai et al. ( 6 3 ) ,but presumably failure to achieve 100% reconstitution of oxidase activity indicates that stability is significantly less than that of native enzyme. Deflavo aldehyde oxidase has been prepared (77c) by a slight modification of the procedure used with xanthine oxidase. Alternative methods of flavin removal have been described by Kanda et al. (78), both for milk xanthine oxidase and for chicken liver xanthine dehydrogenase, and were later applied to xanthine dehydrogenase from turkey liver (76b). Kanda et al. (78) obtained indications of flavin removal on treatment of the enzymes with salts such as sodium salicylate (38; see also 36), KCNS, or KI. Their preferred procedure (which has already been mentioned in Section II,B,2,c) involved treatment of the reduced enzyme with KI. Conditions necessary for flavin removal were quite mild, e.g., for the milk enzyme, 3 M K I at p H 7.8 and 4 O for 30 min. With brief pre-reduction with xanthine, only active enzyme loses its flavin, whereas, if NADH is used for the reduction, then the desulfo form reacts also (76b; see also 7 6 ) .The original finding (63) that deflavo xanthine oxidase is devoid of oxidase activity was confirmed ( 7 8 ) .Stability of the product obtained by this method was said to be good, with 80% recovery of initial activity on adding FAD back again (see also 76a). For this reason, the KI-reduced enzyme method of flavin removal (78) may be superior to the original CaCl, method ( 6 3 ) . c. Enzyme with Alkylated FAD. Riboflavin derivatives are chemically quite reactive and may, e.g., be readily alkylated in positions 4a or 5 with profound effects on their properties (79-81). It is therefore not particularly surprising to learn that enzyme-bound FAD in xanthine oxidase undergoes similar reactions. Thus, the FAD of the reduced enzyme may be alkylated by treatment with iodoacetamide (74,82; see also Section 78. M. Kanda, F. 0. Brady, K. V. Rajagopalan, and P. Handler, JBC 247, 765 ( 1972). 79. P. Hemmerich, V. Massey, and G. Weber, Nature (London) 213, 728 (1987). 80. W. H. Walker and P. Hemmerich, Eur. J . Biochem. 13, 258 (1970). 81. M. Briistlein, W. R. Knappe, and P. Hemmerich, Angew. Chem., Int. Ed. Engl. 10, 804 (1971). 82. Alkylation of the flavin of xanthine oxidase by iodoacetamide was originally

misinterpreted as being alkylation of an essential thiol in the reduced enzyme (83), with concomitant liberation of FAD (&,70). Since alkylated, enzymebound FAD has no absorption at 450 mm (74) and since it remains nonfluorescent when liberated from the protein and hydrolyzed for FAD analysis (36) the flavin remained undetected and was assumed to have been lost (48s). 83. R. C. Bray and D. C. Watta, BJ 98,142 (1966).

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II,B,2,c). The enzymic properties of the product are very similar to those of the deflavo enzyme (Section II,B,3,b). It has no oxidase activity but retains dehydrogenase activity (70,74,84). Use of the alkylated enzyme (which is relativeIy stable) in mechanistic sfudies on xanthine oxidase might provide a useful supplement to work with deflavo enzyme, although little work on these lines seems to have been, done so far. Komai and Massey (74) succeeded in obtaining reversal of the alkylation process, with partial restoration of oxidase activity of the enzyme, by irradiation with visible light. Photodissociation of the alkyl group from alkylated flavins is a known reaction (79).They also found that reaction of reduced enzyme with iodoacetamide was not the only procedure available for alkylation of its flavin. A photoalkylation could be brought about by anaerobic irradiation of the enzyme in the presence of phenyl acetate, and some reversal of this benzylation reaction was achieved on irradiating aerobically (74). The benzylated enzyme was similar to the product from the iodoacetamide reaction. d. Thiol Groups and Proteolysis in Xanthine Oxiidase-Dehydrogenase Interconversions (Including Other Molybdenum Hydroxylases) . There has long been doubt as to whether the molybdenum hydroxylase of rat liver is an oxidase or a dehydrogenase (see, e.g., 86). I n 1968, Della Corte and Stirpe made the surprising observation (86) that storage of freshly prepared rat liver supernatants at -20° brought about dramatic changes in the acceptor specificity of the xanthine “oxidase” in them. Initially, with oxygen as oxidizing substrate, activity was quite low, but this increased some 4-fold or more on storing the supernatants a t -20° for about 6 hr. This change in activity was paralleled by a more than tenfold decrease in the rate of enzyme-catalyzed NAD’ reduction when using xanthine as reducing substrate. These observations suggested that the enzyme of rat liver was initially predominantly an NAD’ dehydrogenase, which was, only subsequently, converted by an unknown mechanism to an oxidase. The conversion process was investigated in a series of papers by Stirpe and co-workers. They finally concluded (87) that the two essential factors involved in the modification of the enzyme were thiol groups and proteolysis. More recently the rat liver enzyme has been further in84. There are discrepancies in the literature regarding the activity of various forms

of xanthine oxidase in the xanthine-ferricyanide assay. Relative to native enzyme, the deflavo enzyme has been reported to have either somewhat diminished (63) or somewhat enhanced (78) activity, whereas the alkylated enzyme has much diminished (74) activity in this system. (See also Section II,C,l,b.) 85. D. A. Richert, R. Vanderlinde, and W. W. Westerfeld, JBC 188, 261 (1950). 86. L. Della Corte and F. Stirpe, BJ 108, 349 (1968). 87. L. Della Corte and F. Stirpe, BJ 126, 739 (1972).

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vestigated by Waud and Rajagopalan ( 8 7 ~ ) These . workers confirmed many of Stirpe’s findings and succeeded in purifying the enzyme both as an NAD+-dependent dehydrogenase (D form) and as an oxidase (0 form). Work on related phenomena in the milk enzyme (88) has also been reported by Stirpe and co-workers. They isolated xanthine oxidase from fresh milk by conventional methods, without addition of proteolytic enzymes (89) but working throughout in the presence of mercaptoethanol. The product had enzymic properties not unlike those of the milk xanthine oxidase preparations of other workers (90). However, after incubating their purified enzyme for 20 min a t 370 with 10 mM dithioerythritol [and apparently not removing the reagent before activity measurements et al. (88) found that the enzyme had become were made ( ~ O U ) Battelli ], analogous to the freshly prepared rat liver enzyme in its properties, i.e., it was, in effect, now an NAD+ dehydrogenase. After treatment with the thiol, a 3- to 4-fold decrease in xanthine oxidase activity was reported, with a roughly 8-fold increase in xanthine-NAD+ reductase activity (measured aerobically). This D form of the enzyme could not be obtained when starting from enzyme prepared by purification methods involving the proteolysis step. It was reported, however, that it could be converted, irreversibly, back again to the 0 form by treatment with proteolytic enzymes, e.g., chymotrypsin gave a 2- to 3-fold increase in the oxidase activity, with NAD’ reductase activity decreasing almost to zero. It was tentatively suggested (88) that the D form of the enzyme contains a n essential thiol group (91),presumably in the vicinity of the flavin, which must remain intact for NAD+ reductase activity to be exhibited. Oxidation of the thiol (occurring spontaneously despite the presence of merceptoethanol), or modification of this group by a variety of thiol reagents (87), causes decreases in NAD+ reductase activity and increases 87a. W. R. Waud and K. V. Rajagopalan, in “Flavins and Flavoproteins” (T.P. Singer, ed.). ASP, Amsterdam (in press). 88. M. G. Battelli, E. Lorenzoni, and F. Stirpe, BJ 131,191 (1973). 89. For effects of proteolytic enzymes on xanthine oxidase, other than on the oxidase-dehydrogenase properties, see Sections II,B,l,a and II,B,Z,b. 90. The preparation of Battelli et a2. (88) had the rather low specific activity of 1.5 IU/mg (cf. Table I) and was not homogeneous by gel electrophoresis. However, further work (F. Stirpe, unpublished) indicates that enzyme purified to apparent homogeneity in the ultracentrifuge behaves similarly. Ma. I t may be significant in relation to these findings that thiols are capable of reducing some molybdenum hydroxylases to a significant extent (gob). 9Ob. M. J. Barber, R. C. Bray, D. J. Lowe, and M. P. Coughlan, BJ (in press). 91. The thiol involved in oxidase-dehydrogenase interconversions appears to be distinct from the thiol (discussed in Section II,B,2,a) which becomes reactive when the enzyme is reduced.

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in the normal oxidase activity. Proteolysis presumably moves the thiol away from the vicinity of the flavin. I n agreement with the hypothesis that the environment of the flavin is different in the 0 and D forms of the enzyme, preliminary E P R studies in the author’s laboratory (91~) have shown that the D form shows a much stronger FADH’ signal under a variety of conditions than does the 0 form, with only minor differences in signal intensity and no differences in signal form for the other E P R chromophores (cf. Section II,B,4). Many of these tentative conclusions have recently been put on a firmer footing by Waud and Rajagopalan (87~). Further work will be required on the thiol content of xanthine oxidase and on the effects of added thiols on the various assay systems before the oxidase-dehydrogenase conversions can be fully understood ( 9 2 ) . However, an analogy may be drawn with lipoyl dehydrogenase (see 93 for recent work). Here, it is well established that a thiol group, presumed to be in the vicinity of the flavin, has a profound influence on the catalytic properties of the enzyme. Modification of this thiol dramatically increases the activity at neutral pH with NADH as reducing substrate and indophenol as oxidizing substrate. With benzoquinone as oxidizing substrate, there is a shift in the reaction mechanism when the group is modified from a two-electron to a one-electron pathway. Work by Stirpe and co-workers on oxidase-dehydrogenase interconversions in the molybdenum hydroxylases from a variety of other sources is of interest in relation to the nature of the natural oxidizing substrate, especially in humans. It was concluded (94) that the native enzyme in all rat organs is probably of the D type, although conversion to the 0 type is often hard to prevent. Similarly, the human liver enzyme, too, appears to be an NAD+ dehydrogenase which may be converted to an oxidase ( 9 6 ) , while the avian enzyme seems to be permanently of the D type (87). e. Desulfo Xanthine Oxidase. This inactive form of the enzyme has 91a. R. C. Bray, M. J. Barber, M. P. Coughlan, H. Dalton, E. M. Fielden, and D. J. Lowe in “Flavins and Flavoproteins” (T. P. Singer, ed.). ASP, Amsterdam (in prem). 92. Work by M. S. Briley and R. Eisenthal [BJ 143, 149 (1974)l has shown that oxidase-dehydrogenase properties of xanthine oxidase are also modified, reversibly, when the enzyme is bound to milk fat globule membrane. The mechanism whereby binding influences the active sites of the enzyme is quite obscure. However, it is of interest that the membrane-bound enzyme is apparently more accemible to NADH (lower K,) than is the free enzyme. 93. M. Nakamura and I. Yamazaki, BBA 267, 249 (1972). 94. M. G. Battelli, L. Della Corte, and F. Stirpe, BJ 126, 747 (1972). 95. L. Della Corte, G . Gozzetti, F. Novello, and F. Stirpe, BBA 191, 164 (1969).

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already been mentioned in Section II,B,l,b. It was formerly referred to as xanthine oxidase-i, or as inactivated xanthine oxidase. As discussed above, it gradually accumulates during storage of xanthine oxidase samples, but transformation of the active enzyme into it is retarded by the presence of salicylate and EDTA. That desulfo xanthine oxidase is a preparation or storage artifact and not a natural product is indicated by data in Table I. The calculation by McGartoll et al. (58) shows that a specially prepared sample of Hart and Bray (51) (see second line of Table I ) , although it contained demolybdo xanthine oxidase, was nevertheless free from the desulfo form, as shown by the high Activity/Mo value. The nature of desulfo xanthine oxidase became apparent from work on the mechanism of inactivation of the enzyme by cyanide reported by Massey and Edmondson in 1970 (62). Following up earlier work on the cyanide reaction (96),they found unexpectedly, using [14C]cyanide, that radioactivity was not bound to the enzyme during the primary inactivation reaction. Instead, for each mole of active center destroyed, 1 mole of [ 14C]thiocyanate was liberated. They then attempted to reactivate the enzyme by restoring the liberated sulfur atom to it, and succeeded in doing this on incubation under specific conditions with sulfide ions (96~). Although only partial restoration of activity was achieved, [ 35S] sulfide could be incorporated into the cyanide-inactivated enzyme and could then be released again as [s5S]thiocyanate on repeating the cyanide treatment. It was proposed (62) that the active enzyme contains an essential persulfide group which reacts with cyanide as follows : Protein-S-S-

+ CN- -+protein-S- + CNS-

Possible involvement of the “persulfide” group in the catalytic reaction is discussed in Section II,C,2,f. Although in later sections, in referring to this group, quotation marks are not used, it must be emphasized that its identity has not yet been fully established. Replacement of coordinated oxygen in molybdenum complexes by sulfur on treatment with H,S has 96. M. P. Coughlan, K. V. Rajagopalan, and P. Handler, JBC 244, 2658 (1969). 96a. In the case of the enzyme from turkey liver reactivation not only by sulfide but also by selenide has been reported ( 7 6 d ) .However, effects of these reagents are complex in that they cause partial inactivation of the functional turkey enzyme as well as partial reactivation of the cyanide-treated form. (The mechanism of this inactivation by sulfide is not understood; the suggestion (76d that it depends on reaction of sulfide with Fe/S I1 seems to rest on slender evidence.) In the case of aldehyde oxidase, on the other hand, reactivation of cyanide-treated enzyme by sulfide could not be achieved (96b). 96b. U. Branzoli and V. Maasey, JBC 249, 4346 (1974). 96c. R. N. Jowitt and P. C. H. Mitchell, Chem. Commun. p. 605 (1966).

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been reported (96c; see also 96d). Furthermore, in the enzyme, the group’s reactions are not always those of a typical persulfide, e.g., it is not acid-labile or sensitive to reaction with a nucleophile such as hypotaurine. Another result not obviously explained by the persulfide hypothesis is the finding (62) that the fully reduced enzyme is not sensitive to cyanide. At the least, interaction of the persulfide with other groups in the enzyme, particularly the molybdenum (6,44,62), has to be postulated to explain these findings. Desulfo xanthine oxidase contains a full complement of Fe, Mo, FAD, and acid-labile sulfur and has a visible absorption spectrum differing only very slightly, though significantly, from that of the active enzyme ( 6 2 ) . Desulfo enzyme prepared by cyanide treatment appears to be essentially indistinguishable from that arising by “spontaneous” inactivation of the native enzyme (44). They have very similar difference spectra relative to the native enzyme and both can be partially reactivated with sulfide (97). Also, both show the same EPR properties (see Section II,B,4,d) and in particular give the characteristic Slow Mo (V) signal. As will be discussed further in later sections, desulfo xanthine oxidase is devoid of enzymic activity .except toward NADH as reducing substrate. f. Other T y p e s of Chemical Modification. A number of chemical modification reactions will be considered in this section. Some of these are dependent on the presence in the xanthine oxidase molecule of the persulfide sulfur discussed in the previous subsection. Reaction of xanthine oxidase with methanol and with formaldehyde has caused considerable interest. It has been known for some 30 years that both these reagents, under mild conditions, can cause loss of activity of xanthine oxidase preparations. However, i t was not until 1971 that Pick et al. (98) showed that the two inactivation processes are closely related. The product retains Fe, Mo, and FAD and has a characteristic difference spectrum relative to native enzyme (96,99).Furthermore, it also shows a highly characteristic Mo(V) EPR spectrum, the Inhibited 96d. A. Kay and P. C. H. Mitchell, JCS,A p. 2421 (1970). 97. There are indications that the desulfo enzyme prepared by cyanolysis is more fully reactivated by sulfide than in the spontaneously generated form (compare 6.3 and 44). However, there may be some doubt about this since the conditions for reactivation are critical (44). In any case, this does not detract from the proposal (44) that loss of the sulfur atom is the primary cause of spontaneous loss of activity by the enzyme since loss of sulfur might sometimes be followed by slower secondary changes rendering reactivation impossible. 98. F.M. Pick, M. A. McGartoll, and R. C. Bray, Eur. J . Bioehem. 18,65 (1971). 99. R. C. Bray, P. F. Knowles, and L. S. Meriwether, in “Magnetic Resonance in Biological Systems” (A. Ehrenberg, B. G. Malmstrom, and T. Vinnglrd, eds.), p. 249. Pergamon, Oxford, 1967.

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signal (see Section II,B,4,d). Study of this EPR signal has revealed a number of important points about the reaction (98). Loss of enzymic activity, which may be partly reversed under specific conditions, closely paralleled development of the signal (98). The Inhibited signal, unlike other Mo(V) signals from the enzyme, is stable in air. Thus, Mo(V) is somehow stabilized in the reaction product. Studies with the deuteriumsubstituted reagents, C2H,0H and 2HC*HO, showed that in the signalgiving species, a single nonexchangeable proton derived from the inactivating agent is coupled to the metal. It was proposed (98) that a formyl residue, C H O , from the reagent becomes bound to the active center in the vicinity of molybdenum during the inactivation. This still seems the most probable explanation, although the precise mechanism of the reaction (loo),particularly when methanol is employed ( I O l ) , remains somewhat unceEtain (see also Section II,C,2,f). Not unexpectedly, it turns out (44) that the persulfide group of the enzyme has to be intact for the reaction with formaldehyde to occur. In agreement with this, it was earlier reported by Coughlan et al. (96) that the reactions of xanthine oxidase with methanol and with cyanide are mutually exclusive. However, although suggestions have been put forward (51, the precise involvement of the persulfide group in the inactivation process is not certain. Inactivation of xanthine oxidase by arsenite has long been known (103-106), and the reaction has features in common with both the cyanide and the formaldehyde-methanol reactions (96). The product again has a characteristic difference spectrum from native enzyme, but in this case there seem to be no indications of a characteristic Mo(V) EPR spectrum (96). Arsenite inactivation is also more readily reversed than is either of the other inactivation processes. (This is particularly so for aldehyde oxidase where arsenite is merely a competitive inhibitor.) Whereas cyanide treatment prevents subsequent reaction (followed spec100. Since H202 must be present during inactivation of the enzyme by formaldehyde (981,the possibility has to be seriously considered that hydroxymethylhydroperoxide is involved in the reaction [see S. Marklund, BBA 258, 9 (1972), and S. Marklund, Acta Chem. Scund. 25, 3517 (1971)l. 101. Although it was originally proposed (102) that a methanol molecule substitutes for one of water in the coordination sphere of molybdenum in the active site of the enzyme, this explanation cannot be reconciled with later data (98). 102. V. Aleman-Alernan, K. V. Rajagopalan, P. Handler, H. Beinert, and G. Palmer, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 380. Wiley, New York, 1965. 103. K. V. Rajagopalan and P. Handler, JBC 242, 4097 (1967). 104. G.Barry, E. Bunbury, and E. 12. Kennaway, BJ 22, 1102 (1928). 105. J. M. Peters and D. R. Sanadi, ABB 93, 312 (1961).

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R. C. BRAY

trophotometrically) with arsenite, the reverse protection is only partial, i.e., arsenite-treated enzyme still, to some extent, seems to react with cyanide (96). Massey and Edmondson (62) have proposed that the arsenite residue binds reversibly between their persulfide group and a neighboring thiol in the enzyme-active center. Other modification reactions for xanthine oxidase which have not been studied in detail may be mentioned briefly. The enzyme is relatively sensitive to inactivation by photooxidation and is also inactivated by H202 ( 3 9 ) , although the mechanisms of these processes are quite unknown. Finally, we note that xanthine oxidase is comparatively resistant to denaturation by urea, although the reagent is a competitive inhibitor of the enzyme (106).Denaturation both of the milk and of the pig liver enzyme, even by 6 M urea, is quite slow (106,107). 4. Redox-Active Groups and Their Magnetic and Optical Properties

a. General Comments. Iron, molybdenum, and FAD in xanthine oxidase would all be expected to be capable of taking up different oxidation states as the enzyme molecule is oxidized or reduced. Though optical spectroscopy has been useful in providing information on oxidation states of the chromophores, EPR has been much more revealing. I n the following subsections, we shall attempt to consider the properties of the chromophores, in their various oxidation states, insofar as is possible considering each chromophore in isolation from the others. Discussion of interactions among the chromophores will be reserved to Section II,B,5, while questions relating to interaction with substrates and to oxidation and reduction processes are considered in Section I1,C. b. “Additional” Redox-Active Groups. Information about additional possible redox-active groups in xanthine oxidase, whose functioning might not be associated with readily observable spectroscopic changes, should, in principle, be obtainable from studies on the stoichiometry of reduction of the enzyme. Without considering the matter in detail here, we may note that reduction of the FAD, Mo, and Fe/S centers in oxidized xanthine oxidase, to levels which seem to be accepted as being reasonable by all workers in the field, would require a total of six electrons per half xanthine oxidase molecule (108). Edmondson et al. (44) and Olson et al. (46; Fig. 4) observed close to this expected stoichiometry in reductive titrations of xanthine oxidase with sodium dithionite, when judging the 106. K. V. Rajagopalan, I. Fridovich, and P. Handler, JBC 236, 1059 (1981). 107. P. E. Brumby, R. W. Miller, and V. Massey, JBC 240,2222 (1964). 108. Based. on the following oneelectron reactions in each half xanthine oxidase molecule: FAD -+ F A D H -+ FADH2; Mo(V1) + Mo(V) -+ Mo(1V); Fe/S I,, -+ Fe/S Ired;Fe/S II,, -+ Fe/S IIred.

6.

MOLYBDENUM HYDROXYLASES

327

end point of the titration by the completion of visible or EPR spectroscopic changes relating to flavin and to iron in the enzyme (109). On the other hand, if the end point was judged instead by appearance of free dithionite in solution (measured a t 315 nm) then some two further reducing equivalents appeared necessary (44) to complete the reaction, bringing the total to eight. The nature of the spectroscopically “invisible” group in the enzyme which accepts these two additional equivalents is uncertain. One possibility might be (cf. 2) that it is a disulfide with unusual properties. The group will be mentioned again in the next subsection in relation t o the origin of one of molybdenum EPR signals (the Slow signal). In any case it is important to note (44) that active and desulfo xanthine oxidases both require the same number of reducing equivalents in dithionite titrations, whichever end point is accepted. Thus, loss of the persulfide group from the active center of the enzyme does not affect the overall stoichiometry of the reduction. c. Flavin Adenine Dinucleotide. It is well known that enzyme-bound FAD, in common with the free coenzyme ( I l l ) , can exist in three oxidation states of which only the semiquinone is paramagnetic and further that each of these can exist in different states of protonation (112-114). The EI’R spectrum of the flavin semiquinone in partially reduced xanthine oxidase was first observed in 1959 (24,25). Care must be taken that the FADH’ EPR signal is not confused with that of the superoxide ion (115) which overlaps with it (Fig. 2). This has not always been done in earlier work. The flavin signal has a g value of 2.0035 k 0.0004 (116) 109. In earlier work work by the same group (36),apparently using the same methods, a stoichiometry of eight electrons per half xanthine oxidase was reported. This discrepancy, which was not commented on (44), must presumably have resulted from inadequate anaerobiosis, in these technically very difficult experiments, in the earlier work. A similar discrepancy seems to relate to reductive titrations with xanthine. By the E,, method, seven electrons were reported (36) to be required and this value was then confirmed (110) by uric acid determinations. Nevertheless, a value of six electrons has been accepted in later work (46‘). 110. V. Massey, H. Komai, G. Palmer, and G. B. Elion, Vit. Horm. (New Yo&) 28,505 (1970). 111. H. Beinert, “The Enzymes,” 2nd ed., Vol. 2, Part A, p. 339, 1960. 112. A. Ehrenberg, G.Eriksson, and P. Hemmerich, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and Morrison, eds.), Vol. 1, p. 179. Wiley, New York, 1965. 113. F. Muller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107.Univ. Park Press, Baltimore, Maryland, 1971. 114. G. Palmer, P. Muller, and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 123. Univ. Park Press, Baltimore, Maryland, 1971. 115. P. F. Knowles, J. F. Gibson, F. M. Pick, and R. C . Bray, BJ 111, 53 (1969). 116. R.C.Bray and T. Vanngtd, BJ 114, 725 (1969).

328

R. C. BRAY

and a linewidth of 19.4 5 1 (117), which decreases in D 2 0 (116). These values and behavior are characteristic of the neutral form of the flavin semiquinone (114). The FADH' E P R signal from the enzyme is considerably more difficult to saturate than is that from metal-free flavoproteins (118).

The precise contribution of flavin, in its different oxidation states, to the visible absorption spectrum of xanthirie oxidase is relatively difficult to. determine owing to the high absorption of the iron chromophores. Its contribution, in the oxidized state, as deduced from the difference spectrum of deflavo enzyme relative to native, is shown in Fig. 1 and is comparable to the spectrum of simple flavoproteins ( 6 3 ) . The semiquinone state is associated with increased absorption in the region of 610 nm (35; cf. 119),as would be expected for the neutral flavin radical ( l l 7 , l 2 O , l 2 l ) , while the spectrum of fully reduced flavin in the enzyme also appears normal ( 6 3 ) .Flavin makes little, if any, contribution to the CD or MCD spectra of xanthine oxidase (60,63,122).Maximum conversion of flavin in the enzyme to the semiquinone decreases with increasing pH, about 20% conversion being achievable.at pH 6, falling to zero a t pH 10 (45). d. Molybdenum. Molybdenum in xanthine oxidase might in principle occur in any valency from six downward, although the lower valencies are strong reducing agents. It seems ( 6 ) , however, that the metal, whatever its valence state, makes relatively little contribution to the absorption spectrum of the enzyme. Thus, knowledge of the state of its molybdenum has rested almost exclusively on E P R studies and detailed work by this method has indeed added greatly to our understanding of the enzyme. A considerable number of different molybdenum EPR spectra may be obtained, under appropriate conditions, from xanthine oxidase. Examination of the conditions under which a given signal appears and disappears can give information about reactions of the enzyme, while examination of the form of the signal can give information about the chemical nature of the signal-giving species itself. It is, however, first necessary to know whether or not an observed E P R signal corresponds to one or more chemical entities. This question has to be answered by comparing the form 117. G.Palmer and V. Massey, JBC 244, 2614 (1969). 118. H.Beinert and P. Hemmerich BRRC 18,212 (1965). 119. R.C.Bray, BJ 83, 11P (1962). 120. F. Miiller, M. Briistlein, P. Hemmerich, V. Massey, and W. H. Walker, EUT.J . Biochem 25, 573 (1972). 121. E.G.Land and A. J . Swallow, Biochemistry 8,2117 (1969). 122. K. Garbett, R. D. Gillard, P. F. Knowles, and J. E. Stangroom, Nature (London) 215, 824 (1967).

6.

MOLYBDENUM HYDROXYLASES

329

of the experimental signal with the standard forms of EPR spectra. If the comparison can be extended to include spectra obtained a t two different microwave frequencies (e.g., 9 and 35 GHz) and to include computer simulations of the spectra, then a more rigorous answer can be given about the number of species involved. In general, each molybdenum E P R signal, from a single chemical species, will correspond to molybdenum atoms in xanthine oxidase molecules (or in the molecules of chemically modified forms of enzyme), which are in a distinct and well-defined chemical environment. Thus, since molybdenum spectra for the enzyme are particularly numerous and well resolved, E P R is clearly a very powerful tool in studying this metal in xanthine oxidase. Many M O W ) compounds as well as some Mo(II1) compounds give well-defined E P R spectra (123) and xanthine oxidase, too, as stated, gives a number of signals which clearly result from this metal. From the first observation (25) onward, it has usually been assumed that these signals from the enzyme arise from molybdenum in the five-valent state. For three out of four known molybdenum signals for xanthine oxidase, all the data do indeed point to the correctness of this assumption. However, the possibility does have to be considered that Mo(II1) may be responsible for the Slow signal, although as will be discussed below, here, too, the balance of evidence would appear to favor Mo (V). E P R cannot, of course, give information (except in a negative way) about the diamagnetic valence states of molybdenum, Mo (VI), Mo (IV), and Mo (11). Molybdenum E P R spectra from xanthiiie oxidase may be observed on solutions of the enzyme at room temperature (98,117,124,125)b u t are more frequently studied on frozen aqueous solutions a t low temperatures ( 6 ) .Solution and frozen spectra of a given sample are always of the same form, showing marked anisotropy, which is not averaged in the solution spectra, because of the high molecular weight of the protein. Thus, the signals always exhibit the typical form of rhombic “powder” spectra (166,127).Molybdenum is a mixture of magnetic ( I = g) and nonmagnetic isotopes, giving six-line and single-line E P R spectra, respectively. The ~ ,relatively low in amplitude and the former, due t o 85Moand 9 7 Mare 123. B. A. Goodman and J . B. Raynor, Advan. Inorg. Chem. Radiochem. 13, 135 (1970). 124. T.Viinngbrd, R. C. Bray, B. G. Malmstrom, and R. Pettersson, in “Free Radicals in Biological Systems” (M. S. Blois, Jr.,et ul., eds.), p. 209. Academic Press, New York, 1961. 125. J. C. Swann and R. C. Bray, Eur. J. Biochem. 26,407 (1972). 126. H.Beinert and G. Palmer, Advan. Ensymot. 27, 105 (1965). 127. P. W.Atkins and M. C. R . Symons, “The Structure of Inorganic Radicals,” p. 268.Elsevier, Amsterdam, 1967.

330

R. C. BRAY

g- nlu..

2.1

2.0

FADH* FADH*

1.v

4,

alpadd.

3.1

1.0

1.8

g nlu..

FIQ.2. Electron paramagnetic resonance spectra encountered in work on milk xanthine oxidme. All the spectra are experimentally recorded ones, except for those from FADH' and from Fe/S 11; these are recorded as difference spectra obtained after subtracting other signals. All samples were prepared at pH 8.2, except for the Very Rapid signal and the superoxide signal, for which higher pH values were employed. Superoxide w m generated by pulse radiolysis (3.9).Spectra were recorded a t 9.3 GHs and 120DK,except for Fe/S I and Fe/S 11, which were a t 50" and 25'K, respectively. Some FADH' is visible in the Rapid (xanthine) sample.

single-line spectra thus dominate the observed signals. Observation of the six-line spectra is facilitated when enzyme enriched in 8 6 Mis~ employed. This has to be prepared by in v w o synt'hesis, by injecting cows with the isotope and isolating the enriched enzyme by standard procedures (73,98).

6.

MOLYBDENUM HYDROXYLASES

331

Resting, oxidized, milk xanthine oxidase normally gives no E P R signals and is assumed to contain Mo(V1). Signals develop only under reducing conditions; EPR data on the enzyme have been reviewed fairly fully by Bray and Swann (6). The parameters and origins of the various molybdenum signals which have been described for the enzyme are summarized in Table I11 (6,99,116,127a,1.28). The nomenclature adopted for the signals is that of Bray and Vanngbd (116),signals being given the names Very Rapid, Rapid, Inhibited, and Slow. These signals are all relatively similar t o one another, e.g., their gav values fall within the range 1.971 -t- 0.006. As will be discussed in Section II,C,2,e, the Very Rapid and Rapid signals appear and disappear in times comparable to the turnover time of the enzyme and are believed to represent catalytically important intermediates. Superficially these signals are related to one another simply by uptake of a proton by the Very Rapid signal, to give the Rapid (6,128), although the precise chemical relationship between the signal-giving species is not certain. I n contrast to these signals, the Inhibited and Slow signals are not involved in turnover. Detailed examination of the Very Rapid, Inhibited, and Slow signals indicates that they all represent single chemical species (6). On the other hand, the Rapid signal corresponds to a family of closely related chemical species, all having very similar EPR parameters (116).The precise form of the Rapid signal observed thus depends on the relative amounts of these species in the sample. Bray and VanngBrd (116) and Pick and Bray (129) proposed that enzyme molecules, in the state of reduction giving the Rapid signal, can enter into complex formation with substrate molecules. Each complex would be a distinct chemical species with its own EPR parameters. Thus, the nature and extent of the complex formation would determine the precise form of the Rapid signal (150,151).Later work in the author’s laboratory (unpublished; also see 46) has shown 127a. D. J. Lowe, Ph.D. Thesis, London University, 1974. 128. D. Edmondson, D. Ballou, A. Van Heuvelen, G. Palmer, and V. Massey, JBC 248, 6135 (1973). 129. F. M. Pick and R. C. Bray, BJ 114, 735 (1969). 130. I t was suggested in Pick and Bray (1.29) that the species giving rise to the Slow signal (i.e., desulfo xanthine oxidase) might also enter into complex formation with substrates, although the evidence for this was not conclusive and later work has rendered it somewhat unlikely. 131. As already mentioned in Section II,B,2,c, it was also proposed (116), that existence of two Rapid signal species, in the absence of complex formation with substrates, might be indicative of the two molybdenum atoms of the enzyme molecule being in slightly different environments. Further unpublished work in the author’s laboratory indicates, however, that this interpretation is unlikely to be correct.

0 0

N

TABLE 111 MOLYBDENUM EPR SIGNALSFROM REDUCED FORMS OF XANTHINE OXIDASE~

Signal

Origin

Very Rapid

Active enryme

Rapid

Active enzyme

Inhibited

HCHO or CHaOH

Slow

treatment of active enzyme Desulfo enzyme

a

Treatment required

Sav

81

IA..I (1H) (G)*

Exchange of interacting 'H with ZHzO

Notes

-

-

12-14d

Yes

3. 9-5.6

No

Several related species Stable in air

16

Yes

-

Reduction for very short times, a t high pH, with xanthinc,c only Reduction with any substrate None

1.977

2.025

None

1.9731.974 1.973

1.9891.994 1.953

Reduction for long periods (e.g., 20 min with dithionite)

1.965

1.956

Data are from Bray and Swann (6) except for the Slow signal for which a revised value (227a) is given.

* The sign of the splittings is not known. Values given are the highest and lowest numerical values observed along the principal axes. A little Very Rapid signal is also obtainable with 1-methylxanthine (128) and a few other substrates (99). Additional smaller and apparently more anisotropic splittings were also reported (226).

FJ P W

E

6.

MOLYBDENUM HYDROXYLASES

333

that not only substrates but also products (particularly uric acid) form complexes affecting the Rapid signal (131a). I n principle, g values and 9 5 M splittings ~ are capable of giving information on the environment of molybdenum in the enzyme, but this is difficult to interpret in the absence of direct information, either on the nature of the ligands or on the symmetry. Only oxygen, sulfur, or nitrogen ligands are of course expected in a protein. Meriwether et al. (139) Buggested, from comparison of the EPR parameters with a limited number of model compounds, that the metal in the enzyme might have one or more sulfur ligands. More recently, knowledge of the structures of Mo (V) complexes and understanding of their EPR spectra has advanced considerably, particularly from the work of Marov and co-workers (133-138). Further studies on the enzyme, especially more detailed work on the Rapid signals and remeasurement of some of the 9 5 M hyperfine ~ splittings, may perhaps be needed before this newer information on the low molecular weight Mo(V) complexes can be made full use of when attempting to elucidate the environment of the metal in the enzyme. However, a complex (133) of molybdenum with quinoline-8-thiol seems particularly interesting. This contains two molecuIes of the chelating agent, each bound to the metal via a nitrogen and a sulfur, with the probable structure shown in Scheme I (where Y represents water or a weakly bound anion). Its EPR parameters (Table IV) are all quite similar to those of the Very Rapid signal (73), which are shown for comparison. The finding of a low molecular weight model compound, with only oxygen, nitrogen, and sulfur ligands and with a type of structure which one might reasonably imagine to occur in a protein and which mimics 131a. It is now clear that under appropriate conditions a considerable number of different species may be obtained. Some slight revision of the details of the analysis of Rapid signals obtained with xanthine, as reported by Bray and Vinngbrd (1161, will probably be needed. 132. L. S. Meriwether, W. F. Marzluff, and W. C. Hodgson, Nature (London) 212, 465 (1966). 133. I. N. Marov, V. K . Belyaeva, Y. N. Dubrov, and A. N. Ermakov, Rwss. J . Inorg. Chem. 17, 515 (1972). 134. I. N. Marov, in “The Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 63. Climax Molybdenum Co., London, 1973. 135. I. N. Marov, Y. N. Dubrov, V. K. Belyaeva, and A. N. Ermakov, Rzcss. J . Inorg. Chem. 17, 1396 (1972). 136. I. N.Marov, E. M. Reznik, V. K. Belyaeva, and Y. N. Dubrov, Russ. J . Inorg. Chem. 17, 700 (1972). 137. I. N. Marov, U. K. Belyaeva, Y. N. Dubrov, and A. N. Ermakov, Russ. J . Inoi-g. Chem. 17, 1561 (1972). 138. I. F. Gainulin, N. S. Garif’yanov, and V . V . Trachevskii, Izv. A k a d . N a u k SSSR, Ser. Khim. 10, 2176 (1969); Bull. Acad. Sci. USSB., Div. Chem. Sci. 10, 2030 (1970).

334

R.

C. BRAY

SCHEME I. The structure of a complex between Mo(V) and quinoline-&thiol, aa suggested by Marov d a?.(133).

the enzyme closely in all its E P R parameters, may indeed seem exciting to those working in this area. Though it may be prudent not to pursue structural analogies too far a t this stage (139), it is indeed tempting to suggest that there is a very similar arrangement of ligands around molybdenum in the Very Rapid signal-giving species to that in the complex of Scheme I. The study of hyperfine interactions, resulting from ligand and other atoms with magnetic nuclei which interact with molybdenum in the enzyme, can also yield valuable information on the environment of the metal. Unfortunately, no nitrogen hyperfine splittings are resolved. This does not, however, exclude the possibility that there are nitrogen ligands of molybdenum in the enzyme (140). For instance, Marov et d.(133) reported, for the compound of Scheme I, no resolved nitrogen hyperfine splitting but indications, nevertheless, of line broadening resulting from unresolved nitrogen hyperfine structure. The only hyperfine structure (other that that from Q 6 M ~which ), is resolved in xanthine oxidase, is due to protons (6,141). This unusual feature of the spectra first became apparent during work with the Q 5 Menzyme ~ by Bray and Meriwether (73). Proton splittings are observed in all the molybdenum signals from the enzyme, except the Very Rapid, and range up to 16 G in magnitude (Table 111).The interacting protons of the Rapid and Slow signals are exchangeable with solvent water molecules ( 6 ) .Only more recently have comparable molybdenum-proton interactions been observed in low molec139. Cotton ( 1 3 9 ~ )emphasizes the possible importance, in substrate binding to enzymes, of 0x0 groups on molybdenum labilizing the trans ligand and so making way for the substrate molecule. On the other hand, work by Stiefel and Gardner (139b) may perhaps argue against an 0x0 group in xanthine oxidase. 139a. F. A. Cotton, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 6. Climax Molybdenum Co., London, 1973. 139b. E. I. Stiefel and K. G. Gardner, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 272. Climax Molybdenum Co., London, 1973. 140. Nitrogen ligands to molybdenum in the enzyme are quite reasonable on general chemical grounds [see R. J. P. Williams and R. A. D. Wentworth, in “Chemistry and Uses of Molybdenum’’ (P. C. H. Mitchell, ed.), p. 212. Climax Molybdenum Co., London, 19731. 141. R. C. Bray, P. F. Knowles, F. M. Pick, and T. Vanngird, BJ 107, 601 (1968).

g

z

3 COMPARISON OF T H E

EPR

PARAMETERS O F T H E

TABLE IV VERY RAPIDSIGN.4L

WITH

THOSE OF

. 4

MO(V)-QUINOLINE-&THIOL COMPLEX [ A ]values (961LIo) (G)

g values

Species

9a"

9. or 911

QU

9,

9..

A,,

9,orAll

A,

A,

A,

Very Rapida Quinoline-8thiol complexb

1.977 1.981

2.026 2.019

1.956 -

-

1.961

1.951 -

34 34

41 50

24 -

24

-

a

b

Ex! 3 % b?

37

From Bray and Meriwether (73). From Marov et al. (1%).

W

W cn

336

R. C. BRAY

ular‘weight complexes (138,142,143). Study of the origins of the interacting protons in the Rapid signal from the enzyme has been of interest in relation to the catalytic mechanism and will be discussed subsequently (Section II,C,2,f). In the case of the Inhibited signal, the proton is nonexchangeable with the solvent and its origin, from formaldehyde or methanol, has already been considered in Section II,B,3,f. The question of whether the interacting protons, particularly in the Rapid signal, are attached directly to the metal to form a molybdenum hydride or whether there are intervening atoms between them has been the subject of some controversy. Bray and Vanngbrd (116) assumed that the signs of the proton hyperfine splittings were all the same and concluded that the maximum anisotropy (i.e., difference in IAl values along the principal axes) was about 2 G. A simple dipolar coupling calculation from this, indicated that the Mo-H distance must be greater than 3 A, thus excluding the possibility of a hydride. On the other hand, plausible catalytic mechanisms for the enzyme may be written, involving a molybdenum hydride. This and other arguments led Edmondson et al. (168) to question the assumption of the splittings having the same sign. These workers pointed out that for a Mo-H distance of 1.5 A, if different signs are allowed, a particular combination of isotropic and dipolar couplings could lead to the observed [A/ values of about 12 G along each of the principal axes. Attractive as such reasoning may be, it does not appear satisfactory when considered in relation to data on the Slow and Inhibited signals (Table 111),as well as to that on signals from xanthine dehydrogenase from 8.izlcalescens (144,145) and from turkey (90b). All these molybdenum signals show the same apparent near isotropy of proton splittings as does the Rapid signal from milk xanthine oxidase, although the actual magnitude of the splittings varies considerably from one signal to another. It seems highly unlikely, in these circumstances, that the proposed (128) fortuitous cancellation of anisotropy could be occurring in all cases. I n any case, the idea of a hydride structure in the enzyme seems to have been abandoned, apparently on chemical grounds, by Massey and co-workers in a later publication (46‘) (see Section II,C,2,f). 142. P. C. H. Mitchell and R. D. Scarle, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 261. Climax Molybdenum Co., London, 1973. 143. E. I. Stiefel, in “Chemistry and Uses of Molybdenum” (P. C . H. Mitchell, ed.), p. 284. Climax Molybdenum Co., London, 1973. 144. The organism referred to in this chapter as Veillonella alcalescens has also, frequently but incorrectly, been called Micrococcus lnctilyticus [see H. Dalton and J. Zubieta, BBA 322, 133 (1973)l. 145. H.Dalton, D.J. Lowe, R. T. Pawlik, and R. C. Bray, BJ (in preas).

6.

337

MOLYBDENUM HYDROXYLASES

The original proposal (116) that molybdenum is not directly bonded to hydrogen must therefore stand. However, the nature of the intervening atom or atoms is not certain and oxygen, nitrogen, or sulfur all seem possible candidates. Bray and Vanngdrd (116) concluded that it could not be nitrogen, owing to the lack of nitrogen hyperfine structure. However, this conclusion must be revised (145a), since Mitchell and Scarle (142) have reported proton splittings of up to 20 G, without nitrogen splittings, in low molecular weight complexes of Mo(II1) believed to have an Mo-NH structure. A further matter of controversy, regarding molybdenum E P R signals from xanthine oxidase, concerns the valency of the metal in the Slow signal. This signal had been known (99) for some while before it became apparent that it resulted from the desulfo form of the enzyme (58,66; see Section II,B,3,6). Massey and co-workers have repeatedly suggested that the Slow signal may result from Mo(II1) (46,110,117), whereas Bray and co-workers have assumed it is from Mo (V) (6,6). The balance of available evidence seems to favor the latter view as discussed below. Information a s to the valency might, in principle, come either from the EPR parameters of the Slow signal or from consideration of the conditions under which the signal appears and disappears. Two alternative reduction schemes to explain appearance of the Slow signal from the desulfo enzyme are given in Eqs. (2) and (3) : Mo(V1) F! Mo(V1)

Mo(V) (no signal)

Mo(V) (Slow signal) F! Mo(1V)

* Mo(1V) Mo(II1) (Slow signal)

(2) Mo(I1)

(3)

Such data as are available from the g values tend to favor Mo(V) in the Slow signal. Thus, Mo(II1) complexes appear to have higher g values than those of related Mo(V) complexes (1&b), whereas the Slow signal has the lowest gay among signals from the enzyme (Table 111). Dithionite titrations of xanthine oxidase (Section II,C,2,d) have revealed a number of points. First, it is clear that since a t least 2-3 moles of dithionite have to be added to the enzyme, per half molecule, before the Slow signal starts to develop (46,146; Fig. 3c), a lower redox potential is re145a. T. Vinngbrd, personal communication. 145b. P. C. H. Mitchell and R. D. ScarIe, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), pp. 140 and 279. Climax Molybdenum Co., London, 1973. 146. H. Beinert, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 416. Univ. Park Press, Baltimore, Maryland, 1971.

338

R. C. BRAY

quired for the desulfo enzyme to give the Slow signal than is associated with any of the chromophores of the active enzyme (see Section II,C,2,f). However, this redox potential cannot be an exceptionally low one, since reduction of enzyme samples with NADH generates the Slow signal (126).It has been suggested (46) that the above stoichiometry in the titrations favors Mo(II1) in the signal. But this argument is complicated by uncertainty about the nature of the “additional” redox-active groups (Section II,B,4,b) (147) and is weakened if it is accepted that large excesses of dithionite can ultimately cause the Slow signal to decrease in intensity (148). This decrease would presumably have to involve, according to Eq. ( 3 ) ,reduction of the Mo(II1) to Mo(I1) (149).Such reduction seems highly improbable on chemical grounds since molybdenum compounds are stated to be generally not reducible below the three-valent state in aqueous solution (150).Finally, perhaps the most convincing evidence for the Slow signal coming from Mo(V) according to Eq. (2) is its appearance in preliminary experiments by the author (91a) in which desulfoxanthine oxidase was reduced by low doses of the hydrated electron in pulse radiolysis (32). It is unlikely that more than one electron could be taken up by the enzyme under the conditions of these experiments. Thus, we assume that the Slow signal, like the other molybdenum signals from the enzyme, results from Mo (V) . (Kinetic considerations relating to the Slow signal will be taken up in Section II,C,2,k). If it is accepted that both Rapid and Slow signals come from Mo(V), then one is in a position to attempt to interpret the difference in their 147. To account for the stoichiometry of the titrations (46) in terms of Eq. (2), one might assume, for instance, that whereas in the active enzyme, the redox potential of molybdenum is above that of the spectroscopically “invisible” electron acceptor, on the other hand, in the desulfo enzyme, this situation has

become reversed. 148. All workers seem to agree that with large amounts of dithionite the Slow signal decreases in intensity but quantitative aspects are less certain. Maximum intensity of the signal is attained with about 4 moles of dithionite per half xanthine oxidase molecule (46,146). Pick and Bray (129) reported extensive decreases in intensity with large excesses of the reducing agent, these decreases occurring at rates dependent on the dithionite concentration. Later work in the author’s laboratory (cf. 1 4 8 ~ )showed similar, though less marked effects. According to data of Massey and co-workers (461, addition of a further 2 moles of dithionite, beyond the quantity required for maximum Slow signal, resulted in a decrease in its intensity of some 15%. 148a. F. M. Pick, Ph.D. Thesis, University of London, 1971. 149. I n earlier work (110,117) Massey and co-workers favored occurrence of Mo(I1) in’ the enzyme but later (46) their data necessitating it seems to some extent to have disappeared (cf. 109). 150. P. C. H. Mitchell, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 1. Climax Molybdenum Co., London, 1973.

6.

MOLYBDENUM HYDROXYLASES

339

EPR parameters in terms of differences in the environments of molybdenum in the active and desulfo enzymes, respectively. The main features (Table 111) are a change from gll > gL to g, > 911 and a decrease in gav of 0.008 to 0.009. Such changes would seem entirely consistent with those which might occur (cf. 1%) on exchange of a single ligand of the metal. It was earlier suggested (5,125) that the data might be explained if a sulfur ligand in the native enzyme, which could possibly be the persulfide sulfur atom (Section IIlB,3,e), had become replaced in the desulfo enzyme b y an oxygen ligand. However, replaccment of sulfur by nitrogen now seems no less possible (151). Perhaps further E P R work with model compounds would now be helpful in elucidating the precise nature of the reaction which takes place when sulfur is lost from the active center of functional xanthine oxidase to yield the desulfo form. e. IronAuZfur Systems. Knowledge of the iron-sulfur systems of xanthine oxidase, like that on the molybdenum, rests primarily on E P R data, but in this case much supplementary information is also available from other physical methods. There are two different iron-sulfur systems in the enzyme, which have, however, so far, been distinguished solely on the basis of their E P R spectra in the reduced state. Each Fe/S system can exist in a signal-free oxidized state or a signal-giving reduced state. The systems have been referred to as the “gaV= 1.95 Fe/S system” and the “gav = 2.01 system” ( 1 5 3 ) ,or alternatively, as “Fe/S system I” and “Fe/S system 11” (128), respectively. Since the latter nomenclature is less cumbersome, it will be adopted here. Integration of the EPR signals indicates (117)that Fe/S I and Fe/S I1 each accounts for one electron per half xanthine oxidase molecule in the fully reduced enzyme. Thus, by analogy with the plant ferredoxins (1541, i t is presumed (though without rigorous proof) that Fe/S I and Fe/S I1 are both two-iron-two-labile sulfur systems, there being one system of each type in each half enzyme molecule, so accounting for the total content of 8 Fe atoms per xanthine oxidase molecule. While helium temperatures are necessary for detailed studies on the 151. Marov el al. (136) found t h a t the complex MoO[CNS14Y (cf. Scheme I) [in which the thiocyanate ions are probably coordinated via N(16.2)] has gav 1.963 and gli 1.931. On the other hand, a complex with the probable formula MoO[CNS]s[SRIY (where SR is thioquinoline, coordinated via S) has g., 1.950 and gll 1.972. Here then, apparently, replacement of S by N causes gll to shift from above g, to below, as in the Rapid-to-Slow transition, although the change in gay is in the opposite sense from that in the enzyme. 152. F. A. Cotton and G. Wilkinson, “Advanced Inorganic Chemistry,” 3rd ed., p. 970. Wiley (Interscience), New York, 1972. 153. D. J. Lowe, R. M. Lynden-Bell, and R. C. Bray, BJ 130, 239 (1972). 154. W. H. OrmeJohnson, Annu. Rev.Biochem. 42, 159 (1973).

w

*

0

TABLE V PROPERTIES OF THE I R O N S U L F U R C E N T E R S OF X a N T H I N E OXIDASE ~

~

Property

Fe/S I

Ref.

1.95 2 . 0 2 2 , l . 9 3 5 , l .899 40" 10

155 153 127a

Fe/S I1

Ref.

EPR properties (reduced enzyme) L

V

911

g%J 98

Fully sharpened at ("K) Microwave power for 50% saturation at 15°K (mW) Optical properties (tentative) AcrZmd (450 nm) mM

Ar,,-,,

(550 ~ ) / A E : % ~ (450 nm)

11.6 0.48

45 45

2.01 2.12,2.007,1.91 25" 60

2.8 0.46

153 127a ld7a

45 45

6. MOLYBDENUM

HYDROXYLASES

341

EPR spectra (Table V),the signal (155) from Fe/S I, though in a much broadened form, is detectable at nitrogen temperatures and was first observed and ascribed to iron in the reduced enzyme in 1961 (96).The signal (153) from Fe/S I1 was not reported until later (117,156) when helium EPR cooling systems had become available. The parameters (Table V) of the Fe/S I signal are entirely comparable with those of typical 2 Fe/2 S iron-sulfur proteins (154) and the indications are, therefore, that Fe/S I must have a structure like that deduced (157) spectroscopically for spinach and parsley ferredoxins and later substantiated (158) by model studies. On the other hand, Fe/S I1 is apparently unique among Fe/S proteins, in having only one g value below the free electron value (153). Although the structural implications of this are not clear, the model of Dunham et al. (157) is presumably not directly applicable to Fe/S 11. Since molybdenum probably makes little contribution, the visible spectrum of deflavo xanthine oxidase must primarily result from Fe/S I and Fe/S 11, and this spectrum (Fig. 1) is indeed not unlike that of other iron-sulfur proteins. However, it is no easy matter to determine the individual contributions of the two systems to the net spectrum. Only recently and very tentatively has an attempt a t this been made, by Olson et al. (46; Table V). The oxidized-minus-reduced difference spectra for Fe/S I and Fe/S I1 seem to have similar forms to one another, but that of Fe/S I is apparently some four times more intense than Fe/S 11. ORD, CD, and MCD spectra of xanthine oxidase have all been recorded (6'0,117,122) and no doubt primarily result from the Fe/S systems but seem to have added relatively little, so far, to understanding of the enzyme and have not distinguished between the two Fe/S centers. Two other types of measurement on xanthine oxidase, though not so far taken to a high degree of sophistication, have nevertheless yielded valuable information on its Fe/S systems. Mossbauer measurements (28,169) have only been made at poor signal-to-noise ratios, since 57Feenriched enzyme was not available (160). However, they did, in 1969 155. J. F. Gibson and R. C. Bray, BBA 153, 721 (1968). 156. W. H. Orme-Johnson and H. Beinert, BBRC 36,337 (1969). 157. W. R. Dunham, G. Palmer, R. H. Sands, and A. J. Bearden, BBA 253, 373 (1971). 158. J. J. Mayerle, R. B. Frankel, R. H. Holm, J. A. Ibers, W. D. Phillips, and J. F. Wieher, Proc. Nat. Acud. Sci. U . S. 70,2429 (1973). 159. C. E. Johnson, R. C. Bray, R. Cammack, and D. 0. Hall, Proc. Nut. Acud. Sci. U . S. 63, 1234 (1969). 160. Iron has not so far been reversibly removed from xanthine oxidase, and in vivo enrichment of the enzyme from cow's milk (as was done for "Mo) would be prohibitively expensive.

342

R.

C.

BRAY

(1591, provide evidence for something which is still of interest, namely, that all the iron atoms of the xanthine oxidase molecule change their environments on reduction of the enzyme. Finally, careful magnetic susceptibility measurements were carried out by Ehrenberg and co-workers (26,27) on xanthine oxidase, a t room temperature, as long ago as 1961. These provided the first evidence for the now well-known fact that iron-sulfur proteins become more paramagnetic on reduction. 5. Magnetic Interactions among the Redox-Active Groups and

Information on Their Relative Locations within the Enzyme Molecule When a macromolecuIe contains more than one paramagnetic center then there may be magnetic interactions among the various centers. The extent to which such interactions are observed will depend on the distances and relative orientations of the centers as well as on the nature of the intervening groups. Consequently, studies on the interactions can, in principle, give information about the structure of the macromolecule. Unfortunately, the theory of interactions of this type is not well understood (but see, e.g., 161). It is to be hoped that the unique way in which paramagnetic centers in a globular protein are fixed relative to one another, in contrast to the random distribution of such centers, e.g., when they are present at low concentrations in inorganic crystals, may attract more physicists to work in this potentially rewarding area. In the case of xanthine oxidase, a t the present time, in the absence of X-ray crystallographic work, magnetic interaction methods seem t o be the only ones capable of yielding information on the relative locations of the Fe/S, molybdenum, and FAD within the intact enzyme molecule. However, their application still seems in its infancy. I n general, if two S = paramagnetic centers are very close together in the same molecule, no EPR signal will be observed. Thus, one might be tempt,ed to conclude from the fact that it is easy to prepare reduccd xanthine oxidase samples showing simultaneously Mo(V), FADH', and Fe/S signals that these centers are not close together in the enzyme. However, such a conclusion would onIy be valid if' it could be excluded that different individual enzyme molecules were in different states of reduction in the sample, so that, for example, some molecules showed Mo(V) signals only, while others showed FADH. signals only, and so on. This raises problems about the various states of reduction of the enzyme which will be considered in Section 1I,C12,f. The first report of magnetic interaction in xanthine oxidase was the finding of Beinert and Hemmerich (118), already mentioned in Section II,B,4,c, that relaxation of the FADH. radical, as indicated by its EPR

+

161. J. S. Leigh, J. Chem. Phys.

a,2608 (1970).

6.

MOLYBDENUM HYDROXYLASES

343

saturation behavior on varying the microwave power, is abnormally fast. In later work (e.g., 162; on aldehyde oxidase) , flavin saturation studies were extended and the conclusion was reached that the interacting paramagnetic centers must be some 10-20 A away from it. However, the nature of the centers interacting with flavin was not established in this work and, indeed, the saturation method does not seem to be capable of yielding detailed information. More recently, Lowe et al. (153) reported that interaction between M O W ) and reduced Fe/S I in xanthine oxidase could give rise to a n additional splitting in the EPR spectrum of the former. This splitting only became apparent as the sample temperature was lowered from about 80° t o 40°K; i.e., appearance of Mo(V) splitting paralleled the normal sharpening of the Fe/S signal as the temperature was lowered. Furthermore, the splitting was not observed in samples having Mo reduced but not Fe/S, confirming that the phenomenon indeed resulted from interaction betweer. the paramagnetic centers. The expected, corresponding splitting of the Fe/S signal has not been observed, partly because of the relatively large linewidth of the signal and partly also, presumably, because every molecule containing reduced Fe/S does not also contain Mo (V) (see Section 1I,C12,f). At the semiempirical level, this molybdenum splitting phenomenon appears to provide a means of distinguishing molecules containing Mo (V) only from those containing both Mo(V) and reduced Fe/S I (cf. 153). This should prove of value in studies of different reduction states of the enzyme. At a more fundamental level, understanding the nature of the interaction and the origin of the apparent virtual lack of anisotropy, should yield information about the geometry of the xanthine oxidase molecule. The splitting phenomenon has been observed not only with milk xanthine oxidase for the Slow and the Rapid signals but also for the corresponding signals from the xanthine dehydrogenase of V . Alcalescens (127a). (As will be discussed in Section III,B, EPR evidence is that the latter enzyme has its molybdenum in an environment quite similar to that of the metal in the milk enzyme. On the other hand, the Fe/S systems of the two enzymes are very different from one another.) For these four molybdenum signals from the two enzymes, the low temperature splittings are all apparently almost completely isotropic, although their actual magnitudes vary some 2- to %fold from one signal to another. It seems highly unlikely that the signs of the splittings are different (cf. discussion of Mo-H distances in Section II,B,4,d). A simple dipolar coupling calculation based on the magnitude of the small detectable aniso162. K. V. Rajagopalan, P. Handler, G . Palmer, and H. Beinert, JBC 243, 3797

(1968).

344

R. C. BRAY

tropies, therefore, puts the molybdenum-Fe/S distance in the enzymes I n effect, at the surprisingly large one of about 30 A or more (6,127~). this indicates the centers to be virtually on opposite sides of the xanthine oxidase molecule. It is difficult to see how superexchange interaction could take place over such distances to provide the observed isotropic splittings, unless there were an extended orbital system of some kind between Fe/S and Mo. It has- tentatively been suggested (5,127a) that there is indeed such a system in the enzyme and that electron transfer between the tenters in catalysis takes place via it. However, more work on the problem is required. Provided sensitivity problems can be overcome, a direct and potentially rewarding way of studying interactions between paramagnetic centers is the method of ELDOR (electron-electron double resonance). As this chapter was being prepared ELDOR studies on xanthine oxidase were being attempted in a t least two laboratories and some preliminary results have been obtained (163). Aside from the studies of magnetic interactions, information about the location of flavin in the xanthine oxidase molecule may be deduced simply from comparison of the properties of native enzyme with those of the deflavo form. There are, of course, no FADH. signals from the deflavo enzyme. However, the forms of the various Mo(V) and Fe/S signals from modified enzyme are indistinguishable from those from the native form (6,63,128). Clearly, therefore, in native xanthine oxidase, in the reduced state, flavin is bound directly, neither to molybdenum nor to iron.

C. CATALYTIC PROPERTTES 1. Reactions Catalyzed a. Specificity: Reducing Substrates (Including Other Molybdenum Hydroxylases) . Attempting to review comprehensively the substrate specificity of xanthine oxidase is a difficult task, which is made still more difficult if a comparison with the specificities of other molybdenum hydroxylases is also included. The enzymes catalyze oxidation of a very wide variety of substrates (1,4) and, to make the would-be reviewer's task more difficult, different workers have measured activities toward reducing substrates using a variety of oxidizing substrates, as well as at different pH values and with varying concentrations of both substrates. Furthermore, apart from the normal complications of two-substrate enzyme-catalyzed reactions, xanthine oxidase reactions are frequently particularly sensitive to inhibition by excess substrate (Section 11,CJ2,j). 163. D. J. Lowe and J. S. Hyde, BBA 377, 205 (1975).

6.

MOLYBDENUM HYDROXYLASES

345

Fortunately, Krenitsky et al. (10) made a very comprehensive comparative study of the reducing substrate specificities of xanthine oxidase and aldehyde oxidase. They selected the physiologically significant pH value of 6.8 and obtained relative oxidation rate data, under standard conditions, on more than 50 compounds which are oxidized a t measurable rates by the milk enzyme, as well as reporting negative results on many more compounds. Apart from this work, the systematic studies of Bergmanp and co-workers (e.g., 164-166) must particularly be mentioned. This and other work has been summarized by Massey (1). The only systematic studies on reactivity of the enzyme toward a wide range of aldehydes still seem to be those carried out by Booth in 1938 (167). Some data on xanthine oxidase are summarized in Table VI and are compared with that on rabbit liver aldehyde oxidase (10) and on the xanthine dehydrogenases from Clostridium cylindrosporum (168) and from turkey liver (169). Comparison of rates from different laboratories on a given substrate of milk xanthine oxidase shows variations, in some cases of 2-fold or more. This is of course not a t all surprising, in view of the variations in experimental conditions, as discussed above (see footnotes to Table VI for details of the conditions used) (10,164-172).However, it does mean that, apart from the direct comparisons of Krenitsky et al. ( l o ) ,some caution must be exercized when attempting to compare the various enzymes. Nevertheless, i t is clear that while all four enzymes listed have closely related specificity ranges yet no two are even approximately identical. To mention a few characteristic features, aldehyde oxidase is distinguished from xanthine oxidase by its low activity toward xanthine, although its activity is by no means low toward all purines. Similarly, the enzymes from Clostridium and turkey are particularly distinguishable from the other two in Table VI by their lower activities toward aldehydes, while they are distinguishable from one another by differing reactivities toward adenine and NADH. It is further interesting that the four enzymes differ not only in the rates of oxidation of different substrates but also in some cases in apparent K , values toward a particular substrate which is oxidized rapidly by all the enzymes. This is espe164. 165. 166. 167. 168. 169. 170. 171. 172.

F. Bergmann, H. Kwietny, G. Levin, and D. J. Brown, JACS 82,598 (1960). F. Bergmann and H. Ungar, JACS 82, 3957 (1960). F. Bergmann, L. Levene, Z. Nieman, and D. J. Brown, BBA 222, 191 (1970). V. H. Booth, BJ 32, 494 (1938). W. H. Bradshaw and H. A. Barker, JBC 235, 3620 (1960). W. F. Cleere and M. P. Coughlan, Comp. Biochem. Physiol. SOB, 311 (19'75). D. Gregory, P. A. Goodman, and J. E. Meany, Biochemistry 11, 4472 (1972). I. Fridovich, JBC 241, 3126 (1966). D. M. Valerino and J. J . McCormack, BBA 184, 154 (1969).

TABLE VI OF SOMEREDUCING SUBSTRATES BY XANTHINE OXIDASEAND RELATEDENZYMES OXIDATION Relative ratesbwith apparent K , values [in brackets] for substrate oxidation by the enzyme listed ~~

Milk xanthine oxidase Krenitsky Bergmann et al." et a l . d . 6

Substrate"

CHsCHO

Acetaldehyde

43'

Other workers/

Rabbit aldehyde oxidase"

Clostridium xanthine dehydrogenasee*g

80

12

94i [6 X 10-3]k

Salicylaldehyde

Turkey xanthine dehydrogenase8.h 3'

7%

Purine 12

Hypoxanthine

Xanthine HO

6,B-Dihydroxypurine

'.,

'%g,H

N

(100) x 10-51

34

130 [ < 10-51

120

170 [ < 10-61

(170)

120

170

(130)

(100) [3 x 10-41

[2

3 [ 2 x 10-31

43 11 x 10-21

[2

x

<1

x

46 10-*I

(170) 10-6; 2 x 10-41 51 [4 x 10-41

[4

x

54

100

[3

(170) x 10-61

P

m

a

z

6.

E rl

V

0

rr3

2 t .rCrJ

MOLYBDENUM HYDROXYLASES

I

22

n

X

2

I

-

c?

r-X

T 0

'& r3 I

r4

co

co

0

m

cv

00

0

2

E

v

r3

R

co

su

g

0

m co

347

TABLE VI (Continued) w

Relative ratesbwith. apparent K , values [in brackets] for substrate oxidation by the enzyme listed Milk xanthine oxidase Krenitsky Bergmann et al.0 et a1.d.e

Substrate"

Other workersf

Rabbit aldehyde oxidasec

Clostridium xanthine dehydroeen&seo'c

-

s Turkey xanthine dehydrogenasee-h

OH I

4-Hydrox y-fhminopyrimidine

23"

91

N LMethylnicotinamide

34

<3 [I

x

0

10-a1

I CHs

NADH

15

<1"

79

16

0.7

70

OH

Allopurinol

NN '

k N N

65

P P

OH

7-Hydroxy- (1,2,5)thiadiazolo (3,4-d)pyrimidine

td

31

320

?c

Pteridine

240 [4

PAmino-7-hydroxypteridine

x 10-51

a,

290" OH

OH

UN2 I

4-Hydroxy-7-azapteridine

1000'

Formulas of oxypurines, etc., are written in the hydroxyl form, where possible, to indicate availability of a proton, although in some cases the keto form is the predominant species. b In each column the figure in parentheses i s arbitrary, other values being expressed relative to this. Krenitsky et nl. (10). Rates are expressed relative to that for purine, taken as 100. Measurements were at pH 6.8, with ferricyanide (1 mM) as oxidizing substrate, with reducing substrates at 0.07 m M for xanthine oxidase or at 2.1 mM for aldehyde oxidase. Bergmann et al. (164-166). Measurements were at p H 8.0, with oxygen (approximately 0.25 mM) as oxidizing substrate, with reducing substrates at 0.06-0.07 mM. Rates are expressed relative to that for xanthine, taken as 170, for comparison with data from Krenitsky et al. (10). f Rates are expressed relative to that for hypoxsnthine, taken as 130, for comparison with data from Krenitsky et al. (10). Bradshaw and Barker (168).Measurements were made at pH 7.5, with 2,BdichIorophenolindophenol(0.06 mM) as oxidizing substrate. Reducing substrates were studied over a range of concentrations, and values quoted correspond to relative apparent V,, values except in the case of NADH. Cleere and Conghlan (169). Measurements were a t pH 7.8 with ferricyanide (1 mM) as oxidizing substrate, with reducing substrates at 0.07 mM. i The substrate concentration used was higher than that for other substrates in this column. i Booth (167). Measurements were made a t pH 7.2 with methylene blue (0.1 mM) as oxidizing substrate, with reducing substrates at optimum concentrations. Gregory et nl. (170). This value represents the apparent K,,, for the nonhydrated form of acetaldehyde, which is the true substrate (171). Measurements were made aerobically a t pH 7.0, with cytochrome c &s oxidizing substrate. Z K ; value as a competitive inhibitor of N*-methylnicotinamide oxidation. The substrate concentration used was lower than that for other substrates in this column. Valerino and McCormack (172). Measurements were made at p H 7.4 with oxygen (approximately 0.25 mM) as oxidizing substrate, with the reducing substrate at 0.05 mM.

$

a

B

0

W

%

350

R.

C. BRAY

TABLE VII SEQUENCE OF POSITIONS I N WHICHPURINE Is OXIDIZED BY XANTHINE OXIDASEA N D RELATED ENZYMES Enzyme

Oxidation sequence

Ref.

Milk xanthine oxidase Human xanthine oxidase Turkey xanthine dehydrogenase c. ey~indrosporumxanthine dehydrogenase V. alcaleseensa xanthine dehydrogenase Rabbit aldehyde oxidase

6 --t 2,6 + 2,6,8 6 2,6 + 2,6,8 6 + 2,6 + 2,6,8 8 + 6,8 + 2,6,8

173 173 174 168

8

176

8

10

~

~~

See footnote (176).

cially marked for purine. Mostly, though not invariably, K,, values are higher for aldehyde oxidase than for xanthine oxidase. There are also, sometimes, important diff erences among the enzymes in the position of the substrate molecule in which the enzyme first carries out its attack, where several different sites of hydroxylation are possible. Data relating to purine are summarized in Table VII (1U,168,173-176). The first three enzymes in Table VII (including milk xanthine oxidase) attack this substrate first in the 6 position, giving hypoxanthine, with well-defined sequential conversion of this primary product to xanthine, then to uric acid. There are no indications of alternative oxidation routes, e.g., via 2,s-dihydroxypurine (173). Since it is well established (173,174; cf. 177) that at intermediate times during the reaction, hypoxanthine and xanthine accumulate in substantial amounts in solution, it follows that purine oxidation must normally take place as three successive twoelectron steps with dissociation of the product from the enzyme after each step. Indeed, it is hardly surprising that this should be the case, since it is reasonable to expect that the purine nucleus would have to be oriented in the active site in three different and specific manners in order to allow oxidation to take place in the 6, 2, and 8 positions, respectively (cf. 178). Presumably the most effective way for the enzyme to 173. F. Bergmann and S. Dikstein, JBC 233, 765 (1956). 174. W. F. Cleere, J. F. Mulhern. and M. P. Coughlan, Comp. Biochem. Physiol. 50B, 323 (1975). 175. S. T. Smith, K. V. Rajagopalan, and P . Handler, JBC 242, 4108 (1967). 176. This organism has also been known as Micrococcus lactilyticua (see footnote

144) * 177. M. M. Jeiewska, Eur. J . Biochem. 36, 385 (1973). 178. P.-S. Song and T. A. Moore, Znt. J . Quantum Chem. 1, 699 (1967).

6.

MOLYBDENUM HYDROXYLASES

35 1

reorient the molecule would be by releasing it and then rebinding. The second group of enzymes in Table V I I attacks purine first in the 8 position instead of in the 6 position. Thus, presumably, in the two sets of enzymes the purine molecule itself must be oriented differently when it is bound in the active sites, reflecting basic structural differences between the sites. An interesting corollary to these findings is the observation (179) that aldehyde oxidase oxidizes N1-methylnicotinamide simultaneously to two products, the 2- and 4-pyridones. These are produced in fixed relative proportions, this proportion, however, varying from one source of the enzyme to another. Presumably this means that a single substrate can be capable of being bound in the active site of a given enzyme in mare than one orientation. Some attempts have been made to consider the specificity of xanthine oxidase theoretically via molecular orbital calculations. However, the differing specificities of the various enzymes make it quite clear that e.g., with purine (cf. Table VII), it cannot be only the structure of the substrate which determines the position of enzymic attack. Therefore, despite the obvious difficulties, molecular orbital calculations ought properly to be made not on isolated substrate molecules but on enzyme-substrate complexes (166,178). With such a wealth of specificity data available one might have hoped to be able to draw conclusions on the nature and geometrical arrangements of substrate binding groups in the active centers of the four enzymes, but even speculations along these lines, let alone firm conclusions, seem to be lacking from the literature at the present time. However, one interesting point on the specificity which emerged from pH-variation studies, must be mentioned : N1-Methylnicotinamide is oxidized by the milk and turkey enzymes, but only a t very high pH values where activity toward xanthine is low (77,169).It was preposed (77), from variations of K , values with pH, that a group in the active center has a pK of 10.7 and that when this is protonsted, purine binding is facilitated, whereas when it is not protonated, binding of the positively charged quaternary compound is favored instead. An interesting corollary is that the enzyme from V . alcalescens (176) does not oxidize N1-methylnicotinamide a t any pH value (175), clearly indicating a difference in the binding site of the enzyme from this source. The wide specificity ranges of these enzymes (Table VI) lead to problems of nomenclature. The fastest reaction catalyzed by milk xanthine oxidase seems to be oxidation of 4-hydroxy-7-azapteridine ( 16 6 ) . Furthermore, there is considerable doubt concerning the biological role of 179. R. C. Felsted, A. E.-Y. Chu, and S. Chaykin, JBC 248, 2580 (1973).

352

R. C. BR.4Y

the enzyme (Section 1,B). Nevertheless, we prefer to maintain the traditional name and shall continue to call it xanthine oxidase. Similarly, we will continue to call aldehyde oxidase by this name, although there is no reason to believe that the enzyme has any biological role in the oxidation of these substrates (10) and although, for example, aldehyde oxidase oxidizes 3-methylhypoxanthine nine times faster than it oxidizes acetaldehyde (Table VI). The use of the traditional names has quite probably hindered research, e.g., in work on molybdenum hydroxylases in Drosophda melanugaster (Section IV,D), by giving workers unfamiliar with the field preconceived ideas about specificity. Nevertheless, renaming enzymes to molybdenum hydroxylase I, molybdenum hydroxylase 11, etc., would only lead to confusion. However, it is suggested that all new enzymes be called molydenum hydroxylases, giving an indication of their specificity, e.g., “an NAD+-specific molybdenum hydroxylase with a reducing substrate specificity rather like that of rabbit aldehyde oxidase,” etc. Oxidation of NADH by xanthine oxidase is quite different in many respects from the oxidation of other substrates. It is unique among reactions catalyzed by the enzyme in that it is formally a simple oxidation, not a hydroxylation, the product being NAD’. Furthermore, in contrast to other substrates, which interact with the enzyme via the molybdenum site (evidence on this point is summarized in Section II,C,2,a), NADH clearly interacts via flavin. Evidence for this comes from complete loss of activity toward KADH in the deflavo enzyme ( 6 3 ) , in contrast to retention of this activity on conversion to the desulfo enzyme by treatment with cyanide (Is&) or in the presence of various inhibitors ( 4 ) . Lastly, reaction of the enzyme with two common reducing agents which are, however, quite properly considered as reducing substrates has to be considered. These are dithionite and borohydride. Dithionite has frequently been used to prepare reduced xanthine oxidase samples, but no detailed kinetic work has been reported on the reduction process (see, however, 41,180,181). Further, enzyme turnover using dithionite as reducing substrate seems well established (182;see also Section II,C,l,e) . Precise rates of reduction of active and of desulfo enzyme by dithionite, which are possibly quite similar to one another, have not been measured, and the mechanism of the reaction is unknown. On the other hand, borohydride has been shown to reduce active xanthine oxidase much faster than it does the desulfo form (44).This has been taken as evidence that 180. G. Stahrer and G. B. Brown, JBC 244,2498 (1969). 181. H. Gutfreund and J. M. Sturtevant, BJ 73, 1 (1959). 182. T-C. Lee, G. Stohrer, M. N. Teller, A. Myles, and G . B. Brown, Biochemistry 10, 4463 (1971).

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hydride transfer from reducing substrate to enzyme is involved in the normal catalytic reaction of active enzyme (see Section II,C,2b). b. Specificity: Oxidizing Substrates. It has long been known that the specificity of milk xanthine oxidase toward oxidizing substrates (electron acceptors) is, like that to the reducing substrates, low (f,4). Apart from oxygen and dyes, such as methylene blue and 2,6-dichlorophenolindophenol, ferricyanide as well as quinones and a large number of other comp o h d s , particularly nitrogen-containing ones, are all reduced. Reduction of NAD' by the enzyme has already been considered in Section II,B,3,d. In the case of the avian enzymes, it has been shown (75b,78)that conversion to the deflavo form abolishes activity toward NAD' as acceptor. Thus, flavin is presumed to be the binding site for NAD' as it is for NADH. It seems that no systematic and quantitative comparative studies of the specificity of the native enzyme toward oxidizing substrates have been carried out using modern methods. However, such work would probably prove comparatively uninteresting since it is likely that for many of the commonly used acceptors, reduction rather than reoxidation of the enzyme must, under most conditions, be rate-limiting in the catalytic process. That this is the case, has definitely been shown when oxygen is acceptor (45,f 81;see Section II,Cl2,e). In the absence of comparative work, only a few particular points can be noted here about individual reactions and some K , values quoted. (Data on other molybdenum hydroxylases are summarized in Section III,C,2). For reaction with xanthine, Fridovich (f82a) found K,,, for oxygen to be 8 X M a t pH 10.0, with saturating concentrations of the reducing substrate. Similarly, at pH 8.5, Massey et al. (35) obtained a value of 5X M . The mechanism of reaction of the enzyme with oxygen will be considered in Section II,C,2,g. The deflavo enzyme (Section II,B,3,b) is devoid of oxidase activity ( 6 3 ) . Interaction of xanthine oxidase with 2,6-dichlorophenolindophenolhas been the subject of considerable study. The reaction is somewhat unusual in that it is markedly inhibited by excess of the oxidizing substrate. Fridovich (183) found K , for the reaction to be about 1.5 X lo-? M , with Ki 1 X M . The inhibition was found to decrease in the presence of organic solvents such as dimethoxyethane, possibly because of conformational changes in the enzyme. Gurtoo and Johns (f84) detected a complex spectrophotometrically between oxidized enzyme and M . Formation of indophenol with a dissociation constant of about 182a. I. Fridovich, JBC 239, 3519 (1964). 183. I. Fridovich, JBC 241, 3624 (1966). 184. H. L. Gurtoo and D. G. Johns, JBC 246, 286 (1971).

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this complex presumably accounts for the inhibition. From studies with thiol reagents, with alloxanthine and with the deflavo enzyme, these workers concluded that the site of inhibitory binding of indophenol must be either molybdenum of the enzyme or a thiol group near to molybdenum. I n agreement with this, conversion of aldehyde oxidase to the deflavo form did not diminish activity toward indophenol (77c). With regard to reduction of cytochrome c by the xanthine oxidase system, it is now known that aerobically this reaction proceeds primarily, if not wholly, not from direct interaction between the enzyme and the cytochrome, but via enzymic reduction of oxygen to superoxide. This free radical, then, in turn, reduces cytochrome c nonenzymically (see Section II,C,lc). There is some controversy as to whether the intact enzyme is capable of reducing cytochrome c anaerobically in the presence of xanthine. Kanda et al. (78; see also 186) found no activity in this assay, although Massey et al. (63) found some 10% of the activity of the aerobic system. On the other hand, somewhat unexpectedly, with the deflavo enzyme, cytochrome c definitely is reduced anaerobically a t quite substantial rates (63,78).Further, as yet unexplained, changes in acceptor specificity in the deflavo enzyme are enhanced reactivity toward ferricyanide and nitro blue tetrazolium (76,78), observed particularly in the case of the deflavo enzyme of chicken liver (see also 77c). Apart from cytochrome c, reaction of two other one-electron acceptors, ferricyanide (4,186) and phenazine methosulfate ( 4 ) , with the native enzyme has to be considered. Reaction with ferricyanide does not seem to have been studied in much detail, although high rates are obtained in this assay (10). Somewhat surprisingly, the reaction is apparently stimulated by oxygen when xanthine is reducing substrate ( 6 3 ) , although to some extent, this may be related to nonenzymic reaction between uric acid and ferricyanide. An interesting feature of reactions both with ferricyanide (10) and with phenazine methosulfate (187) is that these are effective acceptors for oxidation of allopurinol which is a potent inhibitor in most assay systems (Section II,C,l,g). This is attributed (61) to rapid reoxidation by these acceptors of a complex of alloxanthine with xanthine oxidase molecules in which molybdenum is in the four-valent state. This fast reoxidation contrasts with very slow reoxidation of the complex by oxygen. Brown and co-workers have shown that a range of purine N-oxides (some of which are of interest because of their carcinogenic properties) may serve as relatively slow oxidizing substrates for xanthine oxidase, being converted in the process to the parent compounds, without the oxide 185. J. M. McCord and I. Fridovich, JBC 244, 6049 (1969). 186. M. Nakamura and I. Yamaaaki, BBA 327, 247 (1973). 187. D. G. Johns, BBRC 31, 197 (1968).

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function (180,182,188). Another N-oxide which can be reduced by xanthine oxidase is nicotinamide N-oxide (189).This latter reaction is of particular interest, since Murray et al. (190) have shown that with xanthine as reducing substrate and nicotinamide N-oxide as oxidizing substrate, **O is transferred directly from the N-oxide group to the uric acid which is produced. The reaction, therefore, contrasts with the more normal reaction of xanthine oxidase, when using oxygen as oxidizing substrate, in which the oxygen, incorporated into uric acid, can be shown to be derived from water (190-192). c. One- and Two-Electron Reduction of Oxidizing Substrates and Secondary Reactions Following o n One-Electron Reduction. The first suggestion that xanthine oxidase might be capable of reducing oxygen in a oneelectron reaction of the superoxide free radical, 02-, seems to have been made by Fridovich and Handler in 1958 (193,194). It had previously generally been assumed that oxygen reduction to hydrogen peroxide took place via a two-electron mechanism. Subsequent work in several laboratories fully confirmed production of superoxide by the enzyme system, the most direct demonstration of formation of this radical in free solution being provided by EPR work (196-198). It is now clear that the enzyme is a somewhat unusual one in using the one- and two-electron mechanisms of oxygen reduction simultaneously (46,199,600). I n this respect it is, however, analogous to free flavins (201). For xanthine oxidase, high pH, 188. Note that further reduction beyond this level can also take place in some cases, see Section II,C,l,e. 189. K. N. Murray and S. Chaykin, JBC 241, 3468 (1966). 190. K. N. Murray, J. G. Watson, and S. J. Chaykin, JBC 241, 4798 (1966). 191. H. S. Mason, Science 125, 1185 (1957). 192. The fact that xanthine oxidase carries out some reactions by direct oxygen transfer and others by simple electron transfer is a further example of the extraordinary versatility of the enzyme. The mechanism of the direct oxygen transfer reactions will not be considered further in this chapter. It must be noted, however, that at pH 8.9 (although not at pH 7.5) a small but apparently significant amount of direct oxygen transfer from molecular oxygen has been reported to take place during the ordinary xanthine oxidase reaction (190). 193. I. Fridovich and P. Handler, JBC 233, 1581 (1958). 194. In much of the earlier work by this group (e.g., 195) it was assumed that 0,- remained bound to the enzyme rather than being liberated into solution as a free entity. 195. L. Greenlee, I. Fridovich, and P. Handler, Biochemistry I, 779 (1962). 196. R. C. Bray and P. F. Knowles, Proc. Roy. Soc., Ser. A 302, 351 (1968). 197. P. F. Knowles, J. F. Gibson, F. M. Pick, and R. C. Bray, BJ 111, 53 (1969). 198. R. C. Bray, F. M. Pick, and D. Samuel, Eur. J. Biochem. 15, 352 (1970). 199. S. Nakamura and I. Yamazaki, B B A 189,29 (1969). 200. I. Fridovich, JBC 245, 4053 (1970). 201. V. Massey, G . Palmer, and D. Ballou, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), Vol. 1, p. 25. Wiley, New York, 1973.

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high oxygen concentrations, and low xanthine concentration all tend to favor the one-electron pathway relative to the two-electron pathway. It is interesting that reduction of quinones by the enzyme has also been shown by Nakamura and Yamazaki (286,199) to proceed like that of oxygen, both by one- and two-electron routes. There has been some disagreement in the literature regarding the site of interaction of oxygen with the enzyme when one-electron reduction takes place. The deflavo enzyme, which is of course devoid of oxidase activity ( 6 3 ) ,not surprisingly failed to produce superoxide (202).Therefore, Massey and co-workers have very reasonably assumed (1) that superoxide is produced a t the flavin site in the enzyme. On the other hand, Misra and Fridovich (203), on the basis of comparisons of superoxide production by xanthine oxidase, FMN, flavodoxin, and ferredoxins, suggested that the Fe/S centers of the enzyme might be involved in producing the oxygen radical. However, the latest work on the enzyme from Massey’s group (46,4666; see also Section 11,C12,i) seems fully consistent with 02-production at the flavin site and indeed goes a considerable way toward explaining why xanthine oxidase is different from free flavins in its reactions with oxygen. A detailed discussion of the properties of the superoxide radical is outside the scope of this chapter. However, physicochemical studies on it (204-206) have continued in parallel with biochemical work. The redox potential for the system OJ0,- at neutral pH is now known to be -0.33 V (20.4) which is not inconsistent with one-electron reduction of oxygen by flavin of the enzyme (207). Though unstable, the superoxide radical has a finite lifetime in aqueous solution a t room temperature and neutral pH values. It decays in the absence of trace metal impurities, by dispro202. W. H. OrmeJohnson and H. Beinert, BBRC 38, 905 (1969). 203. H. P. Misra and I. Fridovich, JBC 247, 188 (1972). 204. P. M. Wood, FEBS (Fed. Eur. Biochem. Soc.) Lett. 44, 22 (1974). 205. J. Rabani and S. 0. Nielsen, J. Phys. Chem. 73,3736 (1969). 206. D. Behar, G. Czapski, J. Rabani, L. M. Dorfman, and H. A. Schwarz, J . Phys. Chem. 74, 3209 (1970). 207. The new value for the potential, although higher than earlier estimates, still

appears significantly lower than the potentials of free flavins in neutral solution (808). This means that it remains difficult (cf. 808,) to understand, from tt thermodynamic point of view, one-electron reduction of oxygen by free reduced flavins a t pH 6.8 (80.9). On the other hand, the redox potential of the flavin of xanthine oxidase, at pH 8.5, although not known, may well be lower than that of the O,/O,- systems (cf. 808b). 208. H. J. Lowe and W. M. Clark, JBC 221, 983 (1956). 208a. V. Massey, G. Palmer, and D. Ballou, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 349. Univ. Park Press, Baltimore, Maryland, 1971. 208b. S. G. Mayhew, G. P. Foust, and V. Mamey, JBC 244,803 (1969).

6.

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357

portionation to oxygen and hydrogen peroxide, with a second-order rate constant of about lo6 M-I sec-1 at p H 7 (206). The superoxide radical, apart from disproportionation, is a fairly reactive species. Therefore, superoxide generated by the xanthine oxidase system can enter into a variety of secondary reactions; some of these are reductions and some oxidations (209).The most noteworthy secondary reaction is the reduction of cytochrome F by superoxide (185; see also 2 U ) . Discovery of this reaction has gone a long way to resolving earlier reported anomalies and confusion ( 4 ) relating to cytochrome c reduction by the xanthine oxidase system. Other substrates which react rapidly with enzymically generated superoxide (185) are tetranitromethane, which reacts in a very fast second-order reaction with k = 1.9 X lo9 M-' sec-l (212), and epinephrine, which is oxidized to adrenochrome. Nitro blue tetrazolium is also oxidized by superoxide, although it is capable of reacting directly with the reduced enzyme also ( 2 1 3 ) . Doubtless, superoxide can react directly with many other compounds of biological interest and doubtless some further properties of the xanthine oxidase system will in due course be shown to depend on it. However, the essentially diffusion-controlled decomposition (214) of superoxide by bovine superoxide dismutase (185,215) must particularly be mentioned. Competition for superoxide generated by the xanthine oxidase system, between superoxide dismutase and either cytochrome c, or tetranitromethane, or nitro blue tetrozolium, has provided the basis for assays of superoxide dismutase activity (215). A further stage in complexity of the secondary reactions is possible if superoxide radicals, generated by xanthine oxidase, then react with other constituents of the solution to produce new unstable free radical species, which, in turn, further react to produce the products ultimately detected. A case in point is the generation of ethylene from methional (P-methylthiopropionaldehyde) by the xanthine oxidase system. It was proposed by Beauchamp and Fridovich (216) that this reaction, which is inhibited by superoxide dismutase and by catalase, depends on reaction 209. A. different type of secondary reaction, which apparently depends on HZOZ is the coupled oxidation of alcohols in the presence of catalase, rather than 02-, acting as a peroxidase (D. Keilin and E. F. Hartree, BJ 39, 293 (1945); see also Hart et al. (210)). 210. L. I. Hart, E. C. Owen, and R. Proudfoot, Brit. J. Nutr. 21, 617 (1967). 211. J. Rabani and G. Stein, Radiat. Res. 17,327 (1962). 212. J. Rabani, W. A. Mulac, and M. S. Matheson, I . Phys. Chem. 69, 53 (1965). 213. C. Beauchamp and I. Fridovich, ABB 44, 276 (1971). 214. E. M. Fielden, P. B. Roberts, R. C. Bray, D. J. Lowe, G. N. Maiitner, G. Rotilio, and L. Calabrese, BJ 139, 49 (1974). 215. I. Fridovich, Accounts Chem. Res. 5, 321 (1972). 216. C. Beauchamp and I. Fridovich, J. B i d Chem. 245, 4641 (1970).

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R.

C.

BRAY

between superoxide and hydrogen peroxide, yielding the very reactive hydroxyl radical 02-

+ HzOz + OH’ + OH- + 02

(4)

The hydroxyl radical was then supposed to be responsible for the observed reaction with methional. Although other secondary reactions (e.g., 216a) brought about by the xanthine oxidase system have also been suggested to occur via generation of the hydroxyl radical in this way, there is no direct evidence implicating the radical (217) and other mechanisms might in fact be involved. It may be appropriate to mention luminescent reactions of the xanthine oxidase system a t this point, since free radicals are presumably involved in them. Although it has been known (4,221) for some while that luminescense is observed during substrate oxidation by xanthine oxidase in the presence of oxygen together with luminol, nevertheless, it is apparently uncertain (222) whether or not 0,- is involved in this reaction. I n contrast to such studies, a much weaker, but nevertheless quite definite, luminescence may be detected during functioning of the enzyme in the absence of any added chemiluminesrent compounds ( 2 2 3 ) . I n this case it has been proposed (224) that Eq. (4) may be involved in producing species involved in the luminescent reaction, but this is not certain (see also 217). d. Assays for Xanthine Oxidase and Dehydrogenase Activities (Including Other Molybdenum Hydroxylases) . The assay of xanthine oxidase or dehydrogenase activity in general presents no unusual problems. Purified enzyme preparations are normally assayed by following substrate disappearance or product appearance, spectrophotometrically (2.25). Probably the most widely used assay is the one with xanthine 216a. K.-L. Fong, P. B. McCay, J. L. Poyer, B. B. Keele, and H. Misra, JBC 248, 7792 (1973). 217. The rate of the reaction of Eq. (4) is apparently low, if indeed the reaction occurs

a t all (2f8,2f3). Furthermore, any O H produced would decay quite rapidly by reaction with H202 since reaction between these species has a rate constant of about 2 X 10’ M-1 sec-1 (820). 218. P. George, Discuss. Faraday SOC.2, 196 (1947). 219. W. C. Barb, J. H. Baxendale, P. George, and K. H. Hargrave, T r a m . Faraday Soc. 47, 462 (1951). 220. L. M. Dorfman and G. E. Adams, in “National Standard Reference Data System,” vol. 46, p. 35. Nat. Bur. Stand., Washington, D. C., 1973. 221. J. R. Totter, E. C. de Dugros, and C. Riviero, JBC 235, 1839 (1960). 222. G. M. Oyamburo, C. E. Prego, E. Prodamov, and H. Sote, BBA 205, 190 (1970). 223. J. Stauff, H. Schmidkunz, and G. Hartmann, Nature (London) 198,281 (1963). 224. R. M. Arneson, ABB 136, 352 (1970). 225. Fluorimetric methods continue, owing to their high sensitivity, to be particularly useful in special cases, e.g., with pteridine substrates (226).

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359

as reducing substrate and oxygen as oxidizing substrate (see, e.g., 21,36). Assay systems utilizing xanthine are of course in principle simpler than those employing hypoxanthine, since the latter undergoes a two-step oxidation, first to xanthine, then to uric acid. If activity toward a series of reducing substrates of varying spectral properties is to be compared, it is generally convenient to follow the reactions via consumption of the oxidizing substrate. To this end Krenitsky (10) used conversion of ferricyanide to ferrocyanide, followed spectrophotometrically a t 420 nm. No doubt an oxygen electrode could be used effectively for a similar purpose if oxygen were preferred as acceptor. Of assays of xanthine oxidase or dehydrogenase activities which are suitable for studies on crude tissue extracts or similar materials, only a few will be briefly mentioned here. Thus, traditional manometric methods have continued to be useful (e.g., 210), while methods based on the use of radioactive substrates are capable of high sensitivity (e.g., 227,228). Spectrophotometric methods (229,230) based on the reduction of tetrazolium salts are, however, particularly convenient. It may be pertinent to add a word of warning a t this point to those interested in the practical aspects of assaying molybdenum hydroxylases in crude materials. As has been discussed in the previous subsection, many commonly used xanthine dehydrogenase assays (for example, those depending on reduction of cytochrome c or nitro blue tetrazolium) depend, a t least for some of the enzymes either wholly or in part, on nonenzymic reduction of the ultimate acceptor by the superoxide ion, which is in turn generated by the enzymic reduction of oxygen. Therefore, unless it is known that the enzyme under study is entirely without oxidase activity, or alternatively, unless the assay procedure is being carried out under rigorously anaerobic conditions, then the possibility must always be borne in mind that observed apparent changes in xanthine dehydrogenase activities are, in reality, due simply to changes in the lifetime of the superoxide ion in the assay medium. This could be due, for example, to changes in superoxide dismutase activity levels, or even, simply to changes in the level of contamination by trace metal impurities. Where mediation of the assay by superoxide ions is suspected, this may readily be tested for by addition of superoxide dismutase t o the system ( 2 1 5 ) . I n general it would seem wise to avoid the use of superoxide-mediated 226. D. G. Priest and J. R. Fisher, ABB 146, 391 (1971). 227. J. E. Ultmann, P. Feigelson, and S. Harris, Anal. Biochem. 1, 417 (1960). 228. I. Oliver, 0. Sperling, U. I. Liberman, M . Frank, and A. de Vries, Biochem. Med. 5, 279 (1971). 229. R. Fried, ABB 16, 427 (1966). 230. D. J. Bauer and P. L. Bradley, Brit. J. E s p . Pathol. 37, 447 (1956).

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R. C. BRAY

assays for future work on these enzymes, where this is possible, or, a t the very least, to routinely add EDTA to such systems in order to control and minimize trace metal-catalyzed decomposition of the radical. e. Reversibility of the Catalytic Reaction (Including Other Molybdenum Hydroxylases) . Interest in the possibility that purine hydroxylation reactions catalyzed by xanthine oxidase or related enzymes might be reversible is of long standing ( 4 ) . In fact, such reversibility has now been demonstrated for many of the molybdenum hydroxylases, although in most cases these reverse reactions are quite slow. Reversibility studies have been of two types, as depicted in Eqs. ( 5 ) and (6) :

+

(5) (6) One obvious experimental point must be mentioned in view of the lowness of the activities which are involved. I n general, the most unequivocal evidence for existence of dismutation reactions of the type shown in Eq. ( 5 ) will be provided by appearance of the reduced product species, in this case, hypoxanthine. Where disappearance of xanthine or appearance of uric acid is relied on to demonstrate such a reaction for an enzyme which has even a low oxidase activity, then the onus will be on the experimenter to show he really did work under truly anaerobic conditions. The reverse reactions may be particularly important, from a metabolic point of view, in the case of the bacterial xanthine dehydrogenases. For the enzyme from Clostridium cylindrosporum, relatively slow reduction reactions [including the one in Eq. (6) ] , as well as a number of dismutation reactions, were unambiguously demonstrated by Bradshaw and Baker (168). For the V . alcalescens enzyme (176), reverse reactions appear to be somewhat faster. Smith et al. (176) stated that the rate of the xanthine dismutation reaction is 60% of the corresponding rate of aerobic xanthine oxidation. Since the latter reaction occurs (176) a t about 12% of the rate of the fastest forward reaction of the enzyme (which is the one with ferredoxin as acceptor), the dismutation reaction would thus appear to be occurring at about 7% of the maximum forward rate (231). For the avian enzymes, the dismutation reaction has been reported not to occur in the case of the chicken liver enzyme (232). However, with the turkey enzyme, Cleere and Coughlan (169) found a slow but definite dismutation reaction, with a rate of 2.8% of the oxidase rate, i.e., about 2 Xanthine hypoxanthine uric acid Dithionite uric acid + xanthine

+

231. It might be desirable to redetermine this value since uric acid formation only was monitored (cf. ,932); nevertheless, qualitatively at least, the work (176) leaves no room for doubt that the reverse type of reactions do occur readily with this enzyme.

6.

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361

0.05% of the highest dehydrogenase rate which they observed. Finally, for the bovine milk enzyme the backward reactions are also slow although definite reduction, e.g., by dithionite of uric acid to xanthine and to hypoxanthine, has been reported (180,182,633). Existence of the dismutation reaction does not appear to be in any way related to the phenomenon of substrate “activiation” which is observed with some of these enzymes, as manifested by downward curvature of the 1/V against l/S pIots a t high substrate concentrations. Although Smith et al. (17’5) attempted to relate the two phenomena, there seem in fact to be many examples of downward curvature, with particular substrates and particular enzymes, which are not correlated in any way with fast dismutation reactions (168,169,686,Md). f. Turnover Numbers, Activation Energies, and p H Effects. The turnover number of milk xanthine oxidase may be readily calculated from the data in Tables I and I1 for the conditions of the standard assay with xanthine and oxygen (i.e., pH 8.2: 23.5O; 0.1 mM xanthine: solution saturated with air). The value obtained from the calculated Activity/ E,,, ratio of Table I corresponding to 100% pure active enzyme is 12.0 moles of xanthine per active center per second, assuming two centers per mole. Extrapolation to infinite xanthine and infinite oxygen concentration may be made from data of Massey et al. (35), which indicates a 1.35-fold increase in activity above that in the standard assay. The extrapolated limiting turnover number therefore becomes 16.2 sec-l a t pH 8.2 and 23.5O. Massey et al. (35) reported an activation energy, determined under the standard assay conditions as given above and measured in the temperature range 10°-380, of 14,500 cal/mole. Earlier work by Bray et a2. (26) indicated that the activation energy fell off slightly above 20° (234). From their data, values of 16,000 cal/mole in the range 2Oo-25O and of 17,800 between 50 and 20° can be calculated. For the turkey enzyme, in assays with NAD’ as acceptor, an activation energy of 13,375 cal/mole has been reported (169). Detailed data on maximum turnover numbers and activation energies do not appear to be available for other assay systems (see, however, Table VI for approximate relative rate data for other substrates). Effects of pH on the reactions of milk xanthine oxidase are, not unexpectedly, complicated. Although the pH optimum for xanthine oxidation 232. D. G. Priest and J. R. Fisher, Eur. J. Biochem. 10,439 (1969). 233. G. Stohref, Isr. J . Chem. 6, 845 (1968). 234. Ackerman and Brill @S6) also reported that the apparent activation energy

falls with increasing temperature. 235. E. Ackerman and A. S. Brill, BBA 56,397 (1962).

362

R. C. BRAY

by molecular oxygen is frequently quoted ( 4 ) as being a t about 8.3, the exact value depends very much on the precise experimental conditions. This is partly because inhibition by excess substrate (Section II1C,2,j) is markedly pH-dependent (236).Oxidation of xanthine appears to be regulated by a t least three pK values. These are those of xanthine, a t pH 7.5 and 11.8 (237,238) and that of a group in the enzyme with a pK of 10.7 (77; see also Section IIICll,a),This substrate apparently has to bear a single negative charge in order to react, while the group in the active site has to be positively charged. For xanthine oxidation V,,, is constant from pH 11.0 down to 8.5 (46,77) and has decreased only by a factor of 2 on lowering the pH to 7.0 and by a further factor of 2 at pH 5.0 (77). g. Inhibitors. The distinction between an enzyme inhibitor and a reagent which modifies the enzyme chemically is frequently not a very rigid one. I n the case of xanthine oxidase, we have treated the important reagents, cyanide, formaldehyde, methanol, and arsenite, as chemical modifiers of the enzyme and have discussed them in Sections II,B,3,e and II,B,3,f. Earlier work on inhibitors has been reviewed previously ( 4 ) . As would be expected many substrates and product analogs are inhibitors, usually competitive ones, of xanthine oxidase and a number of these have been useful in steady-state kinetic analysis of the enzyme mechanism. Urea and guanidine (182a) as well as salicylate (35) have been employed for this purpose. Synergism between guanidine and thiocyanate in inhibiting the enzyme has been reported by Fridovich (237). Apparently guanidine interacts at the xanthine binding site, although where the thiocyanate interacts seems less clear. Another substrate analog which is a fairly potent competitive inhibitor of xanthine oxidase is ammeline (2,4diamino-6-hydroxy-s-triazine) . This is frequently present as an impurity in guanidine (239).An interesting point is that ammeline inhibits oxidation not only of xanthine but also of N1-methylnicotinamide, in the case of the milk enzyme. On the other hand, with aldehyde oxidase (for which xanthine is not a substrate, Table VI), oxidation of the nicotinamide derivative is not inhibited. This again emphasizes that there must be important differences between the substrate binding sites of these two enzymes. Three particularly potent inhibitors of xanthine oxidase are 230. I. Fridovich and P. Handler, JBC 233, 1581 (1958). 237. I. Fridovich, ABB 109, 511 (1965). 238. J. J. Christensen, J. H. Rytting, and R. M. Izatt, Biochemistry 9, 4907 (1970). 239. I. Fridovich, Biochemistry 4, 1098 (1965). 240. 0.H. Lowry, 0. A . Bessey, and E. J. Crawford, JBC 180, 399 (1949).

6.

MOLYBDENUM HYDROXYLASES

363

2-amino-Qhydroxypteridine-6-aldehyde(4,240), purine-6-aldehyle (941) , and allopurinol. The last named of these will be considered in some detail. During the past decade studies of inhibition of milk xanthine oxidase by allopurinol [ 4-hydroxypyrazolo (3,4-d) -pyrimidine, see Table V I for formula] and related compounds have thrown much light on the mechanism of action of the enzyme. Allopurinol is both a substrate and a n inhibitor of xanthine oxidase (242) and by virtue of the latter property has found clinical application as an anti-gout drug (19). Its potency as a xanthine oxidase inhibitor is extremely high, Spector and Johns (243) reporting an apparent dissociation constant for the enzyme inhibitor complex of 6 X M . Nevertheless, as will be discussed below, under appropriate conditions, the enzyme is capable of continuously converting allopurinol to .alloxanthine [4,6-dihydroxypyrazolo (3,4-d) -pyrimidine, also known as oxipurinol]. As first noted by Johns (187), when xanthine is reducing substrate, allopurinol inhibits oxidation when oxygen is the electron acceptor, b u t it shows relatively little effect when phenazine methosulfate is employed. Later work by Massey and co-workers (61) showed, however, that the obvious interpretation (187), namely, that allopurinol interacts a t the oxygen binding site of the enzyme, is in fact entirely wrong. Contrary to earlier work (a&), it was found (61) that anaerobically, allopurinol reduces xanthine oxidase (see Section II,C,2,f for a discussion of reduction of the enzyme and of the various phases in which it occurs). Reduced enzyme prepared with allopurinol shows normal Fe/S and FADH. EPR signals (61) and generally gives little, if any, Mo(V) signal (245). I n contrast to reduction with xanthine, there is however essentially no slow phase in the reduction process when using allopurinol (61,110; see Section II,C,2,k). Further, it was found (61) that enzyme reduced with allopurinol is readily reoxidized b y oxygen. However, the visible spectrum of the reoxidation product differs slightly but importantly from th a t of normal oxidized enzyme. Flavin and the Fe/S centers apparently make their normal contributions to the spectrum, but there is some increased absorption a t 380 nm and a slightly decreased absorption a t 440 nm. Furthermore, enzyme so treated is inactive in the normal assay system with xanthine and oxygen as substrates. I n this work, identical results were obtained if treatment with allopurinol was replaced b y treatment with alloxanthine plus xanthine. 241. D. A . Gilbert, BJ 93, 214 (1964). 242. G. B. Elion, Ann. Rheum. Dis. 25, 608 (1966). 243. T. Spector and D. G. Johns, JBC 245, 5079 (1970). 244. T. Spector and D. G. Johns, BBRC 32, 1039 (1968). 245. I n rapid freezing experiments allopurinol does give the Rapid signal, but this quickly disappears and is later replaced by some Slow signal (116).

364

R.

C.

BRAY

Massey et al. (61) interpreted these findings in terms of what they termed a “suicide” reaction, in which allopurinol was converted to alloxanthine in the enzyme-active center but then remained firmly bound there, with the molybdenum locked in the four-valent state, while flavin and the Fe/S centers were reoxidized normally. I n keeping with this interpretation, they found that enzyme inactivated by allopurinol could be reactivated and its spectrum restored to the normal one by reoxidation with phenazine methosulfate or ferricyanide. Although oxygen was also capable of carrying out this reoxidation, the reaction was an extremely slow one with it, and, incidentally, had an unusually high temperature coefficient. The conclusion that molybdenum is in the four-valent state in the inhibited complex was derived from studies on the stoichiometry of the reoxidation by ferricyanide. Finally, both normal and deflavo xanthine oxidase were found to be active in the assay with allopurinol as substrate and phenazine methosulfate as electron acceptor (246). Apart from the above evidence, which appears to exclude flavin and Fe/S as sites for interaction with allopurinol and yielded data on the valency of molybdenum in the reduced enzyme, further important information also emerged from studies with allopurinol. It was found (61) from work on the stoichiometry of the inhibition that only active enzyme and not desulfo enzyme normally bound the inhibitor and, further, that in the active enzyme one mole of inhibitor was bound per molybdenum atom, This provided both a means of quantitating amounts of desulfo enzyme in xanthine oxidase samples and evidence for the dimeric nature of the enzyme, i.e., that it has two active sites per mole. Other disubstituted allopurine derivatives also bind to reduced xanthine oxidase in a manner comparable to that with alloxanthine. After reoxidation of flavin and Fe/S, the complex formed by each compound shows (61) a characteristic difference spectrum, relative to native enzyme, which is comparable to but by no means identical with that given by alloxanthine. A further most interesting point is that apparently (44), with appropriate reduction and after prolonged incubation, the desulfo enzyme as well as the.active form is capable of forming complexes with these allopurine derivatives. The complexes so formed have slightly different spectral properties from the corresponding ones with active enzyme. Thus, using 4,6-dimercapto-allopurine, the maximum in the difference spectrum is at 500 nm for active enzyme and a t 540 nm for the desulfo form. These results strengthen the conclusion that although the binding of the inhibitors is presumably directly to the molybdenum, nevertheless 246. The normal enzyme (and presumably the deflavo enzyme) is also active in

the ferricyanide assay system with allopurinol as substrate, see Table VI.

6.

MOLYBDENUM HYDROXYLASES

365

the persulfide sulfur must be very close since loss of it in the desulfo enzyme not only greatly changes the rate of reaction with the inhibitor but also significantly changes the spectrum of the product. Finally, it will be recalled (cf. Section II,B,l,b) that an allopurinol analog was used (44) in the resolution, by affinity chromatography, of active and inactive forms of xanthine oxidase. 2. Mechanism of Action

a. Sites at Which Substrates and Inhibitors Interact with the Enzyme. A central problem in studies of the mechanism of action of xanthine oxidase has been the question of the number and the nature of the sites which are available, in each enzyme half-molecule, for interaction of the various reducing and oxidizing substrates. T w o sites are by now well known and their roles, in a t least some and perhaps most of the various catalytic reactions of the enzyme, seem reasonably clear. These are the flavin site and the molybdenum site. As discussed in Section II,B,3,e the functioning of the molybdenum site is dependent on having a nearby persulfide group intact. Furthermore, the properties of the flavin site seem to depend on the presence or absence of a nearby thiol group (Section II,B,3,d). As will be discussed below, the possibility of additional sites in the enzyme molecule remains open for the time being, although there seems at present to be little, if any, firm evidence necessitating their existence. There is now much evidence, and indeed it seems generally accepted, that reducing substrates, except NADH, interact a t the molybdenum site of the enzyme, whereas oxygen, NAD+,and NADH interact a t the flavin site. Some of the most compelling pieces of evidence relating to the roles of these sites (which have, generally, been mentioned in earlier sections) are the following: 1. The demolybdo enzyme seems to be inactive toward xanthine. 2. Reducing substrates influence Mo (V) EPR signals from the enzyme in a number of specific and direct ways but have no such effects on signals from the other chromophores. 3. When the persulfide group is removed by cyanide treatment, substrates, except NADH, no longer reduce the enzyme. 4. Substrate analogs, particularly allopurinol, prevent reduction of the enzyme by normal substrates but not by NADH. 5. Removal of the flavin abolishes reoxidation of reduced enzyme by oxygen but does not affect its reducibility by, e.g., xanthine. There are many reports in the literature by workers who claimed to have achieved differential inhibition of some of the catalytic reactions

366

R. C. BRAY

of xanthine oxidase. No doubt, those inhibitors which are close structural analogs of reducing substrates interact with the enzyme via the molybdenum site. Let us, therefore, in the 1ight.of the above general comments, look a t some of the reported differential inhibition phenomena. We will not be concerned here with effects of cyanide, methanol, formaldehyde, and arsenite on the enzyme (Sections II,B,3,e and II,B,3,f). There seems to be good evidence that all of the reagents interact near but probably not actually a t the molybdenum site. Some of the other reported differential inhibition phenomena may be divided into three types. Compounds invoIved in the first group are not, in any real sense, xanthine oxidase inhibitors a t all. They do not combine with the enzyme but instead exert their actions by speeding up decay of the superoxide ion, this being produced in the reoxidation of reduced enzyme by oxygen (Section II,C,l,c). Such compounds therefore affect only those reactions, such as aerobic reduction of cytochrome c, which are mediated by the superoxide ion. “Inhibitors” in this category are myoglobin (247,248), carbonic anhydrase (248,249),“tetrazolium reducacid tase inhibitor” (16), and Tiron [ 1,2-dihydroxybenzene-3,5-disulfonic (250,251).It has been shown that effects observed with the last two compounds result from their reacting directly with superoxide, whereas, similarly, the myoglobin and the carbonic anhydrase preparations originally used were contaminated with the enzyme superoxide dismutase (185). Another group of compounds might interact via the thiol in the vicinity of the flavin of xanthine oxidase when this thiol is present in the molecule. Such inhibitors would be expected to affect only the D form of the enzyme (Section II,B,3,d). In this category, apparently, are the actione (87) of N-ethylmaleimide and tetraethylthiuram disulfide ( 4 ) on the rat liver enzyme. NADH competition with NAD’ (252) also seems to come into this category, although no doubt this inhibitor interacts primarily with flavin. Finally, it might be expected that inhibitors could be found which would interact specifically a t the flavin site of the enzyme when xanthine oxidase is in the 0 form. Although there seem to be remarkably few such compounds, one example appears to be the antitrichomona drug, 1-(2’hydroxyethyl) -2-methyl-nitroimidazole (253). 247. 248. 249. 250. 251. 252. 253.

I. Fridovich, JBC 237, 584 (1962). J. M. McCord and I. Fridovich, JBC 243, 5753 (1968). I. Fridovich, JBC 242, 1445 (1967). I. Fridovich and P. Handler, JBC 237, 916 (1962). R. W . Miller, Can. J . Biochem. 48, 935 (1970). L. Della Corte and F. Stirpe, BJ 117, 97 (1970). R. Fried and L. W . Fried, Biochem. Pharmacol. 15, 1890 (1966).

6.

MOLYBDENUM HYDROXYLASES

367

Let us now consider, insofar as they are known, the sites of interaction with the enzyme of various oxidizing substrates apart from NAD’ and oxygen. Whereas reducing substrates, particularly relatively complicated molecules such as purines, might a priori be expected to interact a t a single, specific site only (i.e., a t the molybdenum site) oxidizing substrates such as dyes would not necessarily show such specificity. That all oxidizing substrates do not interact a t the flavin site is clear from the retention of activity in the deflavo enzyme, e.g., to ferricyanide. Further, in view of evidence (Section II,C,l,b; 190) for direct oxygen transfer from nicotinamide-N-oxide to uric acid, there can be no doubt that the oxidizing and the reducing substrate can sometimes interact a t one and the same enzyme site. I n this case, the site involved must be the one a t which xanthine interacts, i.e., molybdenum. Similarly, it has been concluded (6‘1) from fast kinetic studies that phenazine methosulfate interacts via molybdenum. There seems, then, t o be no doubt that both the flavin and the molybdenum sites of xanthine oxidase are available for interaction with acceptors. However, it is not entirely clear whether all existing data could be explained simply in terms of all acceptors interacting a t one or both of these sites only. If it is not so explicable, then it becomes necessary to postulate additional acceptor sites in the enzyme (f?50,254),presumably associated with the Fe/S systems (63,75d). Although further work is needed, there does seem to have been an unjustified tendency for workers to prefer to postulate extra sites in the enzyme, in preference to the idea of individual acceptors reacting a t more than one site. Perhaps it is premature to speculate, but it seems likely to the author that the iron-sulfur centers of xanthine oxidase will be buried within the enzyme molecule (cf. ferredoxin; 255a), thus leaving only flavin and molybdenum sites available. It also seems that reactions of acceptors a t these two sites can probably explain all presently available data, although the field is somewhat confused (255). A direct experimental approach to the question of additional sites in 254. K. V. Rajagopalan and P. Handler, JBC 239, 2022 (1964). 255. Reactions due to 02-explain much early “inhibition” data (661,866); further, the role of coenzyme Q in early preparations of aldehyde oxidase (266)is uncertain. To be fully meaningful, studies of a given inhibitor, affecting two different catalytic activities of a molybdenum hydroxylase, should take account of the complex kinetics of the catalytic reaction and of the poasibilities of substrate inhibition. The concentration of each reducing and oxidizing substrate ought to be varied systematically to determine the mode of inhibition. In no case does this seem to have been done (but, also see 186). 255a. E. T. Adman, L. C. Seiker, and L. H. Jensen, JBC 248,3987 (1973).

368

R. C. BRAY

the enzyme, not apparently tried so far, would be to render both the flavin and the molybdenum sites of xanthine oxidase inoperative simultaneously. This could be achieved by preparing deflavo desulfo enzyme. Reoxidizability of this species by various acceptors could then be studied, following, e.g., on reduction by dithionite. b. Mechanism of Interaction with Reducing Substrates: General Comments. Many workers (e.g., 2,5,&,46,69,178,255b,256) have proposed mechanisms for substrate oxidation by xanthine oxidase, although obviously the earlier ones were based on rather incomplete experimental data. As noted earlier (Section II,C,l,b; 190-192),direct transfer of oxygen from molecular oxygen to the reducing substrate takes place only under special circumstances. A few general points may be mentioned here. It now seems agreed that a nucleophilic reaction mechanism with the reducing substrate must be involved. There is no experimental evidence for the formation of substrate-free radicals (5,257), and molecular orbital calculations (166) are likewise against a one-electron mechanism. On the other hand, one may wonder whether the substrate transfers a hydride ion to the enzyme (44,46,258) or whether it transfers two electrons and a proton (255b). Although, as Massey and co-workers (44) pointed out, specific reduction of the active enzyme by borohydride (Section II,C,l,a) seems to provide strong evidence favoring the former, it may be well to remember that reductions by borohydride are not necessarily simple processes (see, e.g., 259). In the following three subsections, experimental data bearing directly on the reaction mechanism of the enzyme will be summarized from an essentially historical viewpoint. We shall then proceed to consider mechanisms proposed by Olson et al. (46),and this will be followed by discussion of a number of other topics. c. Analysis of Steady-State Kinetics. Detailed analysis of the steadystate kinetics of oxidation of xanthine by oxygen, catalized by milk xanthine oxidase, was first reported by Fridovich (182a) and extended by Massey et al. (35). The finding o f parallel Lineweaver and Burk plots, as the oxygen concentration was varied a t a series of fixed xanthine concentrations, these plots remaining parallel in the presence of competi255b. G. A. Hamilton, Progr. Bioorg. Chem. 1, 83 (1971). 256. E . I . Stiefel, Proc. Nut. Acud. Sci. U.S. 70, 988 (1973). 257. Ackerman and Brill ($36) obtained evidence from magnetic susceptibility measurements for production of substrate free radicals by the xanthine oxidase system. Although they originally assumed these to be xanthine radicals, it now seems much more likely that their results were in fact due to 02-. 258. K. J. Rajagopalan and P. Handler, JBC 239, 2027 (1964). 259. P. M. Treichel, J. P. Stenson, and J. J. Benedict, Znorg. Chem. 10, 1183 (1971).

6.

369

MOLYBDENUM HYDROXYLASES

tive inhibitors, was taken as evidence for a ping-pong mechanism (260) of the type shown in Eq. (7) (661).In this binary complex mechanism,

1 Xanthine

uric acid

(7)

E

EX ZE’U

E’

E‘O,ZEH,O,

E

the two half-reactions occur separately from one another. The half-reactions are, respectively, reduction of the enzyme by xanthine followed by dissociation of uric acid and reoxidation by oxygen, with subsequent dissociation of hyrogen peroxide. Similar conclusions about the mechanism were reached in the case of chicken liver enzyme (IOS), although later work has shown that both for this and for the turkey enzyme ping-pong kinetics are applicable only over a limited range of substrate concentrations (169,626).To account for complex nonlinear Lineweaver and Burk plots for the chicken enzyme, observed over a very wide range of substrate concentrations, a much more complicated scheme, in which the two half-reactions are not separable, has been proposed (226).Nevertheless, the simple scheme of Eq. (7) still seems capable of explaining most data, a t least from the milk enzyme (662). d. Titration of the Enzyme with Reducing Agents. Many studies involving titration of xanthine oxidase with reducing agents have been reported (6,26,S7,SS146,90b,I1 7,146,156,26S) , and some of this work has already been mentioned in Section II,B,4; in particular, the stoichiometry of the reduction has been discussed. Reduction is a complicated process particularly when desulfo and demolybdo enzymes are present; however, experimental results are not in dispute in any serious way and titration studies have been of the greatest importance in attempting to elucidate the roles of the various redox-active groups in reactions of the enzyme. Although reduction has also been followed optically and by magnetic susceptibility measurements, E P R measurements have proved the most revealing. Figures 3a and 3b show early anaerobic E P R titrations of the enzyme with xanthine, reported by Bray et al. (26), while Fig. 3c shows a titration with dithionite by Beinert ( 146). Figure 4 gives more recent data on dithionite reduction from Massey’s laboratory (46). Detailed 260. W. W. Cleland, “The Enzymes,” 3rd ed., Vol. 2, p. 1, 1970. 261. The mechanism of Eq. (7) for xanthine oxidase seems first to have been put forward in 1959 on the basis of stopped-flow data by Gutfreund and Sturtevant (181).

262. But see, e.g., footnote (1%). 263. R. C. Bray, G. Palmer, and H. Beinert, JBC 239, 2667 (1964).

u ? o

1.0

0.5

c

._ c

'"1 I

f.5

Xonthine /total FAD

-i .e >

I

6.0

I

o

I

2

3 4 Xonthine /active centers

5

a

Xonthine /total FAD

1.2

i L

0.8

0.4

r

3.4

3 Dithionite /total FAD

FIQ.3. Titrations of milk xanthine oxidase with xanthine [(a) and (b)] or with dithionite (c), at p H 8.2. For the xanthine titrations, a short reaction time (2 min a t 4 O , with separate samples for each point) was employed; thus, active enzyme only has reacted. The sample used in (b) contained little of the active form and therefore only weak signals were observed. I n the dithionite titration in (c), both active and desulfo enzymes have reacted [although the reaction time had to be extended to 45 min for observation of the Slow Mo(V) signal]. The percentages of active enzyme in the xanthine oxidase samples for (a), (b), and (c) were 37, 6, and 59, respectively. The horizontal scale is generally in moles of reducing agent per half-molecule of total xanthine oxidase. In (a), the scale relating to active enzyme is, however, more appropriate. (A)Mo(V) Rapid, (0) FADH',).( Fe/S I, and ( A ) Mo(V) Slow. (a) and (b) are replotted from Bray et al. (26) and (c) from Beinert (146); contents of active enzyme in Bray et al. ($6) have been recalculated. 370

6.

MOLYBDENUM HYDROXYLASES

371

Dithionite /total FAD

FIG.4. Experimental and computed curves for the titration of milk xanthine oxidase with dithionite a t p H 8.5; (a) fractional amounts of the Fe/S systems in the reduced state, and (b) amounts of reduced molybdenum and flavin. The points represent experimental values from E P R and optical measurements, while the curves are theoretical, computed from the redox potentials given in Table IX. Two different methods of calculating the experimental Fe/S I and FADHz values were used; these and 0 ;other symbols are: (0) FADH', and (A)Fe/S I1 or Mo(V) are shown as Rapid. The vertical scale is calculated to that expected for 100% active enzyme and the Slow signal from the desulfo form is not shown. The horizontal scale is in moles of dithionite per mole of FAD. Reproduced from Olson et a2. (46).

interpretation of the titration curves, which had only recently been attempted when this chapter was prepared, will be reserved for a later section. We simply note here that it is quite clear that all of the redox components of the enzyme do not titrate together and that there are differences between xanthine reduction and dithionite reduction. Samples containing much desulfo enzyme are not readily reducible by xanthine (Fig. 3b). On the other hand, desulfo enzyme is reduced by dithionite and, with relatively large amounts of the reagent, gives the Slow Mo (V) signal (Fig. 3c). It is also important to note that with increasing amounts of either xanthine or dithionite the Rapid signal first increases as Mo(V1)

372

R.

C.

BRAY

is reduced to the signal-giving Mo(V) but then decreases once again as the metal is further reduced to the Mo (IV) state. That this disappearance of the signal is the result of Mo(1V) appearance as was originally proposed (26) now seems to be generally accepted, although other interpretations (2,70) were still being considered comparatively recently. e. Pre-Steady-State Kinetic Studies. Although stopped-flow work on xanthine oxidase has been important, it is exclusively the rapid freezing EPR method (29-32) which has provided information on the functioning of molybdenum in the enzyme and on differences between the two iron-sulfur centers. Most stopped-flow work has been carried out a t 450 nm, where both flavin and Fe/S absorb. At 550 nm, absorption is mainly from the Fe/S centers, while a t around 600 nm the flavin semiquinone also contributes. The earliest stopped-flow work was by Gutfreund and Sturtevant (181) and led these workers to propose the mechanism of Eq. (7). They concluded that formation of Michaelis complexes with xanthine and with oxygen was rapid and that reduction of the enzyme was slower than the reoxidation process. Later, more extensive work by Massey el al. (36) gave a somewhat higher velocity constant for enzyme reduction by xanthine and confirmed that reduction was generally rate limiting in enzyme turnover. Measurements a t 610 nm gave evidence for FADH’ formation. The work of Olson et al. (46)will be considered in later sections. Early rapid freezing experiments on xanthine oxidase were mainly of the “single turnover” type; i.e., they employed an amount of xanthine which was roughly stoichiometric with the enzyme, with an excess of oxygen. Results of the first such experiment by Bray (33) are reproduced in Fig. 5 (664). A Mo(V) EPR signal (presumably mainly the Rapid signal) appeared and disappeared with a time-course close t o that predicted from 450 nm stopped-flow data of Gutfreund and Sturtevant (181). This experiment was interpreted as providing the first direct evidence for involvement of molybdenum in the catalytic processes of the enzyme, with the metal cycIing between the six- and five-valent states. These early experiments also provided some indications of FADH’ and Fe/S involvement in enzyme turnover. More detailed work was reported by Bray et al. (663) and has been widely quoted. These workers distinguished two distinct Mo(V) EPR signals, now called Very Rapid and Rapid. The former signal was detected only as a transient during reduction with xanthine, its intensity being accentuated a t high p H values. I n single turnover experiments, the 264. In the original experiments (33) a second xanthine oxidase sample did not behave as in Fig. 5, but evidence was presented clearly suggesting that this resulted from the presence of an inhibitor in the second sample.

6.

373

MOLYBDENUM HYDROXYLASES

t

b

0

,

I

I00

200

I

300

I

400

Time (msec )

FIG.5. Time-course of development and disappearance of Mo(V) E P R signals from milk xanthine oxidase in a “single turnover” rapid freezing experiment, with xanthine, a t p H 8.2. The curve is theoretical calculated for successive appearance and disappearance of a Mo(V) intermediate, with rate constants of 10.5 sec-l and 21.5 sec-l, respectively. It thus corresponds to the curve predicted from the mechanism of Eq. (7), assuming the first-order intramolecular reduction and reoxidation steps are rate limiting and with the rate constants of Gutfreund and Sturtevant’ (181). The concentration of xanthine oxidase (active centers) was 0.05 m M and those of xanthine and oxygen, 0.06 and 0.6 mM, respectively. The open and filled circles represent two different experiments. Replotted from Bray (33).

two molybdenum EPR signals and also the signals from Fe/S I and free radicals were all clearly distinguished and quantitated on a relative basis [although the FADH’ and 02-radical signals were not distinguished; see (6)].All the signals appeared and disappeared again in times comparable to the turnover time of the enzyme, and one of the most striking features of work was the finding (263) that each EPR signal reached its maximum intensity at a time different from the other signals. The experiment was widely accepted (e.g., 264a) as indicating an electron transfer sequence in enzyme turnover in accordance wit,h the time sequence of the signals, i.e., from substrate to molybdenum, to flavin, to iron, and so to oxygen. However, that this was not the true explanation of the results was sug264a. H. R. Mahler and E. H. Cordes, “Biological Chemistry,” 1st ed., p. 579. Harper, New York, 1966.

374

R. C. BRAY

gested by later work, e.g., Bray et al. (265) studied the effect of varying the oxygen concentration on the signal time-courses in single turnover experiments with l-methylxanthine. They found oxygen-dependent differences in the relative time-courses of the Mo(V) and Fe/S I signals. The tendency of the latter signal to linger on its own was enhanced a t higher oxygen concentrations, whereas the reverse would have been predicted from the original “sequence” model. The true explanation of the signal sequences will be considered further in the next t8wosubsections. Although the single turnover rapid freezing method has thus continued to provide useful results, it has to a considerable extent been superceded in more recent work by direct and separate studies on the reduction (46,128) and the reoxidation (45) processes of the enzyme. Such work is technically more difficult since rigorous anaerobic procedures are required. Results will be considered below. Finally, the “pH jump” variation of the rapid freezing method has also been applied to xanthine oxidase. I n this procedure the pH of reduced enzyme samples is suddenly changed by addition of another buffer and the samples are then frozen within a few milliseconds. The method was first used (128) to show that the Very Rapid and Rapid Mo(V) signals are related to one another by reversible uptake and loss of a proton. Later, however, it was realized (46) that the same experiment also provided information on the rapidity of the internal electron transfer processes of the enzyme (see also 265). f. Reduction of the Enzyme. Let us now consider the reduction of the enzyme in more detail. Morell (47) was the first to make detailed studies on the anaerobic reduction of xanthine oxidase preparations of moderate activity by substrate. He found a fast phase of reduction, as followed at 450 nm, which was presumed to be catalytically significant, followed by a slow phase. Subsequent work on the reduction processes, using either optical or E P R detection, has been extensive (SS,S5,46,117,1~5,1~8,15~, 263). In particular, Massey and co-workers (46,128), have reported comprehensively on the fast part of the reduction, while Swann and Bray (125) concentrated on the slower steps, distinguishing no less than four phases in the overall reduction processes. A very important development in 1972 (44) [foreshadowed in 1970 (58,61)]was the finding that pure active xanthine oxidase, prepared by affinity chromatography, has virtually no slow phase in its reduction by excess xanthine, as followed a t 450 nm. This observation confirmed the original proposal of Morell (47) that the slow phase is not the result of reduction of active enzyme but rather the reduction of inactive enzyme by reduced active molecules (see also Section 11,CJ2,k). 265. R. C. Bray, D. J. Lowe, and M. J. Barber, BJ 141, 309 (1974).

6.

MOLYBDENUM HYDROXYLASES

375

This section is concerned only with the reduction of active enzyme, i.e., with phases I and I1 of Swann and Bray ( 1 2 5 ) .These phases are complete within a few seconds when xanthine is employed to reduce the enzyme. It has been fortunate for experimenters that there are no indications of interference in these two phases by desulfo (or demolybdo) enzyme. At short reaction times, these enzyme forms serve merely as a diluent of the active enzyme. Thus, valid fast reaction kinetic studies on reduction of active enzyme may apparently be made using xanthine oxidase samples having virtually any Actvity/E,,, value. The main findings regarding reduction of the active enzyme by excess xanthine a t p H 8.5 will now be summarized. It should first be noted that with proper allowance for desulfo enzyme reduction of the EPR and optically detectable chromophores of the active form by the substrate proceeds rapidly and goes very nearly, although perhaps not entirely, to completion, as judged by comparison with reduction by dithionite (266). Uptake of six electrons per functional active center (from three xanthine molecules) would be required for this process (108). Stopped flow and rapid freezing data on the time-course of the reduction correlate well with one another (46). All the possible E P R signals are detectable at the shortest reaction times (267’)and these signals then increase in intensity, although not always with simple first-order kinetics. The Fe/S signals increase to steady values, whereas the molybdenum and FADH. signals attain maximum intensities, each a t a different reaction time, then decrease again. The relevant data are summarized in Table VIII. Reduction of the enzyme by 1-methylxanthine is very similar to that by xanthine, except that, first, the overall process is somewhat faster and, second, the Very Rapid signal is not observed a t pH 8.5, although it is detected in small amounts at higher p H values (128). Detailed reduction studies have not been reported for other substrates; however, a few interesting points may be noted. The Very Rapid signal is almost exclusive to xanthine (99) and is always a transient, even in partial reduction experiments employing an excess of the enzyme over xanthine (46). In contrast, the Rapid signal is given by every substrate which has been tested (6) (including even allopurinol ; 125,268), and under appropriate conditions this signal is stable indefinitely. Finally, it is interesting that with 266. It is quite possible that some substrates are incapable of reducing the enzyme fully. This may be the case with salicylaldehyde (cf. 28). 267. I n contrast to these findings a t p H 8.5 ( I d @ , at p H 10.1, the Rapid Mo(V) and FADH signals are not seen a t the shortest, reaction times (15s). 268. In the presence of excess allopurinol, disappearance of the Rapid signal as the result of reduction to Mo(IV), is faster and apparently more complete (136) than is the corresponding process with xanthine (Table VIII).

376

R. C. BRAY

TABLE VIII TIME-COURSE O F REDUCTION OF ACTIVEXANTHINE OXIDASE BY EXCESSXANTHINE AT PH 8.5 A N D 25""

Species Very Rapid Mo(V) Rapid Mo(V) FADH Fe/S I Fe/S I1

Maximum yield (e/active center)

Yield a t longer times (elactive center)

0.075 0.105

<0.05*

0.080 1 .o

1 .o

0

Formation It (msec)

Od

5 15 35

1.o 1.o

80 60

Decay tt max (msec (msec) from max) t

15 65

80 -

40 500" 150

-

-

Data from Olson et al. (46).I n Edmondson et al. (188) somewhat faster reduction was reported when using a somewhat higher xanthine concentration. bAlthough it might have been expect,ed that the Rapid signal would ultimately disappear as a result of reduction to Mo(IV), this has not been reported experimentally. This most likely results from the late stages of reduction by xanthine being slow, as well as from interference from desulfoenzyme [via the onset of phase I11 of Swann and Bray (126)l. Swann and Bray (185) reported that the rate of this process, at a fixed xanthine concentration, was dependent on enzyme concentration. The reason for this is not clear. Data from Bray et al. (863).

salicylaldehyde, in contrast to xanthine and l-methylxanthine, apparently the radical, Rapid, and Fe/S I signals all develop together with similar time-courses (70). Important information relating to the enzymic mechanism has been provided by kinetic studies on reduction of xanthine oxidase using deuterium-labeled substrates. These experiments have revealed two phenomena, namely, changes in the form of the Rapid signal and kinetic isotope effects on the rates of some of the reaction steps. The first observations were made in 1968 by Bray and Knowles (196). They found, using xanthine specifically deuterated in the position a t which the hydroxylation occurs (i.e., the 8 position), that a t very short reaction times the Rapid signal appeared in a modified form. The spectral modification, which disappeared at somewhat longer reaction times, took the form of partial replacement of the characteristic proton doublet of the signal by a single deuterium line. This was interpreted as evidence that direct hydrogen transfer, from [8-2H]xanthine to the enzyme, took place during turnover. The observation was later extended to [!WH]-1-methylxan-

6.

MOLYBDENUM HYDROXYLASES

377

thine (69,70,1@a) and confirmed in another laboratory (168,269,270). So far, little attention has been paid to studying the rate at which 2H disappears from the Rapid signal, although such work might in principle give information on the nature of the proton binding group in the active center. The rate of the exchange reaction with protons from the solvent appears t o be several times faster than the enzyme turnover rate (69,196). The kinetic isotope effects noted by Bray ed al. (69)were then studied in some detail by Edmondson el al. (128). The latter workers compared reaction of deuterated and normal xanthine and l-methylxanthine, working both at pH 8.5 and a t pH 10.0. They reported that so far as overall turnover was concerned there was relatively little difference in Vmaxvalues between the deuterated and nondeuterated substrates. There were, however, quite substantial effects on apparent K , values, e.g., a t pH 8.5 the value for [8-ZH]-l-methylxanthine was 3-4 times higher than that for ordinary l-methylxanthine. With regard to rates of development of the EPR signals, they found with the deuterated substrates, under all conditions, slower development of signals whether from Mo(V), FADH-, or Fe/S. Isotope effects on the half-times of appearance of the individual signals varied from 1.06 to 3.2, the smallest effects usually being on Fe/S signals. Before discussing the interpretation of all the above data on the reduction of xanthine oxida,seby substrates, it will be helpful to consider a few general points. In principle, at any given moment during reductive or oxidative processes, a given xanthine oxidase half-molecule might have accepted any number of electrons between 0 and 6 from the reducing agent. Let us designate the corresponding species as XOO,XO1,, XOZ,, . . . , X06,. Clearly, in principle, each of these species should be regarded as a distinct chemical entity. However, application of this simple concept to interpretation of xanthine oxidase reduction data is a surprisingly recent development. The author was forced to recognize that species generated during reduction by l-methylxanthine, presumably XOne and X04,, are fundamentally different from one another, by E P R studies on 269. *H effects in the spectra of Edmondson et al. (128) seem less dramatic than those in Bray and co-workers (69,70,148a) ; this might be because the quenching times in the rapid freezing procedure were longer in the work at Ann Arbor (32). 270. Although the phenomenon is clear and the original interpretation probably correct, a note of caution must be added. Since the form of the Rapid signal is modified by formation of complexes between reduced enzyme and substrate molecules, it could conceivably be that the ’H “seen” in the signal is on an unreacted substrate molecule, complexing the enzyme, and not on the enzyme molecule itself. Unambiguous evidence for direct hydrogen transfer to the enzyme from study of signal forms would come from work under conditions where substrate complexes of the Rapid signal are excluded.

378

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interaction between Mo(V) and Fe/S I (263;Section II,B15).Interaction between the signal-giving species is not detected in the early stages of reduction but is in the later ones; i.e., interaction in effect serves here as a diagnostic test for appearance of XO,, as the reduction proceeds. A further simple experiment in the author’s laboratory (166)showed that XOne, generated by reduction with 1-methylxanthine, differs in quite a striking manner from XO1, generated from it by brief exposure to small amounts of the one-electron oxidizing agent, phenazine methosulfate (but observed a t the same total reaction time as the control). Comparison of the relevant signal intensity ratios showed that there was substantially more Fe/S I relative to Mo(V) in XO1, than there was in XOI,, thus indicating a distinctly different distribution of elecdrons in the two species. The next question is the rates of the intramolecular electron transfer reactions within the enzyme molecule. In other words, if, e.g., two electrons are introduced into the xanthine oxidase molecule via the molybdenum, how quickly do these electrons redistribute themselves toward an equilibrium situation in which they are, let us assume, distributed in a statistical manner over all the redox-active groups of the enzyme half-molecule? The original assumption (263) was that these intramolecular reactions were relatively slow ones, possibly rate limiting in enzyme turnover (70). However, it has since come to light that all currently available data are interpretable in terms of fast intramolecular reactions (46; see also 165). In the following discussion it will therefore be assumed that the intramolecular oxidation-reduction reactions are relatively fast processes. Later, in Section II,C12,h,the question of the actual rates of these processes will be returned to. We are now in a position to consider the interpretation for the reduction data. Olson e t al. (46) succeeded in interpreting the dithionite titration (Fig. 4) in terms of a simple model. They made the simplest possible assumption, namely, that the behavior of each of the functional groups of the enzyme was governed by characteristic redox potentials, these being independent both of the state of other groups in the same molecule and of the total number of electrons accepted by the molecule. They tried a series of different combinations of redox potentials until, finally, using those given in column two of Table IX, they were able to calculate the theoretical curves shown in Fig. 4, which are in good agreement with the experimental results (2YOa). 270a. Results of EPR titrations of xanthine oxidase with xanthine differ somewhat from those with dithionite (cf. Fig. 3a with Figs. 3c and 4), even after proper allowance for the desulfo enzyme has been made. The stoichiometry is similar with the two reducing agents. However, the behavior of the Rapid MOW) signal is different, particularly in that there is more of this signal, when xanthine

6.

379

MOLYBDENUM HYDROXYLASES

TABLE IX ASSUMEDRELATIVE REDOXPOTENTIALS FOR THE GROUPINGS OF XANTHINE OXIDASEn Relative redox potential (mV) Species

Alone

Fe/S II,,/Fe/S IIred Fe/S I,,/Fe/S Ired FAD/FADH’ FADH./FADH, Mo(V1) /Mo(V) Mo (V)/Mo(IV)

-24 -60 60 -60 -31

0

+

Xanthine bound 0 -24 -60 60 120 57

+ +

+

a The data are from Olson et al. (46)and refer to pH 8.5. All values are relat,ive to Fe/S I1 which is arbitrarily given the value zero.

It was pointed out (46) that in a system of this type the redox potentials determine (under equilibrium conditions and a t a given point in the titration) not only the average extent to which, e.g., the molybdenum atoms of all the molecules in the sample are reduced, but they also determine the proportion of the individual molecules which have accepted 1, 2, or 3, etc., electrons from the reducing agent, per active center, together with the precise statistical manner in which, in each of these reduced enzyme species, these electrons are distributed among Fe/S, Mo, and

FAD. Olson et al. (46) then went on to attempt the much more difficult task of explaining the time-course of reduction of the enzyme by xanthine. They based their explanation on the kinetic scheme and its chemical interpretation which are given in Fig. 6, in conjunction with the redox potentials of Table IX, and they were able, ultimately, to get quite good simulations both of the EPR time-course data in Table VIII and of corresponding stopped-flow data obtained under a variety of conditions. From a kinetic point of view, their scheme (Fig. 6 ) assumes that xsnthine rapidly forms a Michaelis complex with the enzyme, EX, and that formation of this can take place equally well with the oxidized enzyme (i = 0) or with is used in place of dithionite, in the region of 1 to 2 moles of reducing agent per half enzyme molecule (stoichiometry is expressed for active enzyme only, with xanthine, but for total enzyme, with dithionite). No doubt the increase in the Rapid signal results from the formation of complexes of reduced enzyme with xanthine, etc. (Section II,B,4$), although no theoretical titration curves for xanthine reduction seem to have been attempted.

380

R.

C. BRAY

FIG.6. Kinetic and chemical schemes representing the reduction of xanthine oxidase by xanthine, as proposed by Olson et al. (46). E X ( i ) is a Michaelis complex and E’X(i 2) an intermediate in which the xanthine residue haa become covalently bound and two electrons have been transferred to the enzyme and reside mainly on the molybdenum. I n E(i 2) the reaction has been completed and the product has dissociated and the two electrons of the reduced enzyme are now localized mainly on the flavin and Fe/S centers. Reproduced from Olson et al. (46).

+

+

reduced forms of it (i = 2 or 4). The complex is then converted in a first-order process to the intermediate, E’X, which in turn yields the reduced enzyme and uric acid in a further first-order reaction. Each enzyme molecule will go through three cycles of the reactions with successive xanthine molecules (corresponding to i = 0, 2, or 4 in the scheme) , until it is fully reduced. Electron distributions in E,, and E, (and also apparently in the corresponding E X species) were assumed to be governed by the redox potentials of the free enzyme, as deduced from the dithionite titration (see Table IX). On the other hand, in E’X2, E’X,, the redox potentials of the molybdenum are supposed to have been substantially raised by the presence of the bound xanthine residue. Thus, the modified redox potentials given in the right-hand column of Table I X were supposed to govern the electron distributions in these species. I n order to obtain good simulations of the experimental time-courses from the scheme, some further assumptions had to be made. In particular, it was assumed that the rate constant, k,, depended on the value of i, the actual k , value for each of the successive reduction steps being proportional to the fraction of the molybdenum in the corresponding EX complex which was present in the six-valent state. (This was to express the idea that xanthine could react by transferring two electrons only in those molecules of the complex in which the metal was in the fully oxidized state.) The further and rather more arbitrary assumption had to be made that the Very Rapid Mo(V) signal was given solely by the species E’X, of which 65% was in the unprotonated form, whereas the remainder of this species and indeed all of the other Ma(V)-containing species were in the protonated state, thus giving rise to the Rapid signal. On the basis of the kinetic scheme of Fig. 6, with these assumptions,

6.

MOLYBDENUM HYDROXYLASES

38 1

quite convincing simulations of most of the EPR and stopped-flow reduction time-course data were obtained. The value of k , used was 25 sec-I. Kd and k , could not be separately estimated, but the indications were that Kd must be very small. Many of the data could in fact be explained by ignoring the Michaelis complex altogether and putting k , as a secondorder reaction, with a rate constant of.2 x lo6 M-' sec-I. An important feature of the scheme is that it accounts adequately for the transient nature of the Very Rapid signal under all conditions, since the signal is assumed to arise from a species (E'X,) , which always decays by an essentially intramolecular process. The scheme predicts complete reduction of the enzyme by excess xanthine, with full conversion of molybdenum to Mo (IV), which has not apparently been observed experimentally (Table VIII). No doubt it could be modified, though, for incomplete reduction if slow reversibility of reduction was allowed (see Section II,C,l,e). Apart from reduction time-course data, the scheme of Fig. 6 was claimed (46) to explain, and does indeed seem capable of explaining, an impressive range of additional, unusual, phenomena observed with xanthine oxidase. Among these are the following (46): 1. The small apparent lag observed in semilogarithmic plots of the early stages of reduction of the enzyme, followed in stopped-flow experiments a t 450 nm, is a consequence of the three-step nature of the reduction process. 2. Similarly, the apparently faster reduction of deflavo than of normal enzyme, is a consequence of the fact that here only two successive xanthine molecules, rather than three, have to react to give complete reduction. 3. The deuterium rate effects are consistent with k , involving rupture of the C-*H bond. 4. The slowness of the later stages in reduction of the enzyme by NADH (35,166), which reacts in a two-electron reduction via the flavin, appears as a consequence of the fact that in XO,,, all but about 2% of the flavin is present in reduced forms and is therefore not available for reaction with the third substrate molecule. When now considering the chemical, as opposed to the kinetic, interpretation of the scheme of Fig. 6 , it has to be recognized that most of its features are speculative, even if they do seem attractive. I n the reaction step governed by k,, the persulfide group attacks the C-H bond to form an intermediate with the xanthine residue covalently bound to the enzyme, while a t the same time two electrons are donated to molybdenum and the proton is taken up by a nitrogen ligand of the metal. Interaction

382

R. C. BRAY

between the xanthine residue and the metal is supposed to account for the increased redox potentials of the latter in the intermediate (Table I X ) . (It will be remembered that these higher potentials had to be assumed to simulate the kinetics.) The final step ( k 3 ) is hydrolysis of the persulfidnxanthine bond by water, to yield the product. It is assumed (46) that OH- is not involved in this step in view of the fact that V,,, is independent of pH over a wide range (Section II,C,l,f). The scheme seems satisfactory within the limitations that, first, there is no direct experimental evidence whatever for an intermediate with substrate covalently bound to the enzyme via the persulfide group or otherwise, and, second, that in any case the chemistry of persulfides is not well understood (44). The possibility that the persulfide group is an essential ligand of molybdenum (125), rather than a direct participant in the catalytic reaction, also has to be borne in mind. If this were the case, then entirely different types of mechanisms ought to be considered, including perhaps the following scheme, which is slightly modified from one proposed (26Bb)by Hamilton. In this, a hydroxyl originally on molybdenum adds across the carbon-nitrogen double bond adjacent to the 8 position of xanthine, leaving an -OMo(enzyme) residue on the carbon. Then two electrons are transferred to molybdenum, while the hydrogen in the xanthine C-8 position leaves as a proton. One unsatisfactory feature of this scheme is, of course, that as the substrate C-H bond is broken a t a late stage in the reaction, the isotope effects on signal appearance rates are difficult to explain. Nevertheless, this scheme may serve here a t least to reemphasize the provisional nature of many of the chemical postulates of Fig. 6. Despite its unproved nature, however, the scheme of Fig. 6 does have attractive features. If it were located in a suitable orbital, the proton on the nitrogen ligand of molybdenum could presumably give the observed proton splittings of the Rapid signal and the observed deuterium effects. Furthermore, the scheme is founded on the concept developed by Stiefel (266)of linked proton and electron transfers in molybdenum enzymes, this being based on increasing proton affinities of ligands of the metal (such as the nitrogen in Fig. 6) as the valency of the metal decreases. Finally, a point not previously made is that formylation (98) of the same nitrogen atom by formaldehyde could again reasonably be expected, given suitable geometry, to yield the Inhibited signal. It is concluded, therefore that Fig. 6 must be a fair approximatirJn toward the true mechanism whereby reducing substrate molecules interact with the active center of xanthine oxidase. g. Reoxidation of the Enzyme b y Oxygen. The mechanism of oxidation of reduced xanthine oxidase has been much less intensively studied than

6.

MOLYBDENUM HYDROXYLASES

383

has the reduction process. The reason for this is the obvious technical one. It is no easy matter to produce the necessary starting material, i.e., reduced xanthine oxidase, which is either fully reduced and free from excess reducing agent or, alternatively, which has been only partially reduced but t o a known and reproducible extent. Bray et al. (663)made some very preliminary attempts to study directly the reoxidation by molecular oxygen of reduced xanthine oxidase. However, it was not until much more recently that Olson et al. (&), made a systematic, although by no means exhaustive, study of the problem. Virtually no work seems to have been done on the mechanism of reoxidation by electron acceptors other than oxygen. Reoxidation by oxygen was investigated by Olson et a2. (45,QB) in stopped-flow experiments a t various wavelengths and also by rapid freezing EPR. Their most important conclusion was that reoxidation is a twophase process, with the second phase occurring some 10 times slower than the fast one and a t a rate which is considerably slower than enzyme turnover under normal conditions. There were found to be distinct differences in the spectral changes associated with the two phases. In the slow phase, at pH 8.5,there was a substantial increase in absorption at 550 nm. Furthermore, the spectral changes which generally occurred in this phase resembled the difference spectrum (oxidized-reduced) of deflavo xanthine oxidase. The slow phase was therefore taken to be primarily associated with reoxidation of the Fe/S systems of the enzyme. Rapid freezing E P R confirmed this and showed further that it was mainly Fe/S I1 which was being reoxidized in the slow phase, although, to a lesser extent, other EPR-detectable chromophores were also involved. I n their reoxidation work, as in reduction studies, Olson et al. (45) did not remove desulfo xanthine oxidase. However, they showed (spectrophotometrically, a t least) that the reoxidation of normal and cyanidetreated enzyme were indistinguishable from one another. (See, however, discussion of reoxidation of the Slow signal in Section 1I,C12,k.) The twophase behavior was not influenced then by the activity/E450 ratio of the enzyme and, further, it was not influenced when enzymic activity was inhibited by treatment with allopurinol. On the other hand, the relative extents of the two phases could be changed markedly by varying the level of reduction of the enzyme samples. With fully reduced xanthine oxidase, the slow phase amounted to some 2076 of the total change a t 450 nm, but for partially reduced enzyme this figure increased, the slow phase becoming essentially the only one observed a t low extents of reduction. Kinetic studies a t various oxygen concentrations revealed saturation behavior in the fast phase but a pure second-order reaction in the slow phase.

384

R.

C.

BRAY

Finally, production and possible involvement of the superoxide ion,

02-,in the reoxidation processes were studied (46). Yields of the radical (determined by EPR; 197) were relatively low, indicating that HzOzwas the main product of oxygen reduction. Addition of superoxide dismutase did not modify the reoxidation kinetics in any way. This is in contrast to its pronounced effects on autoxidation of free flavins, which depend on reaction between Oz- and FADH’ (201). Olson et al. (46) proposed that the above observations could be explained, if, of the various possible one- and two-electron oxidation steps for reduced xanthine oxidase molecules, those in Eq. (8) were the dominant ones.

I n this scheme, all the steps except the lust one, namely, the oxidation of XOI,, are supposed to involve formation of a Michaelis complex between oxygen and FADHz in the relevant enzyme species, this complex then decaying by a first-order process. It was proposed that, in the t>woelectron reoxidation steps, the flavin cycled twice, rapidly, between the FADHZ and FADH. levels as it transferred two electrons to the bound oxygen molecule. I n this process, the flavin was supposed to become reduced again by intramolecular electron transfer, with this occurring before the product had had time to dissociate, so that HzOz was formed rather t*han02-.On the other hand, in the oxidation of XOz,,there would not be enough electrons available for the second flavin cycle to occur, so that 0 s - would be formed. Similarly, in the last step, there could be no FADHg, so that oxidation was then presumed to occur by reaction bctween the FADH. in XOI, and oxygen. This was taken to be a slow second-order reaction, corresponding to the slow phase of the oxidation process. In free flavins, the corresponding reaction between FADH. and oxygen is fast only for the “red,” not the “blue” semiquinone (271). Olson et al. (46) were able to account for many of the observed features of the reoxidation process in terms of the above model. They first, as in the reduction studies, calculated the electron distributions expected in XOee, X06el . . . , XOI,, on the basis of the redox potentials of Table IX. Then, on the basis of normal extinction coefficients for FADHz and FADH’ in the enzyme, together with assumed values (Table V) for the 271. M. Faraggi, P. Hemmerich, and I. Pecht, FEBS Lett. 51, 47 (1976).

6.

MOLYBDENUM HYDROXYLASES

385

extinction changes associated with oxidation of the two Fe/S systems, they were able to predict quite accurately the percent of slow phase oxidation expected a t 450 or 550 nm, as a function of the extent of reduction of the enzyme. On the basis of the first-order reaction step (decay of the Michaelis complex of oxygen wit.h FADHz in the enzyme) being governed by a rate constant of 205 sec-’, they were also able to simulate satisfacterily the reoxidation time-courses a t the two wavelengths. Although quantitative agreement with the rapid freezing EPR data was somewhat less good, nevertheless, qualitatively, most of the observed features could also be simulated. Finally, the model also roughly accounted for the observed yields of 02-and accounted qualitatively for observations by Fridovich (200) that 02-formation is stimulated by high oxygen concentrations or by low xanthine concentrations. I n order to account for increased formation of FADH’ in the fast phase of reoxidation a t p H 6.0, relative to that a t p H 8.5 (46), it was necessary to assume that some of the redox potentials of Table IX change with pH, although this problem has apparently not been investigated in detail. h. Rates of the Internal Electron Transfer Processes. As has already been discussed in Section II,C,2,e, i t was originally assumed (263) that intramolecular electron transfer reactions among the molybdenum, flavin, and Fe/S centers of xanthine oxidase were relatively slow processes. Newer work, particularly the simulation studies of Olson et aZ. (46) as discussed in the last two subsections, is not consistent with this assumption. All the simulation studies of these workers were based on the postulate of infinitely fast intramolecular electron transfer reactions. Clearly, the quality of the simulations of both oxidation and reduction processes, under a variety of conditions, must go a long way toward justifying the original assumptions. However, two questions arise: Can we put a lower limit on the rates of the intramolecular reactions and are all of them always fast? Regarding the second question, the special case of the Slow signal will be considered in Section II,C,2,k. Regarding the first, it seems clear that if the intramolecular reactions proceeded, not instantaneously, but with half-times of as long as several milliseconds, then this would not affect the quality of the simulations significantly. Olson et aZ. (45) suggested 100 sec-1 as a lower limit for the rates of these reactions. Whether these rates will, in fact, ever be measured experimentally, remains to be seen ( 2 7 l a ) . The question (cf. 29) of whether rapid freezing can stop reasonably fast and purely intramolecular, as opposed to intermolecular, reactions, must also remain open for the time being (but see 271a. Perhaps the “antifreeze” techniques of P. Douzou, Mol. Cell. Biochem. 1, 15

(1973) would be applicable to this problem.

386

R. C. BRAY

272). (In this context, we regard the reaction step governed by k , of Fig. 6, which is stopped by rapid freezing, as an intermolecular reaction involving water.) If it is accepted that all the intramolecular reactions are fast, then it follows that there can be no such thing as a “sequence” in the intramolecular electron transfer processes of xanthine oxidase since all the redox components are permanently a t equilibrium with one another. i. T h e Overall Catalytic Process. We have already considered (Section II,C,2,a) the question of the sites a t which reducing and oxidizing substrates interact with xanthine oxidase, as well as the mechanisms of reduction (Section 11,C,2,f) and of reoxidation (Section II,C,2,g) of the enzyme. It remains only to discuss a few general points about the mechanism. The mechanisms proposed by Olson et al. (46) involve xanthine molecules reducing the enzyme only when molybdenum is in the six-valent state, and oxygen molecules reoxidizing it rapidly only when FAD is in the fully reduced state. This suggested to these workers that the role of the Fe/S centers must be, by virtue of their redox potentials, t o store electrons and so facilitate regeneration of these reactive species during the functioning of the enzyme, thus maximizing the turnover rate. A further point is that in vivo, a given xanthine oxidase molecule would be able to deal relatively efficiently with some random variations in the rate a t which either reducing or oxidizing substrate molecules were presented to it. Presumably, on the basis of the scheme of Fig. 6, differences in the behavior of the various reducing substrates of the enzyme could be accommodated simply in terms of variations in the values of the individual rate constants of the different reduction steps and of variations in the properties of the various E’X species. Differences among the oxidizing substrates presumably depend to a large extent on whether individual substrates interact a t the flavin site, or the molybdenum site, or both. As already noted, further work on the reoxidation mechanisms is called for, however, and the possibility that the Fe/S sites are directly involved in reaction with some oxidizing substrates is not excluded. j . Inhibition b y Excess of Reducing Substrate. It has long been known (4) that xanthine oxidase is susceptible to inhibition by high concentrations of reducing substrates and that the extent of this inhibition is highly pH-dependent (236). No doubt the phenomenon involves some form of “over reduction” of the enzyme, but the precise mechanism remains to be elucidated. It is not clear whether formation of relatively stable com272. C. Capeilkre-Blandin, R. C. Bray, M. Iwatsubo, and F. Labeyrie, Eur. J. Biochem. 54, 549 (1975).

6.

MOLYBDENUM HYDROXYLASES

387

plexes of the reduced enzyme with substrate molecules plays a part. If such complexes are involved, then they might have molybdenum in either the four- or five-valent state. I n the former case the complexes would be analogous t o (but weaker than) those with alloxanthine (61). In the latter, they would be amenable to study by E P R (cf. 129). k . Slow Phases in Reduction of the Enzyme. The fast, and catalytically significant, phases in the reduction of xanthine oxidase by substrates were considered in Section II,C,2,f. It is now necessary t o discuss the slow phases, which have long complicated work on the enzyme and which may take as long as 2 days for completion (117,125).Slow phases in the reduction process are observed both when the reducing agent is in excess ( 4 7 ) and when the enzyme is in excess (26). The latter case has been studied in detail by measurements of difference spectra ( 4 6 ) . The phenomenon is particularly marked when very small amounts of reducing substrate are employed, and it is then attributed to differences in electron distribution in the species XO,, and XO,,. Xanthine initially converts the enzyme rapidly t o XO,, but the ultimate product, under these conditions, is mainly XO,,, this being formed by slow intermolecular reactions with tl,z of the order of 1 hr between XO,, and XO,. Spectral changes in the experiments (46) were generally in accordance with predictions from the spectral properties of these species as discussed previously, and based on the redox potentials of Table IX. It is noteworthy that slow spectral changes, similar to those produced by xanthine, were also observed on partial two-electron reduction of desulfo xanthine oxidase by NADH. When the reducing substrate is present in excess, the slow specctral and E P R changes mainly result from reduction of desulfo xanthine oxidase by the reduced active enzyme. Particularly with xanthine as reducing substrate, the most immediately striking result of these processes is the redevelopment of the Rapid Mo(V) signal, in what was termed by Swann and Bray (125) as phase I11 of the overall reduction process. The rate of phase I11 was dependent on the Activity/E,,, of the sample but for normal preparations had a half-time of the order of 10 t o 20 min. The fact that the most apparent slow E P R change, resulting from the presence of desulfo enzyme, involved signals from active enzyme molecules thus presented something of a paradox (cf. 26). However, this was resolved (46; see also 125) by the assumption that phase I11 involves a one-electron intermolecular reaction between XO,, and oxidized desulfo enzyme. Before the XO,, produced in this reaction (and giving the Rapid signal) could be re-reduced by a two-electron reaction with xanthine, a further one-electron reaction with a molecule of desulfo enzyme would be required. This would account for the extreme slowness of the final stages (phase IV) of the reduction process. Finally, we have t o consider the reduction of desulfo xanthine oxidase

388

R.

C.

BRAY

and the Slow Mo(V) signal. The iron and flavin chromophores of this form of the enzyme behave similarly in dithionite titrations (46) to these components in the active enzyme, while titration behavior relating to the Slow signal was considered in Section II,B,4,d. The term “Slow” was coined for this molybdenum signal by Bray and VanngLrd (116) and is indeed an apt one. However, why the signal should be slow, both in its appearance and in its disappearance, is far from clear, although the experimental data on this point seem unambiguous. In the author’s laboratory the Slow signal is routinely produced by reaction of enzyme for 20 min with excess dithionite ( 5 8 ) .These long reaction times are necessary (189),although the Fe/S centers appear to be fully reduced a t much shorter times (146,148a). Similarly, in reoxidation experiments the Slow signal remains after other signals from the enzyme have ceased to be detectable (163).Thus, the indications seem to be that the Slow Mo (V) signal-giving species is not in rapid redox equilibrium with the other constituents of the desulfo xanthine oxidase molecule (273). However, further work is required. 111. Other Molybdenum Hydroxylases

A. INTRODUCTION

A considerable number of molybdenum hydroxylases, all of them quite closely related to milk xanthine oxidase, are known. They include the various xanthine dehydrogenases from avian, bacterial, and other sources, as well as the aldehyde oxidases ( 2 7 3 ~ )Some . of the more studied of these enzymes are listed in Table X. Sources from which other closely related enzymes have been at least partially purified, but which will not be discussed further here, include calf (274), pig (107,275), and rat (87u,276) livers, as well as butterflies (877) and silkworms (W78).Although each enzyme seems to possess its own characteristic specificity pattern, the latest indications are that the catalytic mechanisms of all 273. Olson et al. (46) reported that in reoxidation experiments the Slow signal disappears at about the same rate as does the Fe/S I1 signal. Since titration data (Sections II,B,4,d and II,C,2,d) indicate that the Slow signal involves a low redox potential, it might have been anticipated that the Slow signal would disappear early in the reoxidat,ion process rather than along with Fe/S 11, which persists (46‘) owing to slow reoxidation of XO1,. 273a. For discussion of nomenclature of these enzymes, see Section II,C,l,a. 274. R. K. Kielley, JBC 216, 405 (1955). 275. P. E. Brumby and V. Massey, BJ &9,46P (1963). 276. P. B. Rowe and J. B. Wyngarden, JBC 241, 5571 (1966). 277. W. B. Watt, JBC 247, 1445 (1972). 278. Y. Hayashi, Nature (London) 192, 756 (1961).

6.

MOLYBDENUM HYDROXYLASES

389

molybdenum hydroxylases must be very similar to that of milk xanthine oxidase. Another enzyme whose specificity falls within the range of that of the molybdenum hydroxylases is oxypurine dehydrogenase from Micrococcus aerogenes (278a). Although this enzyme is stated to contain no molybdenum and no flavin, its reported specificity is somewhat reminiscent of that of aldehyde oxidase and a reinvestigation of ita composition and properties might be worthwhile. A most interesting point about the distribution of molybdenum hydroxylases is that some tissues, e.g., pig liver (179) and apparently also human liver (95,679), contain two such enzymes. The two then have quite different specificity patterns from one another to reducing substrates (i.e., one is a xanthine oxidase and the other an aldehyde oxidase) and the enzymes may readily be separated. In contrast, in other biological materials such as bovine milk there is only one molybdenum hydroxylase, in this case a xanthine oxidase. All the remaining molybdenum hydroxylases have been far less intensively studied than has milk xanthine oxidase. Of the other enzymes, until very recently, aldehyde oxidase had been the most intensively investigated; work on this enzyme has been reviewed by Massey ( 1 ) . However, much work of substantial importance on the xanthine dehydrogenase of turkey liver has recently become available.

B. MOLECULAR PROPERTIES A number of molecular properties of some of the molybdenum hydroxylases, together with an indication of the highest state of purity which has been achieved to date for each enzyme, are summarized in Table X (981-287). Examination of this table in comparison with the corre278a. C. A. Woolfolk, B. S. Woolfolk, and H. R. Whiteley, JBC 245, 3167 (1970). 279. D. G. Johns, J . Clin.. Invest. 46, 1492 (1967). 280. T. A. Krenitsky, J. B. Tuttle, E. L. Cattau, and P. Wang, Comp. Biol. Physiol. 49B, 687 (1974). 281. K. V. Rajagopalan and P. Handler, JBC 239, 1509 (1964). 282. K. V. Rajagopalan, I. Fridovich, and P. Handler, JBC 237, 922 (1962). 283. K. V. Rajagopalan, P. Handler, G. Palmer, and H. Beinert, JBC 243, 3784 (1968). 284. T. Nishino, BBA 341, 93 (1974). Exp. Biol. 11-12 May 1974. 285. R. Andres, Abstr., 6th Annu. Meet. Union Swiss SOC. 286. H. Dalton, D. J. Lowe, R. T. Pawlik, and R. C. Bray, BJ tin press]. 287. V. Aleman, S. T. Smith, K. V. Rajagopalan, and P. Handler, in “Flavins and Flavoproteins” (E. C. Slater, ed.), p. 99. Elsevier, Amsterdam, 1966.

w

CD

0

TABLE X PURITY,MOLECULAR PROPERTIES, AND COMPOSITION OF SOMEMOLYBDENUM HYDROXYLASES" Enzyme ~

Aldehyde oxidase

Rabbit liver

Aldehyde oxidase

Pig liver

Xanthine behydrogenase Xanthine dehydrogenase Xanthine dehydrogenase Xanthine dehydrogenase Xanthine dehydrogenase

Chicken liver

69;63 (77c,281) -

Turkey liver

D. melanogmter V . alcalescens (176) C . cglindrosporum

F n

P

TABLE X (Continued)

E

E P R signals Molybdenum Analysis (mole/mole FAD) Enzyme Aldehyde ox. (rabbit) Aldehyde ox. (pig) Xanthine deh. (chicken) Xanthine deh. (turkey) Xanthine deh. (D.mel.) Xanthine deh. ( V . alc.) Xanthine deh. (C. cyl)

Fe

Labile S

Mo

Resting signals"

Very rapid typed

Rapid typee

Slow type'

Inhibited type0

FADH' Yesk ($83)

3.90 (179)

Fe/S Yesz (283) -

-

Yes (90b)

31

3

2 %

B

Yes (75) Yes (90b)

Yes (886,287)

Data in brackets are to be regarded aa relatively unreliable, usually because they were stated t o be approximate or because experimental details are not given. References are given in parentheses. * An important practical point relating to molecular weight determinations is the finding (169,179,284) that some of the enzymes tend to aggregate unless thiols are present. Here defined as signals, present in the resting (oxidized) enzyme, distinct from the other Mo signals, and which do not change on adding a reducing substrate within the enzymc's turnover time. d Here defined as a signal appearing at very short reaction times with xanthine, without proton splitting, and with 911 > gl.

w

E

eHere defined as a signal appearing in the presence of substrate with indications of proton splitting and 911 > gl. Here defined as a signal obtained under reducing conditions with indications of proton splitting and g1 > 911. 0 Here defined as a signal developing on treatment with MeOH or HCHO, with indications of proton splitting. From ultracentrifuge measurements. i From gel diffusion or filtration measurements. j Early work (282) indicated that this enzyme also contained coenzyme Q. T his seems to have been neither confirmed nor contradicted in later studies. However, since no functional role for this coenzyme has been proposed it seems likely that it w&sa contaminant. I: The observed linewidth was 16 G. T his low value might indicate the presence of coenzyme Q radicals rather than the anionic form of FADH. 2 One species only detected, having nearly axial symmetry. m At 460 nm. Linewidth about 19 G. 0 Two species, each with rhombic symmetry. f

w

(0

N

m 9

6. MOLYBDENUM HYDROXYLASES

393

sponding data on milk xanthine oxidase in Table I1 shows the extreme similarities among the enzymes, particularly in such properties as molecular weights, metal and FAD contents, and extinction coefficients and their ratios. Although the data for individual enzymes generally agree with a composition of 1 Mo, 1 FAD, and 4 Fe/S per half-molecule, as in the milk enzyme, there are strong indications in the low molybdenum contents of some preparations of the rabbit liver enzyme (and perhaps also of the enzyme from Clostridium), that occurrence of demolybdo forms, presumably as natural products, is not unique to milk xanthine oxidase ( 1 8 7 ~ )It. is well established (76b) that the turkey liver enzyme is normally contaminated with a desulfo form, which gives rise to a Slow Mo (V) EPR signal ( g o b ) , As with the milk enzyme, the active turkey enzyme is converted (75b) to the inactive desulfo form on treatment with cyanide. It seems certain that similarly all molybdenum hydroxylases must contain the essential cyanide-sensitive persulfide group. This has indeed been established for aldehyde oxidase (96b),while many of the enzymes have been reported (e.g., 96,103,358) to be sensitive to cyanide. Furthermore, although the enzyme from Veillonella has been stated (176) to be relatively insensitive to this reagent, preparations of it do give a very clear Slow signal (186), thus showing unambiguously the presence of its desulfo form. Future workers on any molybdenum hydroxylase should therefore be on guard against being confused by the almost certain presence in their preparations of the inactive desulfo form, as well the possible presence of the demolybdo form. Despite the obvious similarities, it is possible, however, even at the present limited state of knowledge of some of the molybdenum hydroxylases of Table X, to point to significant differences among some molecular properties. Thus, spectra in the visible region are not all quantitatively the same. Of particular significance may be the fact that, whereas milk xanthine oxidase has E580/E450 = 0.31 to 0.32 (36,63,281), all the enzymes in Table X have higher values for this ratio. Similarly, for rabbit aldehyde oxidase the value of 63-69 for the 450 nm extinction coefficient seems significantly lower than that for the milk enzyme, which is 72 287s. Felsted et al. (179) suggested that the low molybdenum content of their preparations of aldehyde oxidase may have resulted from partial loss of the metal during purification. They gave no direct evidence for this, citing in support only early work by Mahler et al. (287b), which claimed that molybdenum could readily be removed from this enzyme. However, they were apparently unaware that similar claims by the same group relating to molybdenum removal from xanthine oxidase were not substantiated in later work (@; see Section II,B,l,b). 287b. H. R. Mahler, B. Mackler, D. E. Green, and R. M. Bock, JBC 210, 465 (1954).

394

R. C. RRAY

(Table 11). No doubt these variations reflect differences in the precise structure of the iron-sulfur chromophores of the different enzymes. This is more strikingly reflected in the parameters of the Fe/S EPR signals, where these have been studied (288). Whereas the milk and the turkey enzymes both have two types of Fe/S centers, each giving rise to a rhombic g tensor, rabbit aldehyde oxidase (283) and xanthine dehydrogenase from Veillonella (2867, on the other hand, each contains apparently, only one type of Fe/S center, giving an almost axial g tensor. Further, the g values of the two Fe/S systems in the turkey enzyme (90b) are not precisely the same as those of Fe/S I and Fe/S I1 of milk xanthine oxidase (Table V). Similarly, the parameters of the Fe/S in aldehyde oxidase (283) are not identical with those of this center in the Veillonella enzyme (286). With regard to the EPR parameters of Aio(V) and FADH., differences among the enzymes seem small. The only unambiguous linewidth available for FADH’ signals, for the enzymes from Veillonella alcalescens and from turkey liver, indicates that the radical must, like that for the milk . is consistent with the observaenzyme, be of the “blue” type ( 9 1 ~ )This tion of increases in light absorption at long wavelengths on partial reduction by substrates (169,175). Such increases have also been noted for the chicken liver enzyme (76,105). Hence, it seems probable that the semiquinone will prove to be “blue” in all these enzymes. Detailed discussion of data in Table X on the molybdenum EPR signals will be reserved for the next section, since it is particularly relevent to the mechanism of reduction of the enzymes.

C. CATALYTIC PROPERTIES 1. Mechanism of Reduction of the Enzymes The specificities of various molybdenum hydroxylases toward reducing substrates were considered in detail in Section II,C,l,a. This section will summarize available information, derived for inhibition and from EPR studies, on the mechanism of interaction of reducing substrates with the various enzymes. It seems clear that this interaction must in all cases be very similar to that which obtains in milk xanthine oxidase. Electron paramagnetic resonance data on molybdenum signals are summarized in Table X. We first consider the resting Mo (V) signals from the enzymes, ie., signals which are present in the enzymes as they are normally prepared and when apparently in the fully oxidized state. Such 288. R. C. Bray, M. J. Barber, H. Dalton, D. J. Lowe, and M. P. Coughlan, Biochem. SOC. Trans. (in press).

6.

MOLYBDENUM HYDROXYLASES

395

signals have been known for some while for aldehyde oxidase (283) and for the enzyme from Veillonella (287; see also 286) and have had attributed to them considerable significance in the enzymic mechanisms (162,287,289). The resting signals from these two enzymes are quite distinct for one another and each could well result from single chemical species. Neither signal bears any close resemblance to any Mo(V) signal from milk xanthine oxidase, as listed in Table 111. More recently, turkey xanthine dehydrogenase has been studied (90b) and found to give two resting signals, one like the aldehyde oxidase signal and the other rather like that from Veillonella ( 9 1 ~ )Furthermore, . in additional experiments in the author’s laboratory, a resting signal, like that from aldehyde oxidase, has also been obtained, under special conditions, from milk xanthine oxidase samples. It seems clear that the resting signals are not derived from functional molybdenum hydroxylase molecules. Evidence for this is clearest in the case of the turkey enzyme ( 9 0 b ) . Neither of its resting signals changed on adding oxidizing agents and only one changed, and then not readily, in the presence of reducing agents. Most importantly, in rapid freezing experiments, the resting signals remained unchanged in intensity when the active enzyme was reduced with xanthine (see below). It is also significant that one of the resting signals (the Veillonella-like one) could be eliminated from the turkey enzyme by a n appropriate pretreatment of the sample. This treatment consisted simply of putting the enzyme through a cycle of reduction by dithionite and reoxidation by oxygen. Comparable, although less detailed results were obtained with the Veillonella enzyme (286). Here, too, although the resting signal ultimately disappeared on adding reducing substrates, i t did not do so within the turnover time. The precise significance of molybdenum hydroxylase resting signals is not certain and the nature of the modifications in the active centers which are presumed to give rise to these signals remain obscure. It is also not clear whether the modifying reactions can be reversed under any condition. Nevertheless, i t is clear that resting signals cannot result from functional enzyme molecules and that such signals may therefore safely be ignored when turnover reactions of the enzymes are considered. This conclusion must apply t o aldehyde oxidase as well as to the other two enzymes, despite earlier indications to the contrary (90b,9la,162). Reported changes in EPR signals, occurring on adding reducing substrates to molybdenum hydroxylases, after appropriate correction for resting signals is made, seem to be entirely comparable to those which are obtained with milk xanthine oxidase. This conclusion is most appar289. P. Handler, in “FIavins and Flavoproteins” (H. Kamin, ed.), p. 444. Univ. Park Press, Baltimore, Maryland, 1971.

396

R. C. BRAY

ent from work (90b) on the turkey enzyme. A Very Rapid and a Rapid signal develop within the turnover time on adding xanthine to this enzyme under appropriate conditions. Furthermore, in titration studies with this substrate the Rapid signal first increased as the amount of xanthine was increased, then decreased again, corresponding to reduction of molybdenum to the four-valent state. For the Veillonella enzyme, a Rapid signal was also obtained (286) in a time comparable to the turnover. Similarly, although rapid freezing studies have not been reported, the chicken enzyme also gives (75) a very clear Rapid signal, while for aldehyde oxidase, a signal which is probably of the Rapid type is obtained (283). Detailed rapid freezing kinetic studies have been reported only for aldehyde oxidase (162). Unfortunately, however, this work was carried out a t a time when desulfo forms of the enzymes (not to mention resting signal-giving species) were not understood and when the various possible Mo(V) signal types had not been distinguished. However, the work did indicate (162) that as steady-state conditions were approached, Mo (V) and Fe/S signals appeared, with similar time-courses to one another. Similar results relating to signal disappearance were obtained in experiments in which reoxidation of enzyme, reduced by brief exposure to substrates, was studied. On the whole, the behavior of aldehyde oxidase in reductive titration experiments (283) also appears comparable to that of the milk and turkey enzymes, although it seems to have been difficult to reduce molybdenum to the four-valent state. However, it may well be that changes in the active enzyme were partly masked by the presence, which we can safely assume in these experiments, of desulfo enzyme. Other molybdenum signals to be considered are the Inhibited and the Slow signals. Slow signals from desulfo forms of the enzymes from turkey and from Veillonella were mentioned in the previous subsection. The Inhibited signal arises, in the case of the milk enzyme, on inactivation by methanol ( 2 8 9 ~ or ) by formaldehyde. In keeping with this, aldehyde oxidase gives a very clear Inhibited signal on treatment with MeOH (283),while for the turkey enzyme one was obtained with HCHO (90b). Other enzymes do not seem to have been examined for this signal, although sensitivity of several of them to methanol (103,175) suggests that it ought to be observable in other cases also. A striking point about the molybdenum E P R work on these enzymes is the extreme similarity of the parameters of corresponding signals from different enzymes. Thus, none of the g vaues for the Very Rapid, Rapid, 289a. Loss of activity of molybdenum hydroxylases on treatment with methanol frequently parallels that occurring in the presence of arsenite (cf. SG). Effects of both these reagents are discussed in Section II,B.3,f.

6.

MOLYBDENUM HYDROXYLASES

397

Slow, or Inhibited signals from the turkey enzyme (90b) differs by more than 0.002 from the corresponding figure for the milk enzyme. For some of the other enzymes, although significant differences from these two enzymes are observed (e.g., in the parameters of the Inhibited signal from aldehyde oxidase) , they are, nevertheless, relatively small differences which are involved. The EPR data therefore make it virtually certain that the ligands of molybdenum are the same in all molybdenum hydroxylases. There must, however, be substantial differences in other groups in the substrate binding sites to account for specificity differences (Section II,C,l,a), as well as, for example, for differences in sensitivity to inhibition by allopurinol ( 179,290). When all the above data axe considered in conjunction with indications (Section II,C,2,c) that at least under some conditions ping-pong kinetics apply to all the molybdenum hydroxylases which have been tested, then there can be little doubt that all these enzymes are indeed quite similar to one another. 2. Oxidizing Substrates

We now turn to the oxidizing substrates of the purified molybdenum hydroxylases. Data on relative turnover rates of some of the enzymes, with five different acceptors, are summarized in Table XI (10,49,lOS,l68, 169,171,175,~80,68~,~91,291 a ) . Clearly, there are wide differences of acceptor specificity among the enzymes. However, all enzymes listed utilize 2,6-dichlorophenolindophenolquite effectively. Every enzyme has some activity toward oxygen, although in most cases the oxidase activity is low. The exceptions are the mammalian enzymes where oxidase activity is high (292). Similarly, high activity toward NAD+(perhaps accompanied by low activity to ferricyanide) is found only in the avian enzymes. Finally, in the bacterial enzymes, activity both to oxygen and to NAD+ is low and ferredoxin is assumed to be the natural acceptor (293). According to Table XI, then, acceptor specificities divide the molybdenum hydroxylases into three clearly defined groups : oxidases, NAD+ dehydrogenases, and other dehydrogenases. (We have not included cytochrome c as an acceptor in Table XI, since, for many of the enzymes, the question of whether its interaction occurs directly, via 02-,or by both routes, does not seem fully resolved.) 290. N. W. De Lapp and J. R. Fisher, BBA 269,505 (1972). 291. D. B. Morell, BBA 18, 221 (19551. 295a. C. F. Strittmatter, JBC 240, 2557 (1965). 292. See also Section II,B,3,d. 293. Although ferredoxin is a good acceptor for the enzyme from V e d b n e l h it does not, surprisingly, seem actually to have been tested on any of the other enzymes, bacterial or otherwise.

TABLE X I OXIDIZINQ SUBSTRATES OF SOME MOLYBDENUM HYDROXYLASES Relative rateo of reaction with Enzyme Xanthine oxidase

Source

Oxygen

NAD+

Ferredoxin

Milk (cow)

Ferricyanide 100 (10)

Aldehyde oxidase

Rabbit liver

Xanthine dehydrogenase

Chicken liver

2,11 (103,

100 (280)

-

280) Xanthine dehydogenase

Turkey liver

Xanthine dehydrogenase

V. akalescens (17 6 )

Xanthine dehydrogenase

C. cylindrosporum

12 (175)

0 ( 175)

1 (168)

0 (168)

Indophenol 35 (49 1

4,77 (291a, 280)

100 (103)

100

90

( 1 76)

( 1 75)

28 (175)

-

100 (168)

36 (168)

Rates with xanthine as reducing substrate are given unless otherwise stated, the fastest acceptor for each enzyme being assigned a value of 100. Conditions of the memurements varied. References are given in parentheses. Various reducing substrates (other than xanthine) were used.

*I ?

e

6.

MOLYBDENUM HYDROXYLASES

399

Why differing acceptor specificities are shown must now be regarded as one of the major unsolved problems concerning the molybdenum hydroxylases and we can do little more than speculate about its possible mechanism here. Clearly, differences in the spectroscopic properties of the Fe/S centers do not correlate with specificity differences. Thus, insofar as Fe/S E P R is concerned, we have one oxidase (from milk), resembling an NAD’ dehydrogenase (from turkey), while the other oxidase (from rabbit) resembles not these two enzymes but instead an “other” dehydrogenase (from Veillonella) (288).Similarly, available data on the properties of flavin in the enzymes do not help greatly in explaining the acceptor specificities. The flavin in xanthine oxidase, and apparently in the other enzymes also, is of the type giving rise to a “blue” semiquinone radical. FlavoprotGins of this type are not generally good oxidases (294). Thus, in this sense, it is the oxidases which have to be regarded as exceptional among molybdenum hydroxylases. A small clue, however, that flavin might be involved in the differences between the enzymes has been available for some while. Thus, there are indications that partial reduction by substrates [e.g., for the chicken enzyme (IOS)]gives rise to considerably more flavin semiquinone than does the milk enzyme under comparable conditions. This has been put on a more quantitative basis in more recent work on the turkey enzyme (9Ob19Ia).It was found in titration studies that reduction of flavin to FADH, was difficult to achieve so that large amounts of FADH. accumulated. This would seem t o explain low activity to oxygen in this and possibly other dehydrogenases, since at least for free flavins (201,271), the fully reduced form of the coenzyme is more reactive to oxygen than is the LLblue” semiquinone. Perhaps, therefore, it is reasonable to assume that the origins of acceptor specificity differences lie in subtle variations in the environments of the flavin molecules in the different enzymes. These might be related, e.g., to the proximity of thiol groups (cf. Section II,B,3,d) and, in any case, would presumably be reflected both in differences in NAD’ binding and in variations in the flavin redox potentials. The mechanism proposed by Olson et al. (46; Section 111C,2,i) for reoxidation of milk xanthine oxidase postulates that the role of the Fe/S systems is to maximize formation of FADH,, this being required for efficient two-electron reduction of oxygen. On the basis of such a scheme, an enzyme which reacted rapidly with 0,, ought also, provided a suitable binding site were available, to react efficiently with NAD’, and vice versa. Perhaps, however, NAD’ binding sites are lacking in the oxidases, whereas in the dehydrogenases, the redox potentials are not suitable for oxygen reduction to 294. V. Massey, F. Miiller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G . Howell, S. G . Mayhew, R. G. Matthews, and G . P. Foust, JBC 244,3999 (1969).

400

R. C. BRAY

take place via two one-electron steps. [See Sections II,C,l,c and II,C,2,g; the redox potential (204) for the system O,/O,- is slightly lower than that for NAD+/NADH.] Clearly, however, further work on the acceptor specificity differences is called for.

IV. Genetic Studies and the Molybdenum Hydroxylases

A. INTRODUCTION There is now in the literature a substantial body of work, of a more or less genetic nature, involving various molybdenum hydroxylases. By far the most extensive section of this concerns the enzymes in Drusophila melanogaster. There have also been considerable studies on the enzymes in Aspergillus nidu1an.s and, further, there is some related work on humans, which we have to consider. Finally, studies on the enzyme nitrate reductase in Neurospora crassa mutants have a bearing on the composition of the molybdenum hydroxylases. In much of the work we shall consider in this section, there has, as yet, been relatively little interaction between genetics and molecular enzymology. One hopes that such interaction will increase to the benefit of both sides.

B. XANTHINURIA AND GOUT IN MAN The study of purine metabolism in man is an extensive field (see, e.g., 294a) ; this section is concerned only with the molybdenum hydroxylases

involved in the hydroxylation of the naturally occurring purines, hypoxanthine and xanthine, and of the drug, allopurinol, and of genetically determined changes in these enzymes. (Allopurinol is a potent inhibitor of milk xanthine oxidase; see Section II,C,l,g.) Xanthinuria, a rare genetically determined condition in which xanthine oxidase activity is lacking from individual humans, was first described by Dent and Philpot in 1954 (295), although the clinical manifestation which sometimes accompanies it, namely, appearance of xanthine stones, had been known since 1817. Only 18 cases of xanthinuria had been re294a. 0. Sperling, A. de Vries, and J. B. Wyngaarden, eds., "Purine Metabolism in Man," Advun. Exp. M e d . Biol., Vols. 41A and 41B. Plenum, New York, 1974. 295. C. E. Dent and G. R. Philpot, Lancet 1, 182 (1954).

6.

MOLYBDENUM HYDROXYLASES

401

ported up to 1974 ( 2 9 5 ~ ) As . would be expected if xanthine oxdase is completely absent, Engelman et al. (296) found that in biopsy samples from a xanthinuric patient both xanthine and hypoxanthine were oxidized, in zn'tro, a t rates of less than 0.1%of those from normal controls. On the other hand, the same patient as was used in this work is stated (297,298) to have been capable, in vivo, of converting administered hypoxanthine to xanthine, although not to uric acid. It was assumed that this oxidation had occurred after conversion of hypoxanthine t o the nucleotide, oxidation then being followed by hydrolysis, back to free xanthine. Further information on xanthinuric patients has been obtained from studies of the effects of allopurinol administration on their purine metabolism (see 295a). Five such patients have now been examined in this way by various workers, and results reveal surprising differences among the patients. Xanthinurics normally excrete most of their urinary purines as xanthine, with relatively little in the form of hypoxanthine. In some cases, allopurinol administration had no effect on this pattern, while in others, the ratio of these metabolites was reversed by the drug so that hypoxanthine became the major excretory product. Furthermore, some of the patients metabolized allopurinol to alloxanthine, whereas others did not do so, and ability to oxidize the drug did not correlate with its effects on xanthine excretion. It seems that the five patients must be divided, on the basis of these results, into a t least three different groups. We will return to possible explanations of these findings below, after mentioning briefly some data relating to gout. Allopurinol is now widely used in the treatment of gout and hyperuricemia, as mentioned in Section I,B. During such treatment, excretion of uric acid is decreased, while that of xanthine and hypoxanthine is increased (12,298) , as would be expected for inhibition of xanthine oxidase activity (cf. 298a). An unexpected finding (299) , during allopurinol treatment of one gouty patient on a low-purine diet, however, was that some 6,8-dihydroxypurine, a substance not previously reported in human urine, was excreted. 295a. H. A. Simmonds, B. Levin, and J. C. Cameron, Clin. Sci. Mol. Med. 47, 173 (1974). 296. R. Engelman, R. W. E Watts, J. R. Klinenberg, A. Sjoerdsma, and J. E. Seegmiller, Amer. J. Med. 37, 839 (1964). 297. A. Kovensky, G. H. Etchings, E. Metz, and W. R. Rundles, Biochem. Pharmacol. 15, 863 (1966). 298. H. A. Simmonds, Clin. Chim. Acta 23, 353 (1969). 298a. M. M. Jezewska, Eur. J. Biochem. 46, 361 (1974). 299. H. A. Simmonds and W. Sneddon, Clin. Chim. Acta 30, 421 (1970).

402

R. C. BRAY

It is suggested (cf. W96a,W99) that the data on the xanthinurics and the gouty patient may be best explained by the presence in humans, generally, of several different molybdenum hydroxylases, each having its own specificity pattern. Two such enzymes (xanthine oxidase and aldehyde oxidase) have been reported in normal human tissues (96,679) ; and one might even imagine that, a t least in some subjects, a third enzyme could exist, either in addition to the other two, or perhaps in place of one of them. Thus, to extend this idea, a few individuals might possess a genetically determined variant of one or other of the normal molybdenum hydroxylases having a modified substrate binding site and therefore a new specificity pattern. On such a basis, the finding, that xanthinurics although apparently lacking one molybdenum hydroxylase still excrete xanthine rather than hypoxanthine and seem able in vivo to convert the latter to the former, would be explained as resulting from the presence of a molybdenum hydroxylase with properties reminiscent of those of rabbit aldehyde oxidase (10).Thus, this postulated enzyme would oxidize hypoxanthine to xanthine, but not xanthine €0 uric acid and, like the rabbit enzyme, would be insensitive to allopurinol. I n some xanthinurics a modified molybdenum hydroxylase, sensitive to the drug, but still not acting on xanthine, would be present. Similarly, genetically determined modifications in the active centers of molybdenum hydroxylases might account for the failure of some xanthinurics to oxidize allopurinol and might also account, in the gouty subject, for excretion of 2,8-dihydroxypurine. Obviously, there are a number of different hypotheses along the above lines which are capable of explaining the data, and further work is required. However, one point which has now become clear (cf. W96a) must be mentioned: Oxidation of allopurinol to alloxanthine in xanthinurics apparently does not take place, as was originally suggested (300),a t the nucleotide level. C. THE“COMMON COFACTOR” OF NITRATE REDTJCTASE AND DENUM HYDROXYLASES

THE

MOLYB-

In 1964, Pateman et al. (Sol),working on mutants of the fungus Aspergillus nidulans, proposed that there was a common cofactor, given in the name cnx, which was essential for both nitrate reductase and xanthine dehydrogenase activities in this organism. The suggestion was made that 300. R. .A. Chalmers, R. Parker, H. A. Simmonds, W. Snedden, R. W. E. Watts, BJ 112, 527 (1969). 301. J. A. Pateman, D. J. Cove, B. M. Rever, and D. B. Roberts, Nature (London) 201, 58 (1984).

6.

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403

cnx might be somehow associated with molybdenum. Work on the Aspergillus mutants has continued and will be taken up again in Sectioii IV,E. Before considering it, however, we will discuss extensive studies by Nason and co-workers, bearing directly on the nature of what would be presumed to be a very closely related “common cofactor,” from another fungus, Neurospora crassa. Nitrate reductase in Neurospora mama is inducible by nitrate and in 1970, Nason et al. (302), working with mutant strains deficient in the enzyme, known as nit-1, nit-2, and nit-3, reported on intercistronic complementation phenomena, observed in vitro, using cell-free preparations. They found that incubation of induced nit-1 with either uninduced wildtype extracts or, alternatively, with extracts from induced or uninduced nit-2 or nit-3, produced NADPH-nitrate reductase activity. The enzyme so produced had sedimentation behavior = 7.9) indistinguishable from that of the wild-type enzyme. Nitrate reductase activity in this organism is always associated with NADPH-cytochrome c reductase activity. The latter activity was also found on its own, specifically in induced but not in uninduced nit-1 extracts. However, there it had a lower sedimentation coefficient, with szo,w= 4.5. It was therefore assumed that the observed complementation, giving nitrate reductase, resulted from interaction of two or more protein subunits coded by different cistrons, one of these proteins (with szo,w= 4.5) being present only in induced nit-1 and the other being present in uninduced wild-type and other mutant strains. The next, somewhat surprising finding (30.9)was that, in this complementation phenomenon, the second protein component, i.e., the one in nit-2, etc., could be replaced by any of several purified molybdenum hydroxylases, provided these had been pretreated by exposure to a pH of about 2.5. Although the molybdenum hydroxylases employed came from higher animals, it was nevertheless assumed that they were here supplying a protein subunit capable of replacing, both structurally and functionally, the natural one in the fungal nitrate reductase. The complementing species from acid-treated milk xanthine oxidase appeared to have s20,w about 6 and was suggested to be a subunit, half the size of native xanthine oxidase. Complementing activity was reported to be unstable, even a t Oo and neutral pH values. The above somewhat unlikely hypothesis became quite untenable, a year later, when it was found that the replacement of the nitrate reduc302. A. Nason, A. D. Antoine, P. A. Ketchum, W. A. Frazier, and K.-Y. Lee, Proc. Nat. Acad. Sci. U . S. 65, 137 (1970). 303. P. A. Ketchum, H. Y. Cambier, W. A. Frazier, C . H. Mandansky, and A. Nason, Proc. Nat. Acad. Sci. U . S. 66, 1016 (1970).

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tase component by acid-treated molybdenum hydroxylases could be extended to include a large number of other acid-treated molybdenum enzymes (304).Although every molybdenum-containing enzyme tested was active in generating nitrate reductase activity from induced nit-1 , neither molybdate, various molybdenum complexes, nor molybdenum-free enzymes could substitute for them. It was therefore proposed that the molybdenum enzymes were supplying a relatively small “molybdenum cofactor” to the nitrate reductase protein. The various acid-treated proteins were supposed to be acting simply as carriers for this cofactor, while the molecular weight change on complementation (302)was rationalized (304) as resulting from association of protein subunits, present in the nit-1 extracts, under the influence of the cofactor. This interpretation seems a reasonable one and is generally supported by data, to be considered later, on Aspergillus. Further information on the nature of the cofactor is, however, awaited with the utmost interest. Nason et al. (306) claimed to have isolated the cofactor and reported that it had a molecular weight of 1000 or less but gave no details and no indications of its possible nature. Additional work by the same group (306) provided more direct information on the role of molybdenum. When induced nit-1 was converted to nitrate reductase, by treatment with extracts from uninduced wild-type cells grown on Q Q Mthen ~ , nitrate reductase activity and radioactivity moved together during ultracentrifugation on sucrose density gradients. When acid-treated xanthine oxidase was used as the source of the activating material, addition of molybdate stimulated nitrate reductase formation. Stimulation was particularly marked if the xanthine oxidase had been allowed to stand in the acid medium, thus, it was presumed, losing molybdenum. The presence of the xanthine oxidase was, however, essential, and high concentrations of molybdate M ) were required. If the added molybdate was radioactive, then it was incorporated into the nitrate reductase. Molybdenum, once in nitrate reductase was, however, nonexchangeable. The final conclusion (306) was that most probably the nit-1 gene product is a “structural component of the enzymes, more specifically the cofactor moiety which interacts with molybdate.” If this conclusion, and the conclusion that the cofactor has a molecular weight of less than 1000, are indeed correct, then the nit-1 gene product must be a most abnormally 304. A. Nason, K.-Y. Lee, S.4. Pan, P. A. Xetchum, A. Lamberti, and J. de Vnes, Proc. N a t . Acad. Sci. U.S . 68, 3242 (1971). 305. A. Nason, K.-Y. Lee, S.-S. Pan, and R. H. Erickson, in “Chemistry and Uses of Molybdenum” (P. C. H. Mitchell, ed.), p. 233. Climax Molybdenum Co., London, 1973. 306. K.-Y. Lee, S.S. Pan, R. Erickson, and A. Nason, JBC 249, 3941 (1974).

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small one. Gene products are normally thought of as being polypeptides containing not less than, say, 40 amino acid residues. However, other indirect roles for the gene product were not excluded (306),e.g., in transporting, reducing, or helping to incorporate molybdate into the nitrate reductase protein. At this stage we can only speculate as to the possible nature of the low molecular weight cofactor (if such it is) , which is implicated in this work. It might be either organic or inorganic (305). One possibility, which has not, apparently, been suggested hitherto, is that it could be related to the persulfide group of xanthine oxidase and other molybdenum hydroxylases. Although there is of course no positive evidence for such a group in molybdenum enzymes other than hydroxylases, its presence does not seem to have been specifically excluded (cf. Section V) . However, if such a group did turn out to be present, then the specific role postulated for persulfide in xanthine oxidase catalysis [Section 11,CJ2,f; Fig. 6; ( 4 6 ) ] would obviously have to be ascribed to some different grouping in the enzyme’s active center. It may be reasonable to regard the material produced by induced nit-1 cells as an analog of demolybdo xanthine oxidase (Section II,B,l,b). However, the molecular weight difference between the former and nitrate reductase itself seems a point of difference, since demolybdo xanthine oxidase has the same molecular weight as the active enzyme. We now return to work on Aspergillus nidulans, and in particular to that by Cove and co-workers. Their studies (307) on the sedimentation coefficients of nitrate reductase and of NADPH-cytochrome c reductase in mutant strains of this fungus are consistent with the ideas of Nason et al. (30.4) on the origins of nitrate reductase in Neurospora, as mentioned above. It was reported (307), that in wild-type Aspergillus, nitrate reductase had 7.6 S and was associated with cytochrome c reductase. I n mutants lacking nitrate reductase there was, on the other hand, a new species of cytochrome c reductase, with 4.5 S. It was proposed that nitrate reductase is made up from two of these 4.5 S subunits, which associate under the influence of a cofactor specified by the cnx gene, and that the cofactor has to remain bound in the dimer for the nitrate reductase activity to be observed. I n apparent contrast to the work on Neurospora, it was concluded, however, that the Aspergillus cofactor might have a molecular weight as high as 20,000. There are five different genes involved in the cnx mutants of Aspergillus, and in all such mutants both xanthine dehydrogenase and nitrate 307. D. W. MacDonald, D. J. Cove, and A. Coddington, Mol. Gen. Genet. 128, 187 (1974).

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reductase activities are deficient (see 307a). Study of some of these mutants has provided more information bearing on the nature of the common cofactor. Of particular interest are the cnx E and cnx H mutants. Addition of molybdate to the growth medium has been reported (307b) to repair, a t least partially, the cnx E mutation, restoring nitrate reductase activity, together with some xanthine dehydrogenase activity. This would seem to imply that the cnx E gene specifies for something involved in incorporating molybdenum into these enzymes and not for a part of the enzymes themselves. On the other hand, cnx H seems to specify for a structural component of nitrate reductase. Cnx H mutants are temperature sensitive and when the properties of the nitrate reductase in them were compared with those of this enzyme from wild-type cells, i t was found (307~)that not only were the cells affected by temperature but also the enzyme itself, in the mutant, had a diminished thermal stability in vitro. This was taken to imply that cnx H specifies for a part of the nitrate reductase protein. After combining the Neurospora and the Aspergillm data it is not clear whether the common piece of the molybdenum enzymes, indicated by the genetic work, is large or small. I n fact, although one would have expected a similar situation in the two organisms, there appears to be something of a contradiction between the two sets of data (but see 307a). Whereas results on Neurospora appear to exclude a protein component in the ((common cofactor,” some of those on Aspergillus seem to necessitate one. How this will be resolved remains to be seen. Perhaps in any case the closest analogy should be reserved for comparison of nit-I with cnx E. We shall return to the question of the structure of Aspergillus xanthine dehydrogenase in Section IV,E.

D. MOLYBDENUM HYDROXYLASES IN Drosophila melanogaster As has already been mentioned, work on molybdenum hydroxylases in Drosophila melanogaster during the past 20 years or so has been quite extensive. Most work has been concerned, primarily, with understanding the genetics of the orgadism but here we shall concentrate on those aspects seeming to have a direct bearing on the structure and functioning of the enzymes. Much relevant work has been reviewed by Finnerty (307d). 307a. C. Scazzocchio, J . Leas-Common Metals 36, 461 (1974). 307b. H. N. Arst, D. W. MacDonald, and D. J. Cove, Mol. Gen. Genet. 108, 129 (1970). 307c. D. W. MacDonald and D. J. Cove, Eur. J . Biochem. 47, 110 (1974). 307d. V. Finnerty, in “The Biology and Genetics of Drosophila” (E.Novitski and M. Ashburner, eds.), Vol. 1. Academic Press,New York, 1974.

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It now appears to be accepted by geneticists (68,307d) that in Drosophila, three related enzymes, given the names “xanthine dehydrogenase,” “aldehyde oxidase,” and “pyridoxal oxidase,” are subject to the genetic controls described below (307e).The levels of all three enzymes are controlled by two loci, maroon-like (308) (ma-2, 1-64.8),on the X chromosome and low xanthine dehydrogenase (ha, 3-33),on the third chromosome. Three other loci each affect one of the enzymes only and are closely grouped on the third chromosome. Of those, rosy (308) (ry, 3-52.0)is the structural gene for xanthine dehydrogenase (309),aldehyde oxidase (aldoz, 3-56.6f 0.7) is the structural gene for aldehyde oxidase (310), while low pyridoxal oxidase (Ipo, 3-57) has been suggested (311) but not established (307’d)to be the structural gene for pyridoxal oxidase. The first question to consider is whether these enzymes are molybdenum hydroxylases or not and whether, indeed, there is good evidence for there being three separate enzymes. Xanthine dehydrogenase seems fairly well authenticated as a molybdenum hydroxylase (see Table X ) . It has been purified to apparent homogeneity from Drosophila (68), although few properties have been reported as yet (67c,286). An indication that its specificity to reducing substrates (cf. 312,313) may be comparably wide to that of other molybdenum hydroxylases is that a standard assay (314) uses 2-amino-4-hydroxypteridineas substrate. However, activity toward at least some aldehydes seems relatively low (316,516). At least partial separations of aldehyde oxidase from xanthine dehydrogenase have been achieved by various methods (316,316).Therefore, taking into consideration the genetic as well as the separation evidence, there seems little doubt about separate existence of these two enzymes in Drosophila. However, little is known of the properties of aldehyde oxidase. The partially purified enzyme (310,516) has been studied in a limited way, but there is no positive evidence that activity toward 307e. An additional locus, ein, has recently been shown by B. S. Baker [Develop. Biol. 33, 429 (1973) I, to be involved in the regulation of xanthine dehydrogenase. 308. The names “maroon-like” and “rosy” derive from the eye colors of the flies.

Apparently these colors are determined by the presence of certain pteridines, whose levels are related to and, a t least to some extent, controlled by those of the molybdenum hydroxylases. 309. T. T. Yen and E. Glassman, Genetics 52,977 (1965). 310. W. J. Dickinson, Genetics 66, 487 (1970). 311. J. F. Collins and E. Glassman, Genetics 81,833 (1969). 312. E. Glassman and H. K. Mitchell, Genetics 44, 153 (1959). 313. S. D. Parzen and A. S. Fox, BBA 92, 465 (1964). 314. E. Glassman, Science 137, 990 (1962). 315. J. B. Courtright, Genetics 57, 25 (1967). 316. J. F. Collins, E. J. Duke, and E. Glassman, Biochem. Genet. 5, 1 (1971).

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reducing substrates extends beyond aldehydes. Indeed, although genetic evidence for a close relationship between this enzyme and xanthine dehydrogenase is no doubt adequate to justify the assumption that aldehyde oxidase is a molybdenum hydroxylase, there has yet been no direct biochemical confirmation. The only known property we can fall back on for biochemical support is an indication (315)that the molecular weight of aldehyde oxidase is of the expected order of magnitude for an enzyme of this class. Regarding pyridoxal oxidase, we have even less information available. Pyridoxal is a competitive inhibitor of Drosophila xanthine dehydrogenase (317). Furthermore, according to Krenitsky et al. (lo),both milk xanthine oxidase and rabbit aldehyde oxidase can oxidize pyridoxal a t quite significant rates. Therefore, bearing in mind the common genetic controls of pyridoxal oxidase and xanthine dehydrogenase in Drosophila, it may be reasonable t o assume that it, too, is a molybdenum hydroxylase. While the r y and aldox genes are quite well separated, lpo, a t 3-57, is not clearly different from aldox, a t 3-56.6 & 0.7. Thus, one might have suspected that pyridoxal oxidase and aldehyde oxidase are one and the same enzyme. However, there does seem to be quite strong biochemical evidence against this. Collins et al. (316) achieved substantial removal of pyridoxal oxidase activity from aldehyde oxidase by ammonium sulfate fractionation. Furthermore, Dickinson (310)reported that an antibody to aldehyde oxidase failed to cross-react with pyridoxal oxidase. It therefore seems that we are, indeed, dealing with three separate molybdenum hydroxylases. However, further work will be required to establish the extent to which their specificities to reducing substrates overlap, as no doubt they will. Let us now consider the properties of xanthine dehydrogenase, aldehyde oxidase, and pyridoxal oxidase in relation to those of molybdenum hydroxylases from other sources. There is, of course, no a priori reason for expecting the Drosophila enzymes to have properties grossly different from those in other organisms. Indeed, workers in this field clearly ought to be aware of some of the unusual properties of the molybdenum hydroxylases, particularly, perhaps of their wide specificities and of the possibility of their conversion in vitro to inactive desulfo forms. It may be well, also, to remember that there exist naturally occurring nonfunctional demolybdo forms of some of these enzymes (cf. Section 111,B). Hence, occurrence of demolybdo forms of the enzymes in Drosophila would not be in any way unexpected. The.first question we have to consider here is the one of acceptor (oxidizing substrate) specificities and whether it is meaningful to refer to 317. T. T. T.Yen and E. Glassman, BBA 146,35 (1967).

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one of the Drosophila enzymes as a dehydrogenase but to the other two as oxidases. Almost certainly this distinction should not be made. The direct experimental evidence, such as it is, seems to be as follows. There appear to have been few, if any, systematic attempts a t studying acceptor specificities and, certainly, reliable data comparing relative rates for the three enzymes toward different acceptors are not available. Most workers have simply assayed under standard conditions for xanthine dehydrogenase using NAD' or methylene blue as acceptor (e.g., 317) and for aldehyde or pyridoxal oxidase activities with oxygen (e.g., 311). Although artificial acceptors are sometimes used for aldehydes (e.g., 316,516), it has nevertheless been shown (316) that oxygen can function, without any additional acceptor, with them. In the case of xanthine dehydrogenase, on the other hand, there seems to have been surprisingly few attempts to test for oxidase activity, apart from an early report (31.2) that this corresponded to the not negligible value of 5-10% of the dehydrogenase activity. I n the light of information in earlier sections of this chapter, there is clearly much scope for reinvestigating acceptor specificities of the Drosophila molybdenum hydroxylases. Future work ought to take into consideration both the wide ranges of activities toward different oxidizing substrates which are possible (Section 1111C,2) and, even more pertinently, the possibility (Section II,B,3,d) that a given enzyme molecule might be convertible, under appropriate conditions, from a dehydrogenase to an oxidase and perhaps back again (318). One apparently highly anomalous property for a molybdenum hydroxylase has been reported for Drosophila xanthine dehydrogenase by Yen and Glassman (317) and has been quoted in a review (3074. These workers suggested (317) that xanthine dehydrogenase has two active sites, one functioning for purine substrates and the other for pteridines. This was based on inhibition studies, in which pteridines were reported to inhibit the oxidation of pteridine substrates, competitively, and the oxidation of purine substrates, noncompetitively. With purine inhibitors, this situation was reported to be reversed. Examination of the published data (317),however, shows such a conclusion to be quite unjustified. Lineweaver and Burk plots seem to have been drawn in a somewhat arbitrary manner. The inference of two types of active sites seems quite unfounded. Further studies on Drosophila xanthine dehydrogenase have provided considerable information about the enzyme, although this is sometimes of a type not available on enzymes from other sources. We will now at318. There were some possible indications of changes in acceptor specificity during purification of xanthine dehydrogenase in early work on the enzyme (312).

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tempt to summarize some of this work. We first discuss nutritional and maternal effects relating to the enzyme, which, although not properly understood, seem, superficially a t least, analogous to corresponding phenomena observed in the rat. Improving the nutritional status of wild-type Drosophila leads to increased xanthine dehydrogenase activity (319). Comparable effects have long been known in the r a t (276). However, whereas in Drosophila the indications (319) are, apparently, that increased activity may result from activation of a preexisting protein (e.g., by incorporation of a cofactor), this is not the case with the rat, where improved nutrition leads (276) to increased de novo synthesis of enzyme protein. The analogy with the rat enzyme in relation to the maternal effect observed with ma-Z Drosophila mutants (see 320 for summary), is closer. M a 4 mutants, bred from nonmutant mothers, have red eyes and apparently cannot synthesize xanthine dehydrogenase. However, some of the m a 4 gene product is somehow transmitted through the eggs, so that, although xanthine dehydrogenase is scarcely detectable in the egg itself, there are nevertheless substantial amounts of the enzyme (10% of the wild-type level) in newly hatched flies. The comparable finding in rats is that the animals are born without xanthine oxidase (321) and, to attain normal adult levels of the enzyme, require the “xanthine oxidase factor” (322), which is available from milk and which turned out to be nothing more complicated than molybdenum (3f?3,324). Similarly, it seems clear that there must be a close relationship between the m d +gene product and molybdenum, although, as will be discussed further below, the nature of this relationship is quite uncertain. It is also interesting that nutrition somehow comes into the maternal effect (520),as does protein intake into the rat phenomena (321). The finding of naturally occurring electrophoretic variants of Drosophila xanthine dehydrogenase, which map a t the ry locus, was important in identifying this as a structural gene for the enzyme (309).The properties (317) of these variants are also of some interest. It seems that they represent forms of the enzyme with modifications in their amino acid compositions, which do not, however, affect the active site regions. Thus, four variants were found (317)to have differing electrophoretic mobilities but similar K , and Ri values. Electrophoresis has also been used (3.94~) 319. J. F. Collins, E. J. Duke, and E. Glassman, BBA 208, 294 (1970). 320. A. Chovnick and J. H. Sang, Genet. Res. 11, 51 (1968). 321. W. W. Westerfeld and D. A. Richert, JBC 184, 163 (1950). 322. D. A. Richert and W. W. Westerfeld, JBC 192, 49 (1951). 323. D. A. Richert and W. W. Westerfeld, JBC 203,915 (1953). 324. E. C . De Renso, E. Kaleita, P. G. Heytler, J. J. Oleson, B. C . Hutchings, and J. H. Williams, ABB 45, 247 (1953). 324a. T. Shinoda and E. Glamman, BBA 160,178 (1988).

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to distinguish two forms of xanthine dehydrogenase, called I and 11, from wild-type flies. The I form is converted to 11, in what may be a proteolytic reaction. Whether the two forms differ in their oxidase-dehydrogenase properties (cf. Section II1B,3,d) does not seem to have been investigated. Further study (324b) of electrophoretic variants led to the most important conclusion that the ry locus represents a single, unique, and uninterrupted sequence of DNA, having a length, deduced from recombination data, of ca. 3000 nucleotide pairs. This sequence codes for a polypeptide, containing some 10oO amino acid residues, presumed to represent the complete half apoxanthine dehydrogenase molecule (see Section II1B,2,c). In vitro complementation phenomena reminiscent of those related to nitrate reductase in Neurospora have been observed with Drosophila xanthine dehydrogenase. If extracts of ma-1 and ry flies are mixed, a restoration of enzymic activity results. This restoration appears (325) to involve interaction between species from the two extracts, each with molecular weights in the region of 250,000, to give a product, the active xanthine dehydrogenase, also having about this molecular weight. It was suggested that a small cofactor was transferred between the two molecules in this process and that the cofactor might be derived from pyridoxal oxidase of the ry flies. Ma-l flies contain material which cross-reacts with antibody to partially purified xanthine dehydrogenase from wild-type flies (312).Further, the genetic data are consistent (3074 with the ma-1 cistron containing information for a relatively small molecule only. Thus, the ma-l+gene product, although its precise nature, as well as its relationship t o the lxd+ product remains uncertain, seems (307d) closely analogous t o the nit-1 product in Neurospora. It is presumably a small molecule, containing, or otherwise related to, molybdenum. Finally, detailed study of the ma-1 locus has provided information which could have a bearing on the structure of the xanthine dehydrogenase molecule itself. Chovnick and co-workers (326,327) concluded that ma-l is a single cistron exhibiting allele complementation and th a t the biologically active product is presumably a dimer or higher multiple aggregate. The question arises, however (cf. 326) , from the biochemical point of view, of what, precisely, it is that has been shown in this work to be a dimer. We already know that the milk xanthine oxidase molecule, and no doubt other molybdenum hydroxylases also, are dimers (Section 324b. W. M. Gellbart, M. McCarron, J. Pandey, and A. Chovnick, Genetics 78, 869 (1974). 325. E. Glassman, T. Shinoda, H. M. Moon, and J. D. Karam, J M B 20,419 (1966). 326. A. Chovnick, V. Finnerty, A. Schalet, and P. Duck, Genetics 62, 145 (1969). 327. V. Finnerty and A. Chovnick, Genet. Res. 15,351 (1970).

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II1B,2,c). Perhaps the work on ma-1 could simply be a genetic confirmation of this fact. However, there are a number of other possibilities. It seems to the present author that these might range from the rather uninteresting one of some dimeric molecule being involved in a step in the synthesis of the molybdenum site (cofactor) of the enzyme, to there being a twofold symmetry of the groupings bearing the ligand atoms of molybdenum in the enzyme-active center itself.

E. MOLYBDENUM HYDROXYLASES IN Aspergillus nidulans Work on the molybdenum hydroxylases of Aspergillus nidulans is in many respects analogous to that on the enzymes in Drosophila. If in some respects the former is a t a less advanced state than the latter, particularly, e.g., in that the Aspergillus enzymes have not been purified, there are, nevertheless, a number of features of distinct interest. There is firm evidence for two molybdenum hydroxylases only in A s pergillus, given the names “xanthine dehydrogenase I” and “xanthine dehydrogenase 11” by Scazzocchio and co-workers (328). However, the possibility exists that an “alternative pathway” for xanthine oxidation in the organism which has been invoked (9) might in fact involve a third such enzyme. Little biochemical work has been reported to date on either of the xanthine dehydrogenases (307a,328), although molecular weight studies have recently been carried out by gel electrophoresis (329). Apparently, xanthine dehydrogenase I1 has a molecular weight higher, by some 20,000 daltons, than xanthine dehydrogenase I, while the latter is larger than milk xanthine oxidase by a similar amount. Immunological data indicate that xanthine dehydrogenases I and I1 are distinct, although closely related, proteins (328). Substrate specificities have not been systematically investigated. Hypoxanthine rather than xanthine is the substrate routinely used for assaying the enzymes, although apparently either of these substrates will serve (330).Xanthine dehydrogenase I, and apparently also xanthine dehydrogenase 11, has NADH dehydrogenase activity (331). A major specificity difference between the two enzymes is that nicotinic acid is a substrate for xanthine dehydrogenase I1 but not for I. Nicotinic acid does not seem to have been reported as a substrate for molybdenum hydroxylases from other sources. The most nearly analogous substrate for the milk and rabbit enzymes, as reported by Krenitsky 328. C. Scazzocchio, F. B. Hall, and A. I. Foguelman, Eur. J. Biochem. 36, 428 (1973). 329. N. Lewis and C. Scazzocchio, personal communication. 330. A. J. Darlington, C. Scazzocchio, and J. A. Pateman, Nature (London) 206, 599 (1965). 331. C. Scazzocchio, Mol. Gen. Genet. 125, 147 (1973).

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et al. (101, appears to be pyridoxal. A second distinction between the two Aspergillus enzymes is that I1 is less sensitive than I to inhibition by allopurinol (307a,328). The xanthine dehydrogenase of Aspergillus is controlled by a number of different genes (see, e.g., 307a,b, for summaries). Of these, there are five cnx loci controlling the “common cofactor” of the xanthine dehydrogenases and nitrate reductase. Some of these have already been discussed in Section IV,C, in relation to nitrate reductase. I n particular, we note from the cnx E work the direct relationship of the “cofactorll to molybdenum. We also note, from the cnx H work, that the “cofactor” must contain some protein and that it is the precise composition of this protein component of the enzyme which determines the heat stability of nitrate reductase and by implication, also, that of xanthine dehydrogenase, in the cnx H mutants. Thus, more detailed information is available relating to cnx than is available on the analogous ma-1 of Drosophila. Of the other genes affecting the xanthine dehydrogenases, hx B seems . completo be a structural gene for both enzymes ( 3 0 7 ~ )Intracistronic mentation work (332; cf. 328) indicates, at least in the case of xanthine dehydrogenase I, that the gene product is present as a dimer (or polymer) in the enzyme. Genetic data indicated (328) that, in contrast to hx B , the hx A gene codes for a structural component of one of the enzymes only, namely, xanthine dehydrogenase I. It was suggested (328) that this component carries the substrate binding site of this enzyme and further ( 3 0 7 ~ )that ) for xanthine dehydrogenase 11, the hzn C gene may play an analogous role. Two more genes, ua Y (331) and up1 A (328)) have specific roles in the induction of the enzymes. Scazzocchio and co-workers (307a,328),on the basis of the evidence summarized above, have proposed the hypothesis that both xanthine dehydrogenases consist of a protein core, specified by hx B, together with the cnx cofactor (peptide) and another peptide, carrying the substrate binding site. The latter is specified in xanthine dehydrogenase I by hx A , while a different peptide is specified in an analogous manner for xanthine dehydrogenase I1 by hxn C . On the basis of the intracistronic complementation data, together with the information on the dimeric nature of molybdenum hydroxylases generally (Section II1B,2,c), it was suggested (307a) that each xanthine dehydrogenase molecule must consist of six subunits. There would be two hx B cores, in contact with each other, and two each of the c m and hx A units. I n each half of the molecule there would be a cnx unit in close contact with an hx A , but the two cnx units, and similarly the two hx A units would be remote from one another. Interesting though the hypothesis is, it may be that other 332. M. J. Hartley, Genet. Res. 16, 123 (1970).

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explanations of the data are not excluded a t the moment. The biochemical extension of studies on the Aspergillus enzymes is thus awaited with considerable interest.

V. Sulflte Oxidare of liver

A. INTRODUCTION Hepatic sulfite oxidase, which converts sulfite t o sulfate and can use oxygen or other acceptors, has been studied in detail only comparatively recently. The enzyme has been extensively purified from bovine (333), chicken (8),and rat (334) livers. There is also evidence for very closely related enzymes in man (8),as well as in pigs and rabbits (334). The source from which the enzyme has been most fully investigated is bovine liver, and it is this enzyme to which we shall be referring where no other source is specified. The ehzyme contains molybdenum, apparently in an environment quite similar to that of the metal in the molybdenum hydroxylases. It also contains heme, but iron-sulfur and flavin groups are not present. A unique feature relating to sulfite oxidase is that its presence in the livers of various species can readily and directly be detected by observation of highly characteristic Mo(V) EPR signals from the enzyme (334; cf. 334a). The normal biological function of sulfite oxidase is presumed (335) to be in the oxidation of endogenous sulfite, arising from the degradation of sulfur amino acids. However, the enzyme also seems (555) to be instrumental, in the rat, in countering some of the toxic effects of respired sulfur dioxide. Sulfite oxidase is localized in the intermembrane space of mitochondria and its action can be coupled to synthesis of ATP (336).

B. MOLECULAR PROPERTIES Sulfite oxidases from bovine, chicken, and rat livers have all been purified to apparent homogeneity. The bovine enzyme has a molecular weight 333. H. J. Cohen and I. Fridovich, JBC 246,359 (1971). 334. D. L. Kessler, J. L. Johnson, H. J. Cohen, and K. V. Rajagopalan, BBA 334, 86 (1974). 334a. J. Peisach, R. Oltrik, and W. E. Blumberg, BBA 253, 58 (1971). 335. H. J. Cohen, R. T. Drew, J. L. Johnson, and K. V. Rajagopalan, Proc. N u t . 'Acad.Sci. U . S. 70, 3655 (1973). 336. H. J. Cohen, S. Betcher-Lange, D. L. Kessler, and K. V. Rajagopalan, JBC 247, 7759 (1972).

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415

of 115,000 (33613).The subunit molecular weight for this enzyme is 55,000 (SSSu),no doubt indicating a dimeric structure for the enzyme. For the chicken enzyme, the subunit molecular weight is 55,000 and for that from the rat, 58,000 (334).The purified chicken enzyme seems, however, not to be entirely in the dimeric form (8). Sulfite oxidase contains molybdenum and heme in the ratio 1.05 to 1 for the bovine enzyme (337) and 0.93 to 1 for that from chicken (8) and about 1 to 1 for the rat enzyme (334).The heme is present in the form of a b,-like cytochrome, and there are two hemes per enzyme molecule, i.e., one per subunit (33613).Although the above analytical data give no evidence for a demolybdo form of sulfite oxidase under normal nutritional conditions, one is formed, however, during administration of tungsten to rats (58,5513,33713). Sulfite oxidase, in contrast to the molybdenum hydroxylases, contains only two redox-active groups, namely, the molybdenum and the heme. The latter can of course exist in ferri- and ferro-states, these having very In the oxidized state, a low-spin characteristic absorption spectra (35613). heme EPR spectrum with g1 2.93, g2 2.25, and g, 1.53 has also been observed (8) in the case of chicken enzyme. The redox state of molybdenum, on the other hand, which apparently makes little contribution to the visible absorption spectrum, can be monitored only by observation of the EPR signals from the metal when it is in the five-valent state. By analogy with the molybdenum hydroxylases, this metal in the sulfite oxidase might be expected to be six-valent in the oxidized enzyme and to be capable of reduction to the five- and four-valent states, with equilibria among the various valencies when the enzyme is partially reduced. This indeed seems to be the case, although redox titrations of the enzyme have not been reported. Purified, resting bovine sulfite oxidase gives no molybdenum signal, but on suitable reduction with sulfite 50-60% of the metal is converted to the signal-giving form, while reduction with excess dithionite causes the signal to disappear again (337). In the purified enzymes, Mo (V) signals have been elicited by reduction with sulfite and apparently not by any other means. I n mitochondria, as normally prepared, the signals are present, indicating the enzyme to be partially reduced. Dialysis of mitochondria a t low salt concentrations liberates sulfite oxidase into the supernatant fraction and allows it to become oxidized with disappearance of the signals (336). Enzyme in 336a. H. J. Cohen and I. Fridovich, JBC 246, 367 (1971). 337. H. J. Cohen, I. Fridovich, and K. V. Rajagopalan, JBC 246, 374 (1971). 337a. There is evidence (66~) that tungsten administration to rats leads to production both of a tungsten-containing analog of sulfite oxidase and of the W-free demolybdo form.

416

R. C. BRAY

TABLE XI1 E P R PARAMETERS OF Mo(V) SIQNALS FROM BOVINE SULFITE OXIDAGE'

('HI (GI

IAavl

Signal High p H Low p H

Qav

1.965 1.977b

91

1.984 2.000

None lob

Data from Cohen et al. (337) for solutions in 0.1 M Tris-HC1. Accuracy of the measurements was not stated. * Parameters remeasured from the published spectrum, taking account of the apparent, slight deviation from axial symmetry.

dialyzed but uncentrifuged mitochondria1 preparations gave signals on treatment either with sulfite or with NADH. No doubt in the latter case, NADH was acting in an indirect manner to produce some other reducing species which interacted with the enzyme, but this observation does indicate that sulfite plays no specific role in production of Mo(V) signals from the enzyme. Let us now consider the Mo(V) EPR signals themselves. Two signals have been observed (337) and their parameters are given in Table XI1 (338). Inspection of the published spectra indicates that each signal may well correspond to single chemical species, although apparently no 35 GHz spectra have been recorded or computer simulations attempted in order to confirm this. The high pH spectrum (Table XII) has a rhombic form without indications of proton or nitrogen hyperfine structure. The low pH spectrum is nearly axial in form and shows splitting by a single proton, which is exchangeable with protons from solvent water molecules. The high pH and low pH signal-giving species are convertible into one another, simply by changing the p H of the medium, the pK value for their interconversion being 8.2 (337). Presumably the proton taken up during conversion of the high p H species is the one "seen" in the spectrum of the low pH form. Although there is clearly a parallel, both in the signals themselves and in this p H equilibrium, with the corresponding situation relating to Rapid and Very Rapid signals from milk xanthine oxidase, there are nevertheless important differences. First, whereas for the milk enzyme, the high pH species (Very Rapid) is a tran338. "Mo hyperfine structure has been observed (337) for one signal, unambiguously identifying molybdenum as the signal-giving species. Whether MOW) or Mo(II1) is involved has not been proved rigorously but by analogy with the milk enzyme, MOW), is assumed. Disappearance of the signals with dithionite argues strongly that this is correct since reduction to Mo(I1) is very unlikely (cf. discussion of the Slow signal in Section II,B,4,d),

6.

MOLYBDENUM HYDROXYLASES

417

sient detectable only in the time range of about 5-50 msec, in contrast, for sulfite oxidase, both high and low p H signals are stable. With regard to the signals, the low p H sulfite oxidase spectrum is somewhat reminiscent of the Rapid xanthine oxidase signal but for the former, g values are slightly lower and the proton splittings slightly smaller (compare Tables I11 and X I ) . The analogy between the high pH sulfite oxidase and Very Rapid signals is less strong, particularly in that the very high g3 value of Very Rapid is not reproduced. Furthermore, whereas the Very Rapid to Rapid conversion is accomplished with little change in gBY,this is not so for the corresponding conversion of sulfite oxidase. Presumably these differences indicate that, despite similarities, there are some definite differences in the nature or positioning of some of the ligand atoms of molybdenum in the two types of enzyme. An interesting feature of sulfite oxidase E P R spectra is that the precise parameters are relatively sensitive to the presence of anions such as phosphate or chloride (8,334).Although the anions apparently affect the value of the pK controlling interconversion of the two signal-giving species, there seem to be other effects on the spectra also. Presumably the phenomenon is related to inhibition of sulfite oxidase by anions (333). Differences among animal species, in the EPR parameters of M O W ) in the sulfite oxidases, seem small. Most of the points discussed above for the bovine enzyme apply also to the chicken and rat enzymes (8,334). Precise parameters have not always been recorded but we do find some very small but apparently significant differences among them, e.g., the value of gC for the high pH species is 1.950 for the bovine (337) and 1.954 for the rat (334) enzyme. However, no doubt the environment of Mo in all sulfite oxidases is very similar. C. CATALYTIC PROPERTIES Although sulfite oxidase appears to have a virtually absolute specificity for sulfite as reducing substrate, it may well be that a reducing agent such as dithionite ought also to be thought of as a substrate for the enzyme. The specificity with regard to the oxidizing substrate is lower (333) and ferricyanide or cytochrome c, as well as the dyes, indophenol and methylene blue, can serve as acceptor, in addition to oxygen. It should be noted, however, that all activities, including oxidase activity, are relatively low in comparison with that toward ferricyanide. Oxygen is reduced to H,O,, with no indication of formation of 0,- (333). Analysis of the steady-state kinetics indicated a ping-pong mechanism, both in the case of the bovine (333) and of the chicken enzyme (8). K , values for the former were 1.4 X M for sulfite and 5.8 X

418

R.

C. BRAY

M for oxygen; for the latter enzyme they were 2.4 x M for sulfite and 2.2 )( M for cytochrome c. The pH optimum of the bovine enzyme, with oxygen as acceptor, is 8.6 (333). Turnover numbers do not seem to have been calculated. There have been no pre-steady-state kinetic studies reported on the enzyme, but in the steady-state, with ferricyanide as acceptor, it seems that heme is fully oxidized (336%) while molybdenum is substantially reduced (337').On the other hand, with oxygen, heme is fully reduced in the steady state ( 3 3 6 ' ~ ) . Inhibition studies on sulfite oxidase have revealed a number of interesting points. Arsenite inhibits after incubation with the enzyme and so does cyanide (337). Furthermore, treatment with either reagent prevents appearance of Mo(V) EPR signals on addition of sulfite, while cyanide does not affect the visible spectrum of the enzyme, indicating that it is not interacting with the heme. These results raise the question of whether the enzyme ,contains the essential persulfide group, which is present in the molybdenum hydroxylases. In view of the role which has been postulated for this group in xanthine oxidase catalysis (Section II,C,2,f), it would clearly be highly desirable to establish whether or not it is present in sulfite oxidase, also. Information on this point ought to be obtainable either from detection of thiocyanate, which might be liberated in the cyanide treatment, or from appearance of an analog of the Slow EPR signal. If such a signal did exist it would probably be detectable only on partial reduction of cyanide-treated enzyme with dithionite. The finding (337) that methanol does not inhibit the enzyme is probably not evidence against an essential persulfide group in sulfite oxidase, since, although the group is required in xanthine oxidase for reaction of this reagent with the enzyme, nevertheless, it seems probable that persulfide is not the actual site at which a formyl group becomes bound to the milk enzyme in this process (Section IX,C,2,f). Anions inhibit sulfite oxidase only with ferricyanide or cytochrome c as oxidizing substrate and not with oxygen (333). As noted above, they also alter the Mo(V) EPR spectrum, but they do not seem to have been reported to affect the visible spectrum from the heme group. With regard to the mechanism of action of the enzyme, it is assumed (337) that sulfite interacts a t the molybdenum site. Further, the mechanism of this process has been assumed by Stiefel (666) to be quite similar to that of the interaction of reducing substrates with molybdenum in molybdenum hydroxylases and to imolve coupled electron and proton transfers. From the anion inhibition data and from the information on the redox states of molybdenum and heme under steady-state conditions, it has been assumed (337) that oxygen interacts a t the heme site and the one-electron acceptors a t the molybdenum. It is clearly desirable that

6.

MOLYBDENUM HYDROXYLASES

419

this should be confirmed by pre-steady-state kinetic studies. Finally, it would be desirable for work t o be carried out on the stoichiometry of the reduction of the enzyme. This ought to help in elucidating the roles in the catalytic reactions of the enzyme of active center systems, which has accepted, respectively, one, two, or three reducing equivalents. Presumably, only the two-electron reduced form would be involved when oxygen is the acceptor, since 0,- is not formed and sulfite transfers two electrons. Some analogy of the kinetics of sulfite oxidase with those of flavocytochrome b, (272) in this regard might be anticipated. It might also be useful, in the context of the stoichiometry of reduction of the enzyme, to reinvestigate the reported (336a) kinetic differences between behavior of the two heme groups of the sulfite oxidase molecule. The enzyme was reported (336%) to lose some of its activity, reversibly, during turnover with oxygen, with a half-time of the order of 1 min, this process correlating with reduction of half of the heme, the other half having been much more rapidly reduced. Although this explanation fits well with the stoichometry of just half of the heme changing in each phase, an alternative would be that the enzyme initially cycles between the SOo and SO,, forms, but gradually changes over to a less efficient cycling between SO,, and SO,, (where SOo . . . SO,, represent active center systems reduced by 0 to 3 electrons, respectively). Clearly, stopped-flow and rapid freezing EPR studies on the enzyme would be interesting. ACKNOWLEDGMENTS Work by the author was supported by a Programme ,Grant from the Medical Research Council. Comments on the draft manuscript, or parts of it, by the following were of great assistance: M. P. Coughlan, D. J. Lowe, C. Scazzocchio, H. A. Simmonds. Preprints of variou papers received from the above, and also from the following, are acknowledged with thanks: R. Andres, D. J. Cove, R. Eisenthal, V. Finnerty, V. Massey, A. Nason, T. Nishino, G. Palmer, K. V. Rajagopalan, and P. M. Wood. Help from M. J. Barber and D. J. Lowe in the preparation of Fig. 2 is also acknowledged.

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Flavofirotein Oxidases HAROLD J . BRIGHT

DAVID J . T. PORTER

.

I Introduction . . . . . . . . . . . . . . . I1. The Flavin Coenzyme . . . . . . . . . . . . I11. Kinetic Methods Applied to Flavoprotein Oxidases . . . A . Kinetic Strategy . . . . . . . . . . . . B . Steady-State Kinetics . . . . . . . . . . . C . Transient-State Kinetics: The Half-Reactions . . . . D . Kinetic Mechanism . . . . . . . . . . . . E . Computer Simulation Studies . . . . . . . . F. Summary of Major Kinetic Results . . . . . . . IV . The FIavoprotein Oxidases: MoIecular Properties and Kinetic Mechanism . . . . . . . . . . . . . . A . &Amino Acid Oxidase . . . . . . . . . . . B . IrAmino Acid Oxidase . . . . . . . . . . . C . Glucose Oxidase . . . . . . . . . . . . D . Monoamine Oxidase . . . . . . . . . . . E . Old Yellow Enzyme . . . . . . . . . . . V. The Chemical Mechanism of Flavoprotein Oxidases . . . A . The Chemical Mechanism of Flavin Reduction . . . B . The Mechanism of Oxidation of Reduced Flavin by 02 .

. . .

421

. 423

. . . . . .

. . .

. . . . . . .

. .

425 425 . 426 . 432 . 435 . 442 . 443

. . .

445 445 456 . 461 . 466 . 471 . 474 . 474 . 503

.

1 Introduction

The concept of a flavoprotein oxidase is easier to understand than to define by international rules ( I ) . A flavoprotein. or flavoenzyme. is commonly understood to mean a n apoenzyme which. together with its more or less tightly attached flavin coenzyme (either FAD or FMN or deriva1 . International Union of Biochemistry Commission. “Enzyme Nomenclature. Recommendations of the International Union of Pure and Applied Chemistry and the International Union of Biochemistry.” Elsevier. Amsterdam. 1973. 421

422

HAROLD J. BRIGHT AND DAVID J. T. PORTER

tives thereof), catalyzes a redox reaction during which either one or two electrons from the electron donor are transferred transiently to the isoalloxazine nucleus of the flavin coenzyme and then to the electron acceptor. Clearly, then, all flavoproteins (except for those such as E C 2.1.1.21, which do not catalyze a net oxidation of the donor substrate) belong to the class oxidoreductases (1). The term “oxidase” is a recommended name for an oxidoreductase which utilizes O2as the electron acceptor (1). More accurately, perhaps, an oxidase should be defined as an oxidoreductase which catalyzes a reaction in which all of the electrons taken from the donor are transferred to O2 to form any of the reduction products of O2 known in biological systems ( O Z ~HzOZ, , or HzO). The Recommendations for Enzyme Nomenclature ( l ) ,based as they are on the functional groups of the electron donor and acceptor in the case of oxidoreductases, do not allow for flavoprotein oxidases except in a meaningless sense. For the purpose of this chapter and in accord with the understanding of most biochemists, a flavoprotein oxidase is defined as a flavoprotein which catalyzes a reaction having the stoichiometry of Eq. (1) [where -XH = -OH, -NH,, -NHR, or, uncommonly, -CHR, or -C (R) = R’) I

H-C--XH

+

O,->C=X

+

H202

I Metalloflavoprotein oxidases, such as xanthine oxidase, as well as dehydrogenases having weak oxidase activity, form superoxide anion rather than HzO, and are therefore not included in this definition ( 2 ) .Two other characteristics, namely, the ability to form a red (anionic) flavin semiquinone and a flavin-sulfite adduct, would be typical of, but not strictly confined to, flavoprotein oxidases so defined ( 5 ) . At the present state of knowledge, therefore, our definition singles out the simple flavoprotein oxidases in which flavin is the only recognizable prosthetic group that, transiently, accepts electrons originating in the donor substrate ( 4 ) . Of 2. V. Massey, G. Palmer, and D. Ballou, in “Flavins and Flavoproteins”

(H. Kamin, ed.), p. 349. Univ. Park Press, Baltimore, Maryland, 1971. 3. V. Masaey, F. Muller, R. Feldberg, M. Schuman, P. A. Sullivan, L. G. Howell, 5. G. Mayhew, R. G. Matthews, and G. P. Foust, JBC 244, 3999 (1989). R. J. DeSa [JBC 247, 5527 (1972)l has reported that putrescine oxidase is unreactive with sulfite. 4. This definition includes three enzymes which appear to oxidize carbonyl groups, namely, pyruvate oxidase (EC 13.3.3), oxalate oxidase (EC 1.2.3.4),and glyoxylate oxidase (EC 1.2.3.5).The definition of Eq. (1) will hold if the actual (enzyme-bound) substrate to be oxidized by flavin in the first two caws is a decarboxylated substratethiamine pyrophosphate adduct (XH = -C(R)=R’) and, in the third case, the hydrate of glyoxylate(-xH = -OH). Pyruvate oxidase contains thiamine pyrophosphate (I), and it is reasonable to suppose that oxalate oxidase also utilizes this coenzyme.

7.

FLAVOPROTEIN OXIDASES

423

the 80 or so oxidoreductases recognized to be flavoenzymes in 1972, approximately 20 of these (namely, EC 1.1.3.1,4,5,12,13, and 15; E C 1.2.3.3,4, and 5 ; EC 1.3.3.1; EC 1.4.3.1,2,3,4,5,and 9 ; EC 1.5.3.2,5, and 6; E C 1.6.99.1; and Ec 1.7.3.1) would, by our definition, be flavopretein oxidases (1) . The emphasis of this chapter concerns the kinetic and chemical mechanism of flavoprotein oxidase catalysis. There are several cogent reasons for such relatively restricted coverage. First, with the notable exception of monoamine oxidase (EC 1.4.3.4),the precise biological function of the flavoprotein oxidases is, for the most part, rather obscure or, a t best, probably of minor quantitative significance. Although highly exergonic thermodynamically, none (with the exception of pyruvate oxidase, E C 1.2.3.3) is directly coupled to ATP synthesis. Consequently, questions concerning regulation of their synthesis or in vivo activity have little biological significance. Second, the fact that no X-ray crystallographic studies of a flavoprotein oxidase have been published has tended to discourage systematic studies of their protein chemistry, such as sequencing and modification. Third, as with other coenzyme-requiring enzyme systems, model studies directed a t the question of kinetic and chemical mechanism are highly feasible. In the case of flavin, recent advances in our understanding of the mechanism of both the enzymic and nonenzymic reactions have resulted in a most productive liaison between the two approaches. I n addition to its restriction in topic, this chapter deals for the most part with enzymological and model studies related to the mechanism of action of only three flavoprotein oxidases, namely, glucose oxidase (EC 1.1.3.4), L-amino acid oxidase (EC 1.4.3.2), and D-amino acid oxidase (EC 1.4.3.3). This is a consequence, simply, of the fact that, through a disproportionate amount of research effort, we understand these three enzymes far better than any other flavoprotein oxidases. It is intended to demonstrate that the basic molecular mechanisms of these enzymes must be very similar and are probably applicable, in turn, to the other less well-known flavoprotein oxidases. It is our belief that a firm understanding of this class of flavoenzymes, which is probably the simplest mechanistically, will be invaluable for the unraveling of the molecular events which underly the ubiquitous processes of flavoenzyme-catalyzed dehydrogenation and hydroxylation. 11. The Flavin Coenzyme

The flavin coenzyme in its fully oxidized state has the following structure (I) where the R group a t N-10 of the flavin (isoalloxazine) nucleus is either adenosyldiphosphoribityl (FAD) or phosphoribityl (FMN) . Lumiflavin (R = CH,) and its derivatives are frequently used as model

424

HAROLD J . BRIGHT A N D DAVID J. T. PORTER

compounds. Excellent reviews of the molecular physics and chemistry of flavin are available which emphasize structure and reactivity (5-15) , intra- and intermolecular complexation (14,15), flavin free radicals (1618), optical and fluorescence properties (19,20), X-ray crystallographic structure ( 2 1 ), and molecular orbital calculations (22). R

0 (1)

The flavin nucleus exists in three redox states, each of which can adopt three states of ionization (10J2). Although the apoenzymes preferentially bind certain redox and ionization states through a variety of mechanisms 5. G. R. Penzer and G. K. Radda, Quart., Rev., Chem. Soc. 21, 43 (1967). 6. P. Hemmerich and M. Schuman Jorns, FEBS Symp. 29,95 (1972). 7. P. Hemmerich, S. Ghisla, U. Hartmann, and F. Muller, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 83. Univ. Park Press, Baltimore, Maryland, 1971. 8. P. Hemmerich, A. P. Bhaduri, G. Blankenhorn, M. Brustlein, W. Haas, and

W.-R. Knappe, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 3. Univ. Park Press, Baltimore, Maryland, 1973. 9. P. Hemmerich, G. Nagelschneider, and C. Veeger, FEBS (Fed. Eur. Biochem. Symp.) Lett. 8, 69 (1970). 10. P. Hemmerich, C. Veeger, and H. C. S. Wood, Angew. Chem., Znt. Ed. Engl. 4, 671 (1965). 11. P. Hemmerich and F. Muller, Ann. N . Y . Acad. Sci. 212,13 (1973). 12. H. Beinert, “The Enzymes,” 2nd ed., Vol. 2, Part A, p. 339, 1960. 13. G. Palmer and V. Massey, in “Biological Oxidations” (T. P. Singer, ed.), p. 263. Wiley, New York, 1968. 14. T. M. Kosower, in “Flavins and Flavoproteins” (E. C. Slater, ed., BBA Libr., Vol. 8, p, I. Elsevier, Amsterdam, 1966. 15. G. Weber, in “Flavins and Flavoproteins” (E. C. Slater, ed.), BBA Libr., Vol. 8, p. 15. Elsevier, Amsterdam, 1966. 16. A. Ehrenberg, L. E. G. Eriksson, and F. Muller, in “Flavins and Flavoproteins” (E. C. Slater, ed.), BBA Libr., vol. 8, p. 37. Elsevier, Amsterdam, 1966. 17. F. Miiller, P. Hemmerich, and A. Ehrenberg, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 107. Univ. Park Prem, Baltimore, Maryland, 1971. 18. G. Palmer, F. Miiller, and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 123. Univ. Park Press, Baltimore, Maryland, 1971. 19. V. Massey and H. Ganther, Biochemktry 4, 1161 (1965). 20. S. Ghisla, V. Massey, J.-M. Lhoste, and S. G. Mayhew, Biochemistry 13, 589 (1974). 21. P. Kierkegaard, R. Norrestam, P.-E. Werner, I. Csoregh, M. von Glehn,

R. Karlsson, M. Leijonmarck, 0. Rijnnquist, B. Stensland, 0. Tillberg, and L. Torbjornmn, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 1. Univ. Park Press, Baltimore, Maryland, 1971. 22. M. Sun and P.8.Song, Biochemistry 12,4663 (1973).

7.

425

FLAVOPROTEIN OXIDASES

(10,13,19), the counterparts of the following equilibria for free flavins will be predominant in flavoprotein oxidases in the pH range where these retain their native structure (10). The ionizations involve the proton on N-3 for HF, and the proton on N-1 for both H2Fand H,Fr.

Fully oxidized

HFO yellow

-

pK,

-

F,

+

H+

HF-

+

H+

It PKa

Semiquinone (radical)

= 10

6.5

. $ H

(2)

H++eII

pKa Fully reduced (1,5-dihydroflavin)

= 6.2

HsFr "colorless"

-

-H,F; + H+ "colorless"

-

The fully oxidized (E,, E, * * S, and E, * * PI) and fully reduced (Er and E, * S) redox states of flavoprotein oxidases are clearly important, and often obligatory, catalytic intermediates by spectrophotometric criteria. Equally clearly, the free flavoprotein oxidase semiquinones corresponding to H2F and HF', which are produced by partial dithionite titration or photochemically in the presence of EDTA (18), are not catalytic intermediates in the oxidation of physiological substrates, although tight biradical complexes of flavin and substrate have not been eliminated as short-lived intermediates (see Section V ) . Full reduction of the flavin nucleus is accompanied by an abrupt change from a planar t o nonplanar (butterfly wing) configuration (21) corresponding to A< lo4 M-l cm-I in the 450-460-nm region. Thus, static or transient spectrophotometric measurements are readily carried out in this wavelength range. The intermediate E,. * P, (see Section V) in the amino acid oxidase reactions gives A€ x 2-3 X lo3 M-I cm-1 (with respect to E,) a t 550 nm and is conveniently monitored a t this wavelength. Selected aspects of model 'flavin chemistry are discussed in Section V as they relate to the chemical mechanism of flavoprotein oxidase catalysis.

- -

-

-

-

111. Kinetic Methods Applied to Flavoprotein Oxidases

A. KINETICSTRATEGY The flavoprotein oxidases are uniquely amenable to kinetic analysis for three reasons. First, the catalytic cycle can be broken into two

426

HAROLD J. BRIGHT A N D DAVID J . T. PORTER

half-reactions which can be studied separately. These half-reactions are referred to as the reductive (E, S +) and oxidative (E, 0, +) halfreactions. Second, the flavin chromophore is an intrinsic spectrophotometric probe through which the transient oxidized and reduced states of the enzyme can be monitored with high sensitivity. Third, the 0, electrode technique can rapidly generate steady-state kinetic data of high precision. Taken together, these circumstances allow for the correlation of transient and steady-state kinetic behavior over wide ranges of experimental conditions and hence permit the deduction of kinetic mechanism to be carried out in an orderly and logical fashion. For these reasons, the application and interpretation of kinetic measurements will be described in the context of a model investigation, using where appropriate specific examples from the three best understood reactions, namely, glucose oxidase and the D- and L-amino acid oxidases. The investigation takes place in four major stages. First, the steady-state parameters are measured. Second, stopped-flow spectrophotometric measurements of the separate half-reactions are carried out. Third, comparison of the results from the first two stages leads to enzyme-monitored flow studies of the complete turnover process (and perhaps to double mixing experiments) , and then to a working hypothesis for the kinetic mechanism. Lastly, checks and refinements of the kinetic model are conducted through computer simulation studies. This section covers only the major kinetic techniques and findings. Specific aspects are described in Section IV for each enzyme in turn.

+

+

B. STEADY-STATE KINETICS 1. Steady-State Velocity Measurements [SSK ( S / P )]

The flavoprotein oxidases catalyze a two-electron oxidation of substrate which is linked to the two-electron reduction of 0, [Eq. (3) ] :

The initial product P, usually undergoes a further (nonenzymic) solvolysis reaction to form P, (see Section IV). I n principle, therefore, the steady-state initial velocity can be monitored through S, O,, H,02, PI, or P, (or derivatives of P,). In practice, initial velocities are now usually measured through 0, consumption. The 0, electrode technique has superseded manometric methods because of specificity, rapidity of operation, time resolution (response times being 10 sec or less for most Clark membrane electrodes and con-

7. FLAVOPROTEIN

OXIDASES

427

siderably less than this for vibrating platinum electrodes), accuracy, sensitivity (1-10 pLM 0, can routinely be measured with good precision by the membrane electrode) , and the advantage of continuous recording. The major sources of error concern electrode poisons, contamination of the enzyme preparation by catalase, and electrode calibration. I n the presence of excess (nonrate-limiting) catalase the stoichiometry of Eq. (3) becomes that of Eq. (4).

Hence, if the enzyme preparation contains catalase, Eq. (3) can only be achieved in the presence of an effective catalase inhibitor such as CN( 2 3 ) . Alternatively, addition of excess catalase will assure the stoichiometry of Eq. (4) but will halve the sensitivity of the 0, measurements. The solubility of 0, in pure H,O at 25O amounts to approximately respec0.24 and 1.2 mM after equilibration with air and with pure 02, tively, a t 1 atm. Appreciable corrections may have to be applied in the presence of certain solutes (24) including substrate sugars (25). Fortunately, the easily attainable range of 0, concentration encompasses most of the observed K,,, values. Furthermore, the apparent K , values for 0, are often an order of magnitude or so less than those for substrates with the consequence that 0, is almost always the limiting substrate in kinetic experiments. I n some cases (e.g., glucose oxidase) neither PI nor H,O, has significant affinity for any enzyme species ( 2 5 ) . Consequently, the 0, electrode trace, a t a given concentration of S (which remains effectively constant during the kinetic experiment) provides a n infinite set of steady-state velocities as a function of 0, concentration. Such data may be evaluated by the method of tangents or they may be analyzed through the integrated rate equation (see Section III,B,2,e) by computer (26,27) or by manual methods. When these methods are valid, two complete sets of steady-state graphs of high quality can be obtained from as few as four or five kinetic experiments. In addition to rapid evaluation of the steady-state coefficients such experiments have the advantage of providing, through rapid inspection of the 0, trace, useful qualitative [ S ] and +d[O,] information concerning the relative magnitudes of (see Section 111,B12) in the steady-state equation. If +l/[S] << +2/[02] (and if both are larger than + o ) the 0, electrode trace is highly curved. 23. 24. 25. 26. 27.

B. Chance, JBC 194, 483 (1952). J. Robinson and J. M. Cooper, Anal. Biochem. 33, 390 (1970). Q . H. Gibson, B. E. P. Swoboda, and V. Massey, JBC 239, 3927 (1964). H. J. Bright and M. Appleby, JBC 244,3625 (1969). M. K. Weibel and H. J. Bright, JBC 246,2734 (1971).

428

HAROLD J. BRIGHT AND DAVID J. T. PORTER

This shows immediately that the interaction of 0, with ii reduced species of enzyme is a principal rate-determining process in turnover. If the opposite is true, the steady-state velocity remains independent of 0, and the 0, electrode trace has constant slope. Other methods of monitoring steady-state velocities compare so unfavorably with the 0, electrode technique that they are now used only under special circumstances. The reducing substrate, S, is rarely monitored became it is usually far in excess of 0, and its fractional conversion is therefore usually small. An exception is the case where S itself is highly absorbing or fluorescent (e.g., S = NADH in the case of a diaphorase). H,O, formation can be followed spectrophotometrically through its weak tail absorption at 235 nm (t235= 0.058 mM-l cm-l) ( 2 6 ) . This method requires 0-0.1 absorbance sensitivity and should be used with caution since enzyme-bound FAD and FADH, absorb differently a t this wavelength and sny time-dependent change in these species during turnover will be superimposed on the H20, trace. The method is therefore limited to experiments in which E-FAD is approximately 0.5 pM or less. Alternatively, H,O, can be visualized through coupling by horseradish peroxidase to suitable redox dyes (28). This is the basis for the semiquantitative dipstick test for blood and urinary glucose. The product(s) P, can also be monitored. There are numerous methods for monitoring, for example, the a-keto acid product of the amino acid oxidase reactions. a-Keto acids may be followed continuously by spectrophotometry a t 235 nm (69)or, alternatively, through their weak n + T” transition in the 300-340-nm range in which, fortuitously, an approximate isosbestic point for the enzyme can sometimes be located (SO). This method was particularly useful in establishing the kinetics of the transient accumulation of a-imino acid (the protonated form of which has no R. 3 T* transition) in stopped-flow monitored turnover of D-amino acid oxidase (SO), Alternatively, the a-keto acid can be monitored discontinuously through its characteristic 2,4-dinitrophenylhydrazone particularly when, as with the a-chloro-substituted amino acid substrates, there is a need to differentiate between two types of a-keto acid (31). Finally, the strongly absorbing enol tautomer of P, can be determined when it is stabilized by borate (36) or by hydroxide (33). The latter method, though discontinuous, is extremely 28. H. U. Bergmeyer, ed., “Methods of Enzymatic Analysis,” p. 123. Academic Press, New York, 1965. 29. M. L. Fonda and B. M. Anderson, JBC 242,3957 (1967). 30. D. J. T. Porter and H. J. Bright, BBRC 46,571 (1972). 31. C. T. Walsh, A. Schonbrunn, and R. H. Abeles, JBC 246, 6855 (1971). 32. W. E. Knox and B. M. Pitt, JBC 225, 675 (1957). 33. D. J. T. Porter, Ph.D. Dissertation, University of Pennsylvania, Philadelphia, 1972.

7. FLAVOPROTEIN

OXIDASES

429

sensitive and also fairly rapid because the conditions for enolate formation serve also to quench the enzymic reaction. Enolate measurements were used a t high 0, pressures to evaluate the steady-state kinetic parameters of loop B of Eq. (20) for L-amino acid oxidase (33). 2. The Steady-State Rate Equation

The steady-state turnover data, obtained either as initial velocities or when conditions allow (e.g., glucose oxidase as noted previously) as tangents t o the entire 0, electrode trace, are usually plotted in double reciprocal fashion as [ET]/v versus the reciprocal of the variable substrate (0, or S) a t different levels of the fixed substrate (S or 02). The pair of primary plots usually consists of sets of parallel lines and the lines are commonly straight. However, curved parallel lines, as well as intersecting patterns, are also observed on occasion. Although computer fitting of the graphical data will enhance the precision of the derived parameters, i t has been our experience that 0,-electrode velocity data taken in duplicate or triplicate, when fitted by inspection, lead to numerical values for steady-state coefficients which are in excellent agreement with rate constants associated with transients in stopped-flow experiments. This, in our view, is the most stringent criterion for the reliability of steadystate coefficients. a. Parallel (Straight) Line Patterns. The steady-state rate equation corresponding to this case is the following [ Eq. ( 5 )] :

where +,,-I is the maximum turnover number (in sec-l) and and +,-' are apparent bimolecular constants with units of M-l sec-'. We shall routinely use throughout this chapter the 4 notation and method of Dalziel (S4) for the expression of steady-state rate equations because of the close coefficients to rate constants obtained from correspondence of the stopped-flow experiments. The method of Cleland ( S 5 ) does not offer this advantage and, moreover, product inhibition patterns and Haldane relationships cannot easily be determined for flavoprotein oxidase reactions. Any bisubstrate scheme will give Eq. ( 5 ) if it contains, minimally, one first-order (corresponding to &-l) and two second-order (corresponding to +'-' and +*-l) kinetic processes and if, in addition, along the pathway linking the two enzyme species which bind S and 0,, there is a t least one process which is irreversible (or practically so) under the conditions of the experiment (see Section III,B,l). The simplest possible 34. K. Dalziel, Acta Chem. Scund. 11, 1706 (1957). 35. W. W. Cleland, BBA 67, 104, 173, and 188 (1963).

+

430

HAROLD J. BRIGHT AND DAVID J. T. PORTER

scheme consistent with Eq. ( 5 ) is therefore that of Eq. ( 6 ) , where the first-order process may terminate either of the half-reactions. ki = $0-1

E,+S-----,W W or X

- 1-

+ 02-ks

followed by W

= 6s-1

(6)

Y or E.

kr = 40-1

X or by Y

ka- do-'

E,

If n rate-limiting first-order processes are located in either or both of the half-reactions, then the maximum turnover number becomes n

It is quite evident that no decisions concerning mechanism, beyond the generalities just stated, can be made at this point. However, the steadystate rate equation, together with the evaluated coefficients, are indispensable for the correct interpretation of the rapid kinetic measurements discussed in Section II1,C. This is because the kinetics of each transient in the rapid kinetic measurement must correspond to one or more of the steady-state coefficients if the transient enzyme species is an obligatory intermediate in turnover. Glucose oxidase turnover (25) always conforms to Eq. ( 5 ) , or to special cases of i t (depending on the substrate and pH value under consideration). Similarly, the published data for D-amino acid oxidase (3S,37) fit Eq. ( 5 ) , (although there are reasons to believe that nonlinear double reciprocal patterns will be found for this enzyme). On the other hand, the steady-state behavior of L-amino acid oxidase fits Eq. ( 5 ) only under restricted experimental conditions (33). b. Parallel (Curved) Line Patterns. When initial velocities of phenylalanine oxidation by L-amino acid oxidase are measured over wide ranges of 0, concentration at 2 5 O , the plot of [ETJ/vvs. 1/0, consists of families of curved parallel lines, suggesting activation by 0, (33). Each line consists of two linear regions (higher slope a t high 0,) joined by a curved portion. Over the range of 0, conventionally used M to M) the curvature might easily be missed, but the use of 0, a t high pressures, together with the sensitive enolate assay ( 3 3 ) , clearly shows that the slopes of the linear regions differ by as much as 20-fold, depending on the pH. The plots of ET/v vs. l/S on the other hand give parallel straight lines. This behavior cannot be generally expressed in coefficient form,

+

36. V. Massey and Q. H. Gibson, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 23, 18 (1964). 37. V. Massey, B. Curti, and H. Ganther, JBC 241,2347 (1966).

7.

FLAVOPROTEIN OXIDASES

431

although the limiting linear regions of the 0, plot are, of course, described by Eq. ( 5 ) if +o and +, at low 0, are replaced by +o’ and +*’ a t high 0, (with +o > and +2 < 4,’). Rapid kinetic experiments showed that this behavior results from the occurrence of two oxidative pathways rather than the presence of isozymes differing in their intrinsic +o and +, values (33). c. Intersecting Line Patterns. Under rather special conditions such as low pH (SS), or low temperature (38),L-amino acid oxidase generates intersecting patterns conforming to Eq. ( 8 ) :

The eresence of the cross-term coefficient +,*, albeit under limited conditions, indicates that the parallel line patterns which are found most frequently in studies of the amino acid oxidase reactions are probably not resulting from the obligatory release of P, (an irreversible process under initial velocity conditions) prior t o combination with 0, but result instead from the occurrence of some other irreversible first-order process between the binding of the two substrates during turnover. T h a t is to say the amino acid oxidase mechanisms are sequential rather than pingpong, in that all the elements of both substrates exist transiently in a single complex or transition state before the release of either PI or H,Oz. The rapid kinetic studies to be described in Section II1,C clearly show this to be the case. An interesting case of intersecting lines is given by D-amino acid oxidase and p-chloroalanine (39). Here, reversibility between the points of combination of S and 0, is afforded by the independent elimination pathway. Furthermore, since the (po and (pl values for the two pathways are equal, the point of intersection (equal to +o/+l) occurs on the abscissa of the double reciprocal plot. Lactate oxidase and chlorolactate (40) behave similarly and for probably the same reason. d. Inhibition by Substrate. L-Amino acid oxidase is severely inhibited by high concentrations of certain substrates (38).Although the mechanism of this inhibition is now understood (see Section IV), the necessary steady-state rate equations are not easily formulated in + coefficient form. e. The Integrated Steady-State Equation. When the integrated steadystate rate equation can be used, as in the glucose oxidase reaction, the 38. V. Masaey and B. Curti, JBC 242, 1259 (1967). 39. D. J. T. Porter and H. J . Bright, Proc. Znt. Symp. Flavins Flavoproteins, 6th (1975) (in press). 40. C. Walsh, 0.Lockridge, V. Massey, and R. Abeles, JBC 248, 7049 (1973).

432

HAROLD J . BRIGHT AND DAVID J. T. PORTER

conditions of each experiment (S >> 02) force each trace to obey Eq. (9)

Integration gives Eq. (10) : 1 [Ozlo -In-=---t [Ozl

A [Ozlo - [Ozl I [ET] B t B

(10)

+

Comparison with Eq. (5) shows that B = & and A = &, &/ [S]. Plots Although the conditions for of A vs. l/S will therefore yield (p0 and the valid use of Eq. (10) are strict (no product of any kind must interact reversibly with any enzyme species), the greater precision of integral, as opposed to differential, data allows for very accurate evaluation of the steady-state parameters.

C. TRANSIENT-STATE KINETICS : THEHALF-REACTIONS The transient-state kinetics of flavoprotein oxidase reactions, as well as certain enzyme-monitored steady-state kinetics (see Section III,D,2,a) are most commonly studied by stopped-flow spectrophotometry. Temperature-jump and other relaxation measurements cannot easily be applied to flavoprotein oxidase half-reactions because of their unfavorable thermodynamics. Excellent reviews of the capability and limitations of the stopped-flow technique are available (41,42). 1. Reductive Halj-Reaction The reductive half-reaction (RHR) is carried out anaerobically in the stopped-flow spectrophotometer by mixing E, with S and monitoring at a wavelength a t which the signal from the transient enzyme species of interest is maximal. In order to obtain AA values of 0.1 or so, the concentration of enzyme-bound flavin after mixing must be M or greater. This is about lo5 times greater than that required in a typical SSK (0,) experiment. Two wavelengths are commonly used, namely, 550 nm for the amino acid oxidases and 450 nm for glucose oxidase. In the case of the amino acid oxidases, three spectral states of the enzyme are observed in the RHR (36,38,43-46) namely, fully oxidized (E, 41. Q.H. Gibson, “Methods in Enzymology,” Vol. 16, p. 187, 1969. )

42. H. Gutfreund, “Enzymes” Physical Principles.” Wiley, New York, 1972. 43. V. Massey, H. Ganther, P. E. Brumby, and B. Curti, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 335. Wiley, New York, 1965. 44. K. Yagi, Advan. Enzymol. 34, 41 (1971). 45. T. Nakamura, S. Nakamura, and Y. Ogura, J. Biochem. (Tokyo) 54, 512 (1963). 46. D. J. T. Porter and H. J. Bright, BBRC 36,209 (19691

7.

433

FLAVOPROTEIN OXIDASES

and E, * * S), free fully reduced (E,), and a transient intermediate having long wavelength absorbance which, for the purpose of this discussion, is considered to be E, * * P,. The formation and decay of E, * P, is monitored a t 550 nm, at which wavelength none of the other species absorbs. Typically, the rise and fall of 550 nm absorbance following mixing of E, and S occurs as two exponential processes, the first of which can be saturated by S. This behavior is most simply explained by the scheme of Eq. (11):

-

-

-

If the rise in E, by Eq. (12) :

Eo*..SLE,...P1-tEr+P~

(11)

k-1

k-1

-

ka

kn

ki

E,, S f

PI is a single exponential process, this is described ki(k2

kobs =

kl[Sl

+ k-z)[Sl + + kz + k-2 +

k-ik-2

(12)

k-1

In most cases, the plot of l/kObsvs. 1/S is linear, indicating that either Ic-, or k-, (or both) is effectively zero. It should be noted that this condition is both necessary and sufficient to cause double reciprocal plots of steady-state data to be straight and parallel in the event that E, * * PI reacts with 0,. In the case that k-, is zero, the abscissa intercept of the double reciprocal plot is k,/(k-, k,) while the ordinate intercept is k2-l, Eq. (13) :

-

+

When k-, and k.., are nonzero (as judged from the nonlinearity of the double reciprocal plot of k , b s vs. S), Eq. (12) can be rearranged to Eq. (14) : cd

1 koba

- b/d

=-

da

- bc

+-d a d2- bc (h)

+

(14)

+ +

where a = k,(k, Ic-,), b = I c - , h ~ ~ c, = k,, and d = k, k-, k-,. Although plots of Eq. (14) obtained by curve-fitting procedures will be linear, there will be no correlation of the slope with the slopes obtained from plots of SSK (S/P) data. Fortunately, nonlinear plots of double reciprocal R H R data are rare. The decay of 550 nm absorbance in the second exponential from the RHR experiment is independent of the concentration of S and, as shown by double mixing experiments (43))is unaffected by a structurally differ* PI can be made ent substrate. Provided that the formation of E, sufficiently rapid (in order to get total conversion of E, to E, * * PI),

- -

-

434

HAROLD J. BRIGHT AND DAVID J . T. PORTER

the extinction coefficient of E, * ' * P, and the value of k , can be easily measured. Special cases of Eq. (11) are often encountered. First, saturation, of E, * P, formation by S may not be achieved. This may result from large values of k, and/or (k-l k,)/kl [see Eq. (13)], from limited substrate solubility, or from combinations of these factors. This feature is commonly found in flavoprotein oxidase reactions (see Section 5,A,2,a). It is not likely to result from the condition k, [S] << k? >> k-l (25) because kobais associated with a substrate deuterium kinetic isotope effect ( 4 6 ) . Therefore, when l/k2 is less than the mixing time of the stoppedflow apparatus (- 2 msec), whether or not substrate solubility is a limiting factor, the RHR in such cases is best described by Eq. (15) :

-

+

Second, E, * * P, may escape detection entirely if k , > > k,, in which case the RHR must be monitored a t 450 nm. This appears to be the case with basic substrates of D-amino acid oxidase (44). I n the glucose oxidase RHR only the fully oxidized (E, E, * * S) and fully reduced species are observed spectrophotometrically (25,47,48). E, * PI, if it has a significant lifetime, must have the same spectrum as E,. Consequently, the RHR is always monitored at 450 nm where E, and E, * * S absorb maximally relative to E,. Exponential decay of 450 nm absorbance is observed. If this process can be saturated by S, it is explained most simply by Eqs. (13) and (16).

+

-

-

E,

+S

6. I

k-i

E,

lc2

. . . S + Er

+ PI

(16)

Whether the rate of dissociation of P1from E, * * * PI is actually very rapid, as is implied by Eq. (16), is not known. Stopped-flow monitored turnover experiments (see Section III,D,2,a) suggest that it may occasionally be significant in catalysis. Most substrates do not show saturation behavior in the RHR, and they behave as if the bimolecular collision of E, and S were the rate-determining process in flavin reduction, Eq. (17).

+ S 4k Er + Pi

(17) However, the values of k, (spanning the range from 3 M-' sec-l to lo4 M-' sec-l) indicate that this is not a diffusion-controlled process and that there must be more than one event preceding flavin reduction [see Eq. (15) and Section V] . E,

47. T. Nakamura and Y. Ogura, J . Biochem. (Tokyo) 52, 214 (1962). 48. S. Nakamura and Y. Ogura, J. Biochem. (Tokyo) 63,308 (1968).

7.

435

FLAVOPROTEIN OXIDASES

2. Oxidative Half-Reaction

Oxidative half-reaction (OHR) is carried out by mixing E, (usually prepared anaerobically by reducing E, with a slight excess of S) with 0, and monitoring the appearance of oxidized enzyme a t 450 nm. I n all cases, the stopped-flow measurements conform to a simple bimolecular process with no evidence for saturation (although it should be noted that 0, concentrations are ordinarily limited to about 0.5 mM in the OHR and would therefore be insufficient for the detection of a labile E, * * 0, complex) (%5,S6,38,39,48).

+ O t 3 Eo(H1O~)

( 18) PI in the glucose oxidase RHR, the presence As is the case with E, and lifetime of E, * H 2 0 , is not detected in a OHR experiment because there are no spectrophotometric changes following the oxidation of flavin. This species, or an intermediate kinetically equivalent to it, can be detected, however, in enzyme-monitored turnover experiments (see Section

- -

- -

E,

+

III,D,2,a).

D. KINETICMECHANISM 1. Correlation of Steady-State and Transient Kinetics At this point, the results from the stopped-flow measurements of the

RHR and OHR, in their most general form, can be summed to construct the following catalytic cycle [Eq. (19) ] : kr

E,

\/

-+

h ki

k2

S - - E , . . . S ~ E , * .

1

,.

H24

Er

Whether or not this cycle accurately represents the turnover mechanism can only be determined by examining each of the steady-state 4 coefficients [Eqs. (5) and ( 8 ) ] in turn to see whether they correspond quantitatively to single rate constants (or functions of rate constants) as measured directly in the rapid reaction studies of the half-reactions. The coefficient of the steady-state rate Eq. ( 5 ) is easily identified in all cases because it is numerically equal to the slope of the double reciprocal plot of the RHR data [Eq. (13) or (15)]. These identities between the transient and steady-state kinetic parameters definitely

436

HAROLD J. BRIGHT AND DAVID J. T. PORTER

establish Eq. (11) or (16) as an obligatory reaction sequence in turnover. They also establish that is independent of enzyme concentration, at M over which the steady state least through the range from to and transient kinetics are routinely measured, and that there is one FAD per active site. Such uniform correlation with the rapid kinetic measurements is not, however, observed with the +o and terms. In the case of glucose oxidase, +z-l= k4 for all substrates (25). This krIOzl ide'ntity establishes E, E,(HzOz) as an obligatory reaction in turnover. With 2-deoxyglucose, 40-l = kz, showing that flavin reduction is the sole rate-determining first-order turnover process. The kinetic mechanism for this substrate is therefore accurately represented by Eq. (19), with the probable qualification that ka >> kz. For mannose, xylose, and galactose, which fail to saturate either in turnover or in the RHR, the kinetic mechanism is simply Eq. (17), with 41-1= k,. Glucose presents special problems. Its dl-l and $2-1 coefficients are represented by k, and kd, respectively, but the rate-determining first-order process or processes making up 4o-l cannot be detected in either the RHR or OHR (65). Solutions to this problem are provided by stopped-flow measurements of the behavior of oxidized enzyme species in turnover, which are described in Section III,D,2,a. In the case of D-amino acid oxidase, and the nonbasic a-amino acids, there is no quantitative correspondence between +o-l and +2-1 on the one hand and k, and k, of Eq. (19) on the other ( S S , 4 5 ) . The maximum turnover number, +o-l, for example, is usually a t least an order of magnitude greater than k, but is considerably smaller than k,. This rules out E, as an obligatory turnover intermediate and forces the conclusion that 0, must react directly with E, * * P, and that the dissociation of P, from E, * * * PI must be the rate-limiting first-order process in turnover, Eq. (20), loop B.

+,

-

-

-

Eo * * P,

7.

FLAVOPROTEIN OXIDASES

437

The pathway of loop B can be verified by double-stopped flow experiments (see Section 111,D,2,b). L-Amino acid oxidase is interesting because, depending upon experimental conditions, either loop A or loop B of Eq. (20), or both, can be operative ( 3 3 , S S ) . This behavior, which reflects competition between k 6 [ 0 2 ]and k,, gives rise to curved parallel line steady-state patterns with 0, as variable substrate (33). The rate constant k , is pH-dependent, being small a t low p H values and much larger a t high pH values. Turnover a t high p H and moderate 0, concentrations, therefore, takes place through loop A of Eq. (20). Under these conditions +,,-l = k3 and +,-' = k,. At much higher 0, concentrations, loop B becomes dominant and +o-l = k , and +*-l = k,. The curved region of the double reciprocal plot of the steady-state data with 0, as variable substrate corresponds to the condition when both loops of Eq. (20) are important. At lower pH values, the transistion from loop A to loop B occurs a t lower 0, concentrations because the value of k , is reduced. These examples illustrate how either complete or partial turnover mechanisms can be constructed by taking together the results from transient- and steady-state kinetic experiments. Neither type of experiment alone is sufficient to determine mechanism. I n those cases where the turnover mechanism is still undefined (e.g., glucose as a substrate for glucose oxidase) , stopped-flow monitored turnover experiments can be of great help, as discussed in the following section. 2. Confirmation of Mechanism by Further Rapid Reaction Measurements

a. Enzyme-Monitored Turnover [ S S K ( E ) ] .We have noted that the three term steady-state rate equation, Eq. ( 5 ), requires a t least two bimolecular processes (responsible for +1-1 and +,-l and involving S and O,, respectively) and at least one unimolecular step which is rate limiting at saturating S and 0, (and which is evaluated as +o-l). Transient kinetic studies of the RHR and OHR lead, directly or indirectly, to the location and characterization of all such turnover processes in the amino acid oxidase reactions [see Eq. (20)]. However, in the case of glucose oxidation by glucose oxidase, only and +2 were so characterized through the halfreaction studies because first-order processes associated with spectrophotometric changes could not be detected in the half-reactions. The problem of +o was solved by Gibson et al. (25) using an elegant and highly precise method first developed by Chance in his early studies of peroxidase (49). In this, the SSK(E) method, the substrate dependence of certain enzyme species is monitored during turnover. Although high enzyme concentrations (M ) and, consequently, rapid reaction techniques are required, the method is based on steady-state rather than transient-state 49. B. Chance, JBC 151, 553 (1943).

438

HAROLD J. BRIGHT AND DAVID J. T. PORTER

analysis because at least 10 or so turnovers are involved and the rate of change of substrate and product concentrations greatly exceeds that of any enzyme species. The SSK(E) method equates the area swept out a t any time by one or more absorbing enzyme intermediates during turnover (in the plot of absorbance vs. time) to the amount of limiting subtrate which has been consumed at that time. We illustrate the experiment in the case of glucose oxidase and glucose. Assume that each of the half-reactions is terminated by a single first-order process such as product release as shown in Eqs. (21)* kr

-

ks

41-1

Eo+S-Er*..P1+Er+P1

Eo

kk=+i-1

E,

$01-

.

ks *

*

H2024

Eo

+ HzOz

If [S]>> [02], the steady-state Eq. (22), with E,, = E, will hold through the entire turnover experiment.

+ E,

dt

Provided that the extinction coefficients of Eo and Eo . identical, Eq. (23) will relate the area, A t , swept out by E,, a t time, t , to the amount of O2which has been consumed a t time, t.

Equation (23) can be used in two ways. First, the turnover number v/[ET] can be computed at any concentration of 0, by the method of rectangular approximation suggested by Gibson et al. (25). These data generate excellent parallel line double reciprocal plots yielding +, and c$*. In addition to its precision, this method also has the advantage of establishing whether or not the turnover kinetics in stopped-flow experiments of all types M enzyme) are identical to those obtained by SSK(S/P) methods M enzyme). Second, and more importantly for this discussion, Eq. (23) allows computation of the turnover number of the “oxidized fraction of enzyme” alone. Plots of At,t,~/[Oz]o (in units of sec) versus a series of l/[S] should have slope = l/k, and ordinate intercept = l/k5. If +, = 1/k6, then i t is concluded that k3>>k6 and the process E, H,Oz+E,+H,Oz (or some other kinetically equivalent first-order decay of oxidized enzyme) is the only rate-determining first-order process in turnover. If +,, > l/ks (and if

-

7.

439

FLAVOPROTEIN OXIDASES

l/k5 # 0) both k , and k , are kinetically significant and k, can be evaluated from Eq. (24) :

If l/k, = 0, then k , is the only rate-determining first-order process in turnover. The following conditions, in general, must be fulfilled for valid application of the SSK (E) method. 1. One of the substrates must be in sufficient excess over the other (the limiting substrate) that the concentration of the former remains essentially constant during the complete turnover experiment. 2. All enzyme species which react with the limiting substrate must be spectrally distinct from those which react with the substrate in excess, and all of the latter species must have the same extinction coefficient at the wavelength of interest. 3. No product must accumulate to a concentration greater than about one-tenth of the value of its dissociation constant for interaction with any of the enzyme species. 4. If a branch point exists in the turnover mechanism none of the steps originating a t the branch-point species must be a bimolecular process involving the limiting substrate.

The last condition is best illustrated by Eq. (25), which'is an accurate representation of the L-amino acid oxidase reaction under certain conditions (33).

J

Eo . * P,

\P*1

k, EO

kr[Sl

-

/

\Odk

~

E,**.P,

(25)

Er

The total area swept out by the oxidized species E, and E, * * P is given by Eq. (26) and is clearly not simply proportional to the amount of 0, consumed.

The SSK(E) method is therefore invalid for the L-amino acid oxidase reaction when both loops A and B of Eq. (20) are operative. It should be noted that substrate inhibition (which is also highly characteristic

440

HAROLD J. BRIGHT AND DAVID J. T. PORTER

of the L-amino acid oxidase reaction because of the formation of S) will not by itself invalidate the SSK (E) method. E, * * The SSK(E) method has been successfully applied to the glucose oxidase reaction (25,50,51) [loop A of Eq. (20) operative] and to the D-amino acid oxidase reaction (37) (loop B operative). The equation appropriate for the latter case is the following [Eq. (27)] :

-

-

In this case, E, * * P, was monitored at 550 nm and this trace used to construct the area corresponding to E, PI plus E,, assuming, justifiably, that the concentrations of E, * * S and E, are negligible. This example also illustrates the need to subtract the area resulting from any oxidized species present a t reaction equilibrium or pseudo-equilibrium (resulting from slow hydrolysis of PI, for example) in order to obtain A t in Eq. (23). However, flavoprotein reactions are highly irreversible, thermodynamically, and this correction would rarely be required. The SSK(E) results of Massey et al. (37), which were claimed to be in good agreement with SSK(S/P) results, should be compared with those of Shiga and Shiga (52),which show that the turnover number of D-amino acid oxidase is dependent on the enzyme concentration. b. Double Stopped-Flow Measurements. The double stopped-flow (DSF) method, despite its potential, has not been used, to our knowledge, in flavoprotein. oxidase reactions, although Massey et al. (43) generated E, * * * PI from alanine in the D-amino acid oxidase reaction and then reacted this by a second manual mix with methionine in order to demonstrate that a second amino acid (chosen in this case to be kinetically distinct from the first) is not involved in the conversion E, * PI + E r +P,. Three important applications of the DSF method are briefly discussed here. * PI with 0, in the First, the evidence for the direct reaction of E, * amino acid oxidase reactions, though compelling (36,45),is indirect. Unlike E,, E, * * * PI is not stable in the absence of oxidizing agents, being generated as a transient intermediate. This problem can be met in a DSF experiment, however. If E, * PI is genexated anaerobically in the first mix and then reacted in the second mix with 0, and a tightly binding inhibitor which reacts specifically, irreversibly, and rapidly with E, [such as benzoate (43) in the case of D-amino acid oxidase), turnover is

-

-

6

- -

-

50. F. R. Duke, M. Weibel, D. S. Page, V. G. Bulgrin, and J. Lut,hy, JACS 91, 3904 (1969). 51. H. J. Bright and Q. H. Gibson, JBC 242,994 (1967). 52. K. 'Shiga and T. Shiga, BBA 283, 294 (19721

7.

441

FLAVOPROTEIN OXIDASES

quenched after one cycle and the following scheme applies after the second mix [Eq. (28) 3 :

The rate of disappearance of E, is given by Eq. (29) :

- -

P, a t 550 nm after the second mix

Since k , can be independently evaluated from RHR experiments, plots of Eq. (29) permit calculation of k, and also, from the magnitude of the first term, provide a measure of the reversibility of the RHR. It should * I resembles E, spectrally and is be noted that I is chosen so that E, entirely distinct spectrally from E, * * P,. Second, the DSF method can be used as follows to evaluate kl,a step which was again only indirectly deduced (36,455)except for the case of SSK(E) measurements of the glucose oxidase reaction ( 2 5 ) . I n this case, a tightly and rapidly binding competitive inhibitor for E, is chosen whose I is spectrally distinct from E,. Anthranilate (19) would complex E, be such an example for D-amino acid oxidase. Again, E, * PI is generated in the first mix and is then reacted with anthranilate and very high 0, in the second mix. Only E, * PIis present a t the end of the second mix under these conditions, and the rate of formation of E, * I (measured a t 550 nm) will correspond to the rate of conversion of E, * * * PI to E, P, [Eq. (30) 1. The rate constant, k,, may therefore be measured directly.

- -

-

-

-

- -

+

&-..I-%

fast

Third, the DSF method can be used to establish the locus of H,02 Plis release in loop B during turnover [Eq. ( 2 0 ) l . To do this Er * generated in the first mix and then mixed in turn with 0, and a rapidly

-

442

HAROLD J. BRIGHT AND DAVID J. T. PORTER

responding indicator for H,O, such as horseradish peroxidase. The kinetics of complex I of peroxidase are monitored a t 375 nm and compared with computer-generated traces of complex I formation which allow for H,O, release a t either k, or k,, the latter rate constants having been already evaluated for DSF experiments as described above. The measurements clearly showed (39) that H,O, is released in the k, step of Eq. (20) *

E. COMPUTER SIMULATION STUDIES The complexity of the entire multistep turnover mechanism [e.g., Eq. (20)] is such that computer studies are a useful, and sometimes necessary, adjunct to the intuition and experimental expertise of the investigator. The computer may be used during the collection and primary analysis of the kinetic data (41) or it may be utilized almost as an independent experimental method by which the pieced-together mechanism can be tested under stringent, and perhaps novel, experimental conditions. Chance was the first to use this method in this capacity during his classic studies of peroxidase (49) while Gibson et al. were probably the first to apply it to flavoprotein oxidases ( 2 ~ 5 ~ 3As 6 ) .a rule, the strategy is to determine whether the complete mechanism and its associated rate constants (both of which, as we have illustrated, are likely to have been deduced in piecemeal fashion from a series of more or less direct measurements under a variety of experimental conditions) is capable of regenerating, with reasonable precision, the time course of all experimentally observable substrates, products, and enzyme species in both half-reactions and turnover experiments (~C5,~6,SS,S6,~8,5l). It is, of course, rare, with a given substrate and experimental conditions, that all of the individual steps in a complicated mechanism will be partially rate-limiting during all, or even a part, of the turnover reaction. This condition was closely met, however, in computer simulation studies of the turnover of L-amino acid oxidase in which both loops of Eq. (20), together with substrate inhibition, the nonenzymic conversion of PI to PZand tautomerization of PI, were shown to satisfactorily reproduce the time course of E, * * * PI and O2(33). L-Amino acid oxidase provides another interesting example of the usefulness of computer simulation. I n this case, the importance of the reversibility of E, * - - PI E, PI was not k-a apparent from either SSK(S/P) or R H R reactions because, for different reasons, the transient buildup of PI (which hydrolyzes nonenzymically to Pz) does not occur in these experiments. However, PI accumulates readily to the level of k9/k--3 in SSK(E) experiments, with the result that the calculated time dependence of E, * PI in turnover, predicated on k-8 = 0, is greatly in error. This problem was quickly revealed by

+

+

7.

FLAVOPROTEIN OXIDASES

443

computer simulation studies and, subsequently, solved quantitatively (33,63).The redundance, or otherwise, of certain steps originally deduced from the behavior of substrates of different structure (e.g., heavy isotope substitution in the case of 2H) can be easily tested by simulation (51). Likewise, spikes and other notable characteristics of SSK(E) oscilloscope traces are highly diagnostic of mechanism and ought to be capable of simulation by any credible mechanism (33,SS). I n general, therefore, there is no uniform strategy for the application of simulation studies to flavoprotein oxidase mechanisms. Nevertheless, it is self-evident that complex turnover patterns from SSK(E) experiments in particular, though amenable usually to qualitative interpretation on the part of the investigator, can and should be subjected to quantitative simulation in order to demonstrate the adequacy, if not the uniqueness, of the derived mechanism. When such simulation studies have been satisfactorily carried out over as wide and as stringent a range of conditions as possible, the mechanism upon which they are based [having been typically deduced from a progressive correlation of SSK (S/P) , RHR, OHR, and SSK(E) experiments] can be taken as a reasonable working hypothesis until such time as new experimental data render it untenable.

F. SUMMARY OF MAJORKINETICRESULTS The kinetic behavior of the flavoprotein oxidases conforms to a single mechanism, as shown in Eq. (20). Differences in their behavior merely reflect relative differences in certain rate constants and are therefore = [H,O,] = 0) trivial. The complete steady-state rate equation ([PI] corresponding to Eq. (20) is the following [Eq. (31)] : [ETI 1)

-

I kr

+ k2 k-ik-2 +-k-IkikdS1 kik2(ka + kdOzl[Sl + kz(kak7 + ksks[021)kd[Oz]+ kzkskskr + k&6k7(kZ + k-z)[02] kZkrkSkr(k8 + kS[oZ])[oZ1 +

(31)

Equation (31), in its entirety, is never required in practice. Rather, special cases of it are characteristic of each of the enzymes and further q proximations of these special cases fit the behavior of specific substrates. The behavior of glucose oxidase is most simply explained if k, >> ka[O,]. Hence, loop A of Eq. (20) is operative, for which the steady state rate equation is Eq. (32) :

This conforms to the experimental rate law [Eq. ( 5 ) ] given by glucose. 63. D. J. T.Porter and H. J. Bright, BBRC 46,564 (1972).

444

HAROLD J . BRIGHT AND DAVID J. T. PORTER

However, the RHR and SSK(E) measurements show that k3 and k , are the only kinetically significant first-order processes in turnover and that Eq. (15) holds. Consequently, the behavior of this substrate is given by Eq. (33) :

With mannose, xylose, and galactose, neither S nor 0, saturates in turnover and the RHR experiments show that the second term of Eq. (33) entirely dominates the steady-state rate equation. 2-Deoxyglucose is the only substrate for which flavin reduction ( k , ) completely determines &, as shown by comparison of SSK(0,) and RHR results. Its behavior is expressed by Eq. (34) :

D-Amino acid oxidase, on the other hand, utilizes loop B of Eq. (20) (except in the case of basic amino acid substrates) because kG[02] >> k,. In this case, Eq. (31) becomes Eq. (35) :

However, the fact that the SSK double reciprocal 'patterns are parallel and straight [conforming to experimental Eq. ( 5 ) ] and that DSF experiments show that the reductive half-reaction is highly irreversible (as a result of small values of k-, and/or k-,) clearly demonstrates that the fourth (cross-product) term of Eq. (35) is negligibly small. The same conclusion was reached by Palmer and Massey (13). In the case of leucine and alanine, it can be conclusively shown that both k-, and k-, are effectively zero (see Section IV,A) . Very recently, it was shown (54) that arginine utilizes loop A of Eq. (20) and yields a rate equation which is probably identical to that given by glucose and glucose oxidase [Eq. (33) I. L-Amino acid oxidase exhibits a variety of kinetic patterns (depending on the substrate and experimental conditions) all of which can be accommodated by the full mechanism of Eq. (20). Thus with phenylalanine, both loops of Eq. (20) are operative at 2 5 O . However k-, is extremely small a t pH values greater than 6 (33) and Eq. (36) results:

This equation predicts curved, but parallel, double reciprocal plots of SSK(0,) data with 0, as variable substrate, as are actually observed 54. K. Yagi, M. Nishikimi, A. Takai, and N. Ohishi, BBA 341, 256 (1974).

7.

FLAVOPROTEIN OXIDASES

445

(3.3). At very high 0, concentrations (k,[O,] >> k,) , Eq. (36) reduces to Eq. (35) (with the fourth term being effectively zero), while a t very low 0, concentrations (k, >> ka[Oz]) Eq. (33) results. Both of these expressions conform to the experimental rate law of Eq. ( 5 ) . At Oo (38), or at very low pH values (SS), k-, and k-, become significant compared to k, and intersecting double reciprocal patterns of SSK(0,) data result. These conform to Eq. (35) and to the experimental rate law of Eq. (8). Thus, despite the bewildering variety of kinetic behavior exhibited by the flavoprotein oxidases, all of these enzymes conform to the general scheme of Eq. (20). IV. The Flavoprotein Oxidases: Molecular Properties and Kinetic Mechanism

A. D-AMINOACIDOXIDASE Purification of the benzoate-holoenzyme complex of D-amino acid oxidase from pig kidney is based on the procedure of Kubo et al. (55).Massey et al. (56) improved this method by the addition of calcium phosphate chromatography, and the procedure has been carefully documented by Brumby and Massey ( 5 7 ) .Some difficulties in obtaining homogeneous preparations by these methods have been noted (58).Recently, Curti et al. (59) have added a step involving DEAE-Sephadex chromatography of the benzoate-holoenzyme complex at pH 8.3. On the other hand, Tu et al. (60) have introduced a DEAE-Sephadex chromatography of the apoenzyme. Both of the latter preparations appear to be homogeneous on SDS gel electrophoresis. It should be noted that there is a slight discrepancy between the value of the extinction coefficient of the apoenzyme given by Curti et al. (59) and that reported by Tu et al. (60). Resolution of the benzoate-holoenzyme complex is accomplished as described by Yagi and Ozawa (61) or by gel filtration of a mixture of reduced enzyme and benzoate ( 3 9 ) . Resolution of the holoenzyme itself is best achieved by the method of Massey and Curti (62). 55. H. Kubo, M. Yamano, M. Iwatsubo, H. Watari, T. Soyama, J. Shiraishi, S. Sawada, N. Kawashima, S. Mitani, and K. Ito, Bull. SOC.Chim. Biol. 40, 28 (1958). 56. V. Massey, G. Palmer, and R. Bennett, BBA 48, 1 (1961). 57. P. E. Brumby and V. Massey, Biochem. Prep. 12, 29 (1968). 58. S. W. Henn and G. K. Ackers, JBC 244, 465 (1969). 59. B. Curti, S. Ronchi, U. Branzoli, G. Ferri, and C. H. Williams, Jr., BBA 327, 266 (1973). 60. S. C. Tu, S. J. Edelstein, and D. B. McCormick, ABB 159, 889 (1973). 61. K. Yagi and T. Ozawa, BBA 56, 420 (1962). 62. V. Massey and B. Curti, JBC 241, 3417 (1966).

446

HAROLD J. BRIGHT AND DAVID J. T. PORTER

The molecular weight of the monomeric form of pig kidney enzyme has been measured to be between 38,000 and 39,000 (59,600). The amino acid analyses of Tu et al. (60) and Curti e t al. (69) agree reasonably well. Curti et al. determined the N-terminal amino acid to be methionine and the C-terminal amino acid t o be leucine (69). These results confirm the conclusion of Kotaki e t al. (63). The ratio A274/A4BZwas determined to be 9.5 by Curti e t al. (59), compared to the value of 10.0 reported by Brumby and Massey (57). Chemical modification by sulfhydry1 group reagents (64) and by glyoxal have been reported (65). Recently, T u and McCormick (66) have modified holenzyme through photochemical activation of FAD. It was suggested that tyrosyl and cysteinyl residues are near the active site since loss of enzymic activity paralleled photooxidation of these groups. The fluorescence and absorbance spectra of the oxidized enzyme have been reviewed in detail (13). Recently, Ghisla e t al. (20) have found that the flavin of reduced enzyme has a fluorescence emission spectrum. Perturbation of the spectrum of the oxidized enzyme by small molecules is well known and has been discussed in detail by Yagi et al. (67) and Massey and Ganther ( 1 9 ) . Anthranilate and other enamine-like ligands are of particular interest because the spectra of their complexes with E, are very similar to the spectrum of the enzyme obtained during turnover of p-chloro-a-amino acids (see Section V,A,3,a). The binding of FAD to D-amino acid oxidase is demonstrably reversible with a dissociation constant of 5.35 X lo-’ M (60). Kinetically, FAD binding is a two-step process, the appearance of catalytic activity being correlated with the second step (6g).No evidence exists for (labile) covaIent interactions. The regions on the FAD molecule available for noncovalent interactions with the enzyme may be classified into three groups: (1) adenylate, (2) ribitol, and (3)the isoalloxazine. McCormick e t al. (68) demonstrated that the adenylate portion of FAD is practically essential for binding and catalytic activity. Substitution of this portion of the molecule by other purines and by pyrimidine ribofuranosyl groups resulted in FAD analogs which are essentially inert catalytically. I n addition, substitution of the ribofuranosyl group by deoxyribofuranosyl decreased coenzyme activity twofold and binding tenfold (68). Both the length of the flavin side chain and the presence of a 2‘63. A. Kotaki, M. Harada, and K. Yagi, J . Biochem. (Tokyo) 61, 598 (1967). 64, A. H. Neims and L. Hellerman, Annu. Rev. Biochem. 39,867 (1970). 65. A. Kotaki, M. Harada, and K. Yagi, J. Biochem. (Tokyo) 64, 637 (1968). 66. S. C. Tu and D. B. McCormick, JBC 248,6339 (1973). 67. K. Yagi, T. Oaawa, M. Naoi, and A. Kotaki, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 237. Univ. Park Press, Baltimore, Maryland, 1968. 68. D. B. McCormick, B. M. Chasay, and J. C . M. Tsibris, BBA 89, 447 (1964).

7.

FLAVOPROTEXN OXIDASES

447

hydroxyl group contribute to complex stability and coenzyme activity. Thus, the K , value for 2’-deoxyriboflavin-adenine dinucleotide is fivefold greater than that for FAD, and a flavin side chain of five carbons is necessary for coenzyme activity but not necessary for binding (69). The role of the pyrophosphoryl group in binding has not been studied, although resolution of the holoenzyme by 1 M KBr suggests that it may be important ( 6 2 ) . The importance of aromatic amino acids in the binding of flavin to proteins has been established by X-ray crystallographic studies in the case of flavodoxin, where it was found that tyrosyl and tryptophanyl residues are aligned plane-parallel to the isoalloxazine of F M N ( 7 0 ) .Tu and McCormick (71) have suggested that there is a tyrosine-tryptophan pair present a t the active site of D-amino and oxidase which contributes to the FAD-binding a h i t y through coplanar interaction with the isoalloxazine ring. The N-3 position of the isoalloxazine ring appears to be important for FAD binding since alkylation of this position decreases the ability of FAD to bind. In addition, deprotonation of N-3 also decreases the affinity of FAD, and Tu and McCormick have concluded that the N-3 proton is probably necessary for binding ( 7 1 ) . The importance of hydrophobic forces in the binding of FAD has recently been shown by Naoi et al. ( 7 2 ) .Using the FAD-competitive hydrophobic probe, 4-benzoylamido-4’-aminostilbene-2,2’-disulfonate,they found enhancement of its fluorescence when bound to the apoenzyme. I n solution, an intramolecular complex between adenine and the isoalloxazine ring of FAD exists (73) as has been found for riboflavin and adenosine in the crystalline state (7’4). Striking homologies have been found in the topography of nucleotide coenzyme binding sites ( 7 5 ) . D-Amino acid oxidase undergoes a concentration-dependent dimerization (58,76),and there are various reports (62,77,78) that the turnover number is dependent upon the enzyme concentration. This presumably 69. B. M.Chassy and D. B. McCormick, B B A 110, 91 (1965). 70. R. D. Anderson, P. A. Apgar, R. M. Burnett, G. D. Darling, M. E. LeQuesne, S. G. Mayhew, and M. L. Ludwig, Proc. Nut. Acad. Sci. U.S. 69,3189 (1972). 71. S. C. Tu and D. B. McCormick, Biochemistry 13, 893 (1974). 72. M.Naoi, A. Kotaki, and K. Yagi, f. Biochem. (Tokgo) 74, 1097 11973). 73. M.Kainosho and Y. Kyogoku, Biochemistry 11, 741 (1972). 74. D.Voet, and A. Rich, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 23. Univ. Park Press, Baltimore, Maryland, 1971. 75. M. Buehner, G.C. Ford, D. Moras, K. W. Olsen, and M. G. Rossmann, Proc Nut. Acad. Sci. U . S . 70, 3052 (1973). 76. E . Antonini, M. Brunori, M. Rosaria Brurzesi, E. Chiancone, and V. Masaey, JBC 241, 2358 (1966). 77. K. Shiga and T. Shiga, A B B 145,701 (1971). 78. K. Yagi, N. Sugiura, and H. Ohama, J . Biochem. (Tokyo) 72, 215 (1972).

448

HAROLD J. BRIGHT AND DAVID J. T. PORTER

reflects the fact that the turnover number of the monomer is different from that of the dimer. Yagi et al. (79) found that the enzyme is completely dissociated by 2 M urea and that the V,,, value for the monomer is greater than that for the dimer. Their results were somewhat obscured by the fact that urea is a competitive inhibitor of D-alanine and that a complete kinetic analysis of the monomeric enzyme reaction was not carried out. Nonetheless, the fact that the value of Vmax,app in an airequilibrated solution was five times greater for the monomer than for the dimer suggests that the rate differences reside in IC, and/or k , of Eq. (20) since these are the processes which control the turnover number in such an experiment. The finding by Massey et al. (37) that V,, is constant over a wide range of enzyme concentration suggests either that k , may be the step which is slowed in the dimer reaction or that dissociation of the dimer was insignificant under their conditions. It would seem prudent, in view of the uncertainties involved, to obtain steady-state data for this enzyme from SKK(E) experiments if these are to be correlated with transient kinetic data, since the enzyme concentrations required in the two kinds of experiment are comparable. Regardless of the enzyme concentration, the steady-state rate equation has three terms, corresponding to parallel line double reciprocal patterns [Eq. ( 5 ) ] . Such studies have been mostly with alanine (36,80,81). Dixon and Kleppe (80) interpreted the parallel line patterns to mean that the enzyme cycled in ping-pong fashion between E, and E,. As pointed out in Section I11 no such conclusion can be drawn from SSK(0,) data alone. Furthermore, both Massey and Gibson (36) and Nakamura et al. (45,82) had already conclusively shown by rapid reaction measurements that the enzyme must oscillate between E, and E, * PI [Eq. (20)] with the susbstrates tested. The steady-state rate equation corresponding to the general scheme for this enzyme [Eq. (31)] reduces to the experimental rate law of Eq. ( 5 ) when k , is effectively zero and when k-, or k-,, or both, are very small. Double stopped-flow experiments (see Section III,D,2,b) confirm the latter condition. Substitution of &-deuterium for a-hydrogen in valine resulted in a kinetic isotope effect of 2-fold on with little effect on either &, or #2 (83).These results substantiate the conclusion that flavin reduction (involving C-H bond scission in the substrate) is usually not a significant contributor to the term in flavoprotein oxidase turnover. Systematic

- -

+,

79. K. Yagi, N. Sugiura, H. Ohama, and N. Ohishi, J . Biochem. (Tokyo) 73, 909 (1973). 80. M. Dixon and K. Kleppe, BBA 96,368 (1965). 81. J. F. Koster and C. Veeger, BBA 151, 11 (1968). 82. T. Nakamura, J. Yashimura, and Y. Ogura, J . Biochem. (Tokyo) 57, 554 (1965). 83. K. Yagi, M. Nishikimi, A. Takai, and N. Ohishi, BBA 321, 64 (1973).

7. FLAVOPROTEIN

OXIDASES

449

studies of the pH dependence of the steady-state coefficients have not been reported. Reductive half-reaction experiments with this enzyme, as with L-amino acid oxidase, have been predicated on the long wavelength absorbance of E,* *PIwhich was first detected in a flavoprotein reaction by Beinert (84). Although the chemical identity of E,;.-P1 is not fully established (see Section V,A,B,b), there is a reasonable consensus concerning the kinetics of E;-*PI in this reaction (36,45,. The rise of E;--P, after the anaerobic mixing of Ea and S occurs as a single exponential ( k a b s ) at 550 nm, as does its subsequent decay ( k 3 ) .The value of k, is entirely independent of S. The small amount of biphasic character in k o b s (85) has now been interpreted as resulting from the presence of the monomer (86). Double reciprocal plots of kobs vs. S,except in the case of a series of phenylalanines (87‘), seem always to be linear. I n the case that a finite value of kobs is achieved a t infinite S (e.g., leucine) the entire RHR experiment is described b y Eq. (12). Linearity of kzt vs. S-’ indicates th a t k-z or k-1, or both, are very small. Thus, the linearity of this plot is determined by the same factors which cause the parallel line patterns in the case of SSK(02) data. The RHR experiments involve the production only of a n active site equivalent of P,, and k_,[P,] is inoperative; this is evidenced by the fact that AAE5,,returns to its zero time value a t the end of the experiment. During turnover, however, PI may accumulate transiently a t concentrations approaching or exceeding k 4 k 3 , in which case k-JP1] becomes significant (53). Saturation of kobs is often not observed, indicating th at k2 is very large or th a t the apparent affinity of S ((k-1 k 2 ) / k l ) is weak (or that both of these factors are operative). A third pattern of behavior is seen with the basic amino acid substrates where E, . . PI does not accumulate and the R H R must be monitored a t 450 nm (54). This result is most simply explained by the condition ka >> k2. The RH R results in general point to two important conclusions regarding the kinetic mechanism for the overall reaction. First, k, is usually much too large to be a significant contributor to Q ~ - I . I n the opposite sense, k , (except in the case of the basic amino acid substrates) is far too small to allow E, to be an obligatory intermediate in turnover. Consequently, the first-order process (es) controlling the maximum turnover rate must be located in the oxidative pathway (36,45).

-

+

84. H. Beinert, JBC 225, 465 (1957). 85. V. Massey, G. Palmer, C. H. Williams, B. E. P. Swoboda, and R. H. Sands,

in “Flavins and Flavoproteins” (E. C. Slater, ed.), p. 133. Elsevier, Amsterdam, 1966. 86. K. Yagi, M. Nishikimi, and N. Ohishi, J . Biochem. (Tokyo) 72, 1369 (1972). 87. J. G. Voet, D. J. T. Porter, and H. J. Bright, unpublished results.

450

HAROLD J. BRIGHT AND DAVID J. T. PORTER

The effect of a-deuterium substitution in the amino acid substrate on for E,..-P1 formation in the RHR depends on the substrate used. Furthermore, since the deuterated substrate is usually the DL racemate, competitive inhibition by the L isomer has to be considered. Thus, Ki for L-leucine is 1.4 mM (88). However, provided that the ratios of kobs are obtained with the racemates of both the deuterated and nondeuterated substrates, valid kinetic isotope effects can be measured. There are several interesting cases. With leucine and valine, the saturating value of k o b s , corresponding to flavin reduction controlled by k,, gives a deuterium kinetic isotope effect of 3-fold (83).Norvaline, on the other hand, has a saturable kobsbut gives no kinetic isotope effect. This result strongly suggests that a two-step kinetic scheme for E; * .P, formation may be too simple and that Eq. (52) is required instead. I n Eq. (52), both E,***S and E,’**.S must resemble E, spectrally and C-H bond scission occurs in the third step. With valine, the second step is faster than the third, while with norvaline the opposite would hold. Other merits of Eq. (52) are discussed in Section V,A,2,a. The RHR behavior of leucine and alanine are also worth noting, since two separate lines of experimental evidence point to a common feature of their behavior. First, the leucine concentration dependence of k o b s is given by Eq. (37) : B 1 _ -A+(37) kobs

kobs

PI

Yagi (4.4) studied the pH dependence of A and B in Eq. (37) and found that A reflects the titration of a basic group in the enzyme while B is or both [see Eq. (12)] entirely independent of pH. Either k-, or must be zero in order for Eq. (37) to result from the RHR data. €3 can only be pH-independent if k, >>k-,. Furthermore, k, > k-, because the spectrum of the rate-limiting species in E, * * * P, formation has the character of oxidized, rather than reduced, enzyme. Therefore, the R H R data conform to Eq. (38):

We conclude, therefore, that both k-, and k-, are remarkably small in the case of leucine. This is also the case with alanine, since the linear double reciprocal plot of k o b s vs. a-deuterated alanine is parallel to that for a-protonated alanine (39).Arguments similar to those given for leucine show that this will only occur if both k-, and k-, are effectively zero. These RHR experiments, together with the double stopped-flow 88. K. Yagi, M. Nishikimi, N. Ohishi, and A. Takai, FEBS (Fed. EUT. Biochem. Soc.) Lett. 6, 22 (1970).

7.

FLAVOPROTEIN OXIDASES

45 1

measurements (see below and Section III,D,2,b) provide the explanation for linearity of the R H R double reciprocal plots and for the parallel line patterns obtained with SSK(0,) data [Eq. ( 5 ) ] . Koster and Veeger (81), on the other hand, have concluded from an analysis of SSK (0,) data taken in the presence of the competitive inhibitor benzoate that the rate at which E,- * .PI reverts to E, is 50 sec-l in the case of alanine. This value is too large to be consistent either with the linearity of R H R double reciprocal plots or with the parallel line patterns seen with SSK(0,) data (36). Consequently, we have directly measured the overall rate of the process E,...P, E, by means of double stopped-flow spectrophotometric experiments (39). The rationale of this method was given in Section III,D,2,b. In the first mix, E; - * P I was generated anaerobically from alanine and then mixed with benzoate (I) which is a specific competitive inhibitor reacting rapidly and irreversibly with E,. The subsequent decay of E , - * . P to E,*..I and E, was monitored a t 550 nm. When 0, = 0, the required expression is Eq. (39) [see Eq. (29) ] :

In 0.1 M potassium pyrophosphate a t 25" and pH 8.3, the value of kobs was 0.038 sec-l while k,, measured separately, was 0.017 sec-l. The value k-, k-l) was therefore 0.021 sec-l. This is the effective of k-,k-,/(k, rate constant for E, * P, + E, and clearly demonstrates the irreversibility of E , - . - P , formation. This result probably explains why Walsh et al. (31) failed to obtain anaerobic exchange of the a-H of alanine with solvent tritium. The rate of decay of E;..P, to E, (k,) is highly dependent on the nature of the substrate. Yagi has shown that k , varies over several orders of magnitude (44). The pH dependence of k , is remarkably similar for all amino acids and similar also to the corresponding dependence of k, in the L-amino acid oxidase reaction (33). The value of k , tends to lie on a plateau in the neutral pH range, but rises indefinitely as the pH is raised above 9. Many studies, through emphasis on pH 8.3 and substrates with small k , values (such as alanine), have tended to obscure the fact that like L-amino acid oxidase, D-amino acid oxidase can utilize both the A and B loops of Eq. (20) under appropriate conditions, although i t is true that, in general, the k , values for D-amino acid oxidase substrates [except for basic amino acids ( 4 4 ) ]tend to be smaller than those for L-amino acid oxidase. The value of k , is independent of a-deuterium substitution in the sub-

+ +

-

452

HAROLD J. BRIGHT AND DAVID J. T. PORTER

strate (44) as is the case also in the L-amino acid oxidase reaction (46). This suggests that the conversion of E,. *Pl to E, P does not involve the transfer, in any way, of the hydrogen originating a t the a-carbon of the substrate and thereby supports the contention that this process does indeed represent the release of imino acid from reduced enzyme. Under most conditions, and with most substrates, the value of k , is substantially smaller than +,,-'. This rules out E, as an obligatory intermediate in turnover under such conditions and prompted the early suggestion that E,. * .P1must react directly with 0, (36,45). Rao et al. (89) evaluated Ic,, governing the reaction of E, with 0, [see Eq. (20)] as 1.9 x lo4 M-l sec-' at pH 8.3 and 25O. Nakamura et al. (46), using the method of Chance (49), estimated that the rate of reaction of E r a * .PI with 0,, governed by k,, was 1.2 x lo5 M-l sec-l at pH 8.3 and 20° during alanine turnover. Massey and Gibson (361,through computer simulation of SSK(E) data, estimated a very similar value of 1.5 X lo5 M-' sec-'. None of these methods is direct, however, and we have obtained a value of Ic, from direct measurements in double stopped-flow spectrophotometric experiments (39). The design of such experiments is very similar to that used for measuring the irreversibility of the conversion E,. * .PI + E, [see Eq. (39)] except that 0, is present in this case and Eq. (29) applies. The rationale for this method was given in Section III,D12,b. Measurement of kObsof Eq. (29) as a function of O2 leads to a value of 1.7 X lo5 M-' sec-' for k,. This is in excellent agreement with other results (36,&,89) and confirms the validity of the mechanism of Eq. (20). Rao et al. (89) have shown recently that electron acceptors other than 0, will react with E,...P,and with E,.Ferricyanide and tetranitromethane were less reactive than 0, in oxidizing E, * -PI but more reactive than 0, in the oxidation of E,. Although the scheme of Eq. (20) is supported by data from a variety of laboratories, the nature and location of the first-order step(s) controlling maximum flux through loop B are not known with certainty. It has been suggested as release of PI from E, PI for two reasons (36,45). First, the value of &,-I is substrate structure-dependent. Second, the enzyme exists in an oxidized state during SSK(E) turnover experiments with saturating S and 02. However, the rate-determining process could also involve release of H20zfrom H202 * . E, * . PI or a weighted mixture of consecutive PI and H20z release. This question can also be answered by the double stopped-flow technique in an experiment similar in design to that required to directly measure k s (see Section III,D,2,b). * anthranilate complex is monitored a t 550 nm (19). Briefly, the E,

+

-

-

-

-

-

-

89. N. A. Rao, M. Nishikimi, and K. Yagi, BBA 276,350 (1972)

7.

453

FLAVOPROTEIN OXIDASES

E, . . PI is generated anaerobically in the first mix from alanine and then mixed with high 02 and anthranilate (at concentrations sufficient to convert E, t o E,-anthranilate very rapidly and completely). The rate ks0a of the process E, * PI E, . * PI is also very fast under these PI after conditions with the result that the enzyme is entirely E, the second mix and the transient E, * . . PI + E, is monitored simply anthranilate. The rate of formation of free E, a t p H 8.3 through E, and 25" was 9.8 sec-I, which agrees very well with = 10 sec-' obtained from SSK(Oz) experiments under the same conditions (39). These results confirm the assignment of k7 [Eq. (20)] as the single firstorder rate-determining process in turnover as well as its location in the pathway after the oxidation of E, * * PI by 02. However, they do not establish whether it is the release of HzOzor PI which is being measured. To answer this question, it is technically easy to trap H z 0 2with horseradish peroxidase and to monitor the appearance of complex I and I1 of this enzyme spectrophotometrically a t 375 nm (90). The pseudo-firstorder processes in the D-amino acid oxidase reaction were adjusted by control of S and O2 concentrations to give the following simplified scheme [Eq. (40)]:

-

-

-

1000 sec-'

sec\7j:i:: EO

10

(40)

Eo***P

If H,O, were released a t a step (a) in Eq. (40),a burst in complex I formation is predicted, whereas a lag is predicted if the point of release is ( b) . At pH 8.3 and 25O, there is a burst in H,O, formation (reaction initiated by mixing E, O2 peroxidase with S 0,) which, by computer simulation, can be shown to correspond exactly to the release of H,O, a t step (a) of Eq. (40) (59).Turnover of alanine and leucine are therefore accurately described, kinetically and chemically, by Eq. (41):

+ +

+

Attempts have been made to apply the Hammett linear free energy relationship to D-amino acid oxidase in order to determine whether flavin 90. B. Chance, ABB 22, 224 (1949).

454

HAROLD J . BRIGHT A N D DAVID J. T. PORTER

reduction is favored by electron withdrawal or donation to the substrate a-carbon. Neims et al. (91) measured the effect of ring substituents in series of phenylalanines and phenylglycines on the steady-state velocity of the D-amino acid oxidase reaction. The velocity measurements were not extrapolated to infinite 0, and were of the form of Eq. (42):

Comparison of the values of v a t different 0, concentrations clearly showed that +o is a function of the ring substituent. However, b0-' represents k, the dissociation of P, from E,. .P, [Eq. (41)], a t least in the case of the unsubstituted amino acids. The plot of log v vs. u was biphasic in the case of phenylglycines, with a p value of 5.44 for electron donating substituents and -0.4 for electron withdrawing substituents. The authors interpreted these results to mean that electron donating substituents slowed substrate a-proton removal whereas electron withdrawing substituents forced a change in rate-determining step from flavin reduction to product release. The only reliable method available to investigate substituent effects of this kind is to systematically measure, by RHR stopped-flow measurements, the effect of both ring substituents and a-deuteration on the kinetics of E, P, formation. We have carried out such stopped-flow studies with substituted phenylalanines and could find no interpretable relationship between log k, and u (87). It is clear that the rate of dissociation of E,. * *Plis highly sensitive to the nature of the R group of the product ( 3 6 ) . Furthermore, plots of log Ki versus u for a series of substituted benzoates (which specifically react with E, giving complexes analogous to E,. .PI) were biphasic (99) and approximately the mirror image of the results of Neims et al. (91). If this variation in stability of E,. . -1is caused chiefly by variation in the rate of dissociation of the E , * - . Icomplexes, such results would tend to confirm that the effect of substituents on the steady-state velocity (91) originates in E; -Pl(k7)rather than E,. .S (k?). One may ask whether, from our present state of knowledge at least, a-p studies on flavoprotein oxidases can even in principle provide reliable chemical information. Ideally, the substituent should exert its electronic effect on the rate-determining transition state between E,. * * S and E,-.-P, and should have no effect on the ground state energy level of E,.* -S. Unfortunately, this.appears not to be the case even with leucine and valine, where the difference in electron donating power of the R groups should be negligible. The k , values for leucine and valine a t pH 8.3 are 70 and 2000 sec-l, respectively, and both substrates exhibit deuterium kinetic isotope effects on this step (44). It seems highly likely that

-

--

-

-

91. A. H. Neims, D. C. DeLuca, and L. Hellerman, Biochemistry 5, 203 (1966). 92. J. F. Koster and C. Veeger, BBA 167, 48 (1968).

7. FLAVOPROTEIN

455

OXIDASES

the 30-fold difference in reactivity between these substrates originates in large part from unknown differences in ground- and transition-state binding interactions rather than from the differences in electron density F& the a-carbon of the bound substrate. It follows that kinetic substituent effects, even when these are correctly assigned to the process of interest (flavin reduction in this case), are not likely to resolve the question of whether the a-hydrogen of the substrate is removed as a proton, hydride, or hydrogen atom. A kinetically competent flavin- or protein-substrate adduct has never been trapped during the oxidation of physiological substrates, in contrast to the case of nitroalkanes (see Section V). Attempts to do so through the use of BH,- resulted in the reductive fixation of one flavin equivalent of alanine as r-N- (1’-carboxyethyl) -1ysyl-apoeneyme ( 9 3 ) .However, this labeled apoeneyme was subsequently shown to be catalytically active after the addition of FAD, thus demonstrating that neither the lysyl residue nor the (presumed Schiff’s base) adduct formed between it and the departing product is an obligatory participant in the catalytic mechanism (94). The latter conclusion is substantiated by the fact that the product, P,, is the a-imino, rather than the a-keto, acid (see below). The case for P, being the a-imino acid, rather than a-keto acid, is strong. First, Yagi et al. (4)measured transient pH changes during oxidative turnover of cyclic and noncyclic amino acids which corresponded to release of protons during turnover. However, in the case of noncyclic amino acid (leucine), proton release was quickly followed by proton uptake. Under the conditions used ( M enzyme, pH 8.3) the transient accumulation of a-imino acid corresponding to leucine would be favored, since the rate of enzymic synthesis is high and the pH is greater than the pK, value of the a-immonium group (thus diminishing the fraction of the reactive immonium species for the subsequent nonenzymic hydrolysis to a-keto acid and ammonia). In the case of proline, the cyclic a-imino structure is resistant to hydrolysis [see Eq. (43) 1. proline

co;

I

leucine

c=o + I co;

R~NH:

(43)

t R I C(H,NH,R‘) I

c0,93. D. S. Coffey, A. H. Neims, and L. Hellerman, JBC 240, 4058 (1965). 94. V. Massey, B. Curti, F. Miiller, and S. G. Mayhew JBC 243, 1329 (1968)

456

HAROLD J . BRIGHT AND DAVID J. T. PORTER

Hafner and Wellner (95) obtained direct evidence, by the recovery of racemic a-amino acid after borohydride addition, for the a-imino acid product. However, these results were not quantitative and they claimed that the major free product which accumulates transiently under these conditions is the carbinolamine. Subsequently, Porter and Bright (SO), using phenylalanine as substrate, showed that phenylpyruvate (Pz)formation was characterized by a lag period which increased as the pH was raised. Quantitative borohydride trapping experiments showed that this lag was almost entirely accounted for by the transient accumulation of a-imino acid, in agreement with the behavior of analogous imine-carbonyl interconversions.

B. L-AMINOACIDOXIDASE L-Amino acid oxidase is widely distributed (96-100), but the enzyme from snake venom (101,109) has received by far the most attention. The highest levels of ophidian L-amino acid oxidase are to be found in the rattlesnakes Crotalus adamanteus and Agkistrodon piscivorus, and the enzymes from these two sources appear to be very similar (101,103). Wellner and Meister (103) crystallized the C . adamateus enzyme and showed by sedimentation studies that it has a molecular weight of approximately 130,000. Three electrophoretically distinct components (designated A, B, and C in order of decreasing charge) were detected in variable amounts, but no catalytic differences were observed. The amino acid compositions of the A, B, and C species differ (104), and dissociation into two different polypeptide chains has been achieved (105) suggesting the possibility of the combinations aa, ab, bb. However, isoelectric focusing revealed a t least 18 isozymes (106). Despite these complexities, whose molecular bases are obscure, kinetic heterogeneity is not observed. The C . adamanteus enzyme has two tightly bound FAD molecules per mole holoenzyme and a characteristic FAD absorption spectrum (103). 95. E. W. Hafner and D. Wellner, Proc. Nut. Acad. Sci. U.S. 68, 987 (1971). 96. M. Nakamo and T. S. Danowske, JBC 241, 2075 (1966). 97. P. K. Stumpf and D. E. Green, JBC 153,387 (1944). 98. S. G. Knight, J . Bacterial. 55, 401 (1948). 99. J. Roche, P. E. Glahn, P. H. Manchon, and N. V. Thoai, BBA 35, 111 (1959). 100. J. Struck and I. W. Sizer, ABB 90,22 (1960). 101. T. P. Binger and E. B. Kearney, ABB 2.9, 190 (1950). 102. E. A. Zeller, Advan. Enzymol. 8, 459 (1948). 103. D. Wellner and A. Meister, JBC 235, 2013 (1960). 104. D. Wellner and M. B. Hayes, Ann. N . Y . Acad. Sci. 151, 118 (1968). 105. A. De Kok and A. B. Rawitch, Biochemistry 8, 1405 (1969). 106. D. Wellner, Annu. Rev. Biochem. 36,669 (1967).

7.

FLAVOPROTEIN OXIDASES

457

Two mechanisms have been proposed for the L-amino acid oxidase reaction and it is convenient to first consider that formulated by Wellner and Meister (107).Their experiments were confined to SSK(0,) measurements, and a major question which they sought to answer was the origin of the inhibition of high concentrations of certain substrates which is characteristic of this enzyme. The essence of their proposal was that the two FAD molecules act as a unit within a single active site. At low and moderate substrate concentrations, the enzyme was postulated to oscillate between E- (FAD) and E- (FADH) , while a t high substrate concentrations the enzyme oscillated predominantly in a second cycle between E- (FADH) and E- (FADH,) 2. If the postulated biradical E- (FADH) were oxidized more rapidly than E- (FADH,) 2, then turnover velocity would be predicted to be substantially inhibited when the oxidative processes were rate-determining (i.e., high S and low 02). Several objections to this mechanism were raised by Massey and Curti (38,108), of which the most cogent was the following. When the semiquinoid state [ E- (FADH)* ] of L-amino acid oxidase was generated anaerobically by irradiation in the presence of EDTA, it was found (in common with other flavoprotein oxidases) to be entirely inactive with reductive substrates. This evidence is both necessary and sufficient to rule out the Wellner-Meister mechanism. A variety of steady-state and rapid kinetic studies carried out subsequently has gradually resulted in the adoption of a mechanism [Eq. (2011 which involves one FAD per active site and which can explain the phenomenon of substrate inhibition as well as other features of the reaction. At 25O, under routine conditions, parallel line double reciprocal plots are obtained (33). However, at low pH (33) or at low temperature (38) converging line patterns result. A third pattern is observed, a t least with phenylalanine, when the range of 0, concentration is extended to high values through the use of high 0, pressures. I n this case, the lines are again parallel and straight with S as variable substrate, but distinctly curved (or rather biphasic) and parallel with 0, as variable substrate (33) Stopped-flow R H R measurements pointed to several important conclusions (38). First, the transient long wavelength absorbing species occurring chronologically between E, and E,, with the majority of substrates a t p H 7.8 and low temperature, was formed a t a rate exceeding the maximum turnover number but decayed a t a rate which was considerably smaller than the maximum turnover number. Hence, the transient 107. D. Wellner and A. Meister, JEC 236, 2357 (1961). 108. V. Massey and B. Curti, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 226. Univ. Park Press, Baltimore, Maryland, 1968.

458

HAROLD J . BRIGHT AND DAVID J. T . PORTER

intermediate, but not E,, qualified as an obligatory intermediate in turnover. Second, the electronic spectrum of the transient intermediate differed significantly, depending on the substrate used. The intermediate must therefore contain some element of the substrate and cannot be, for example, E-(FADB)z (which, in any case, has a totally different spectrum). Third, the decay of the intermediate was entirely independent of the substrate concentration. This makes it extremely unlikely that the conversion of E, * S to the intermediate requires the participation of a second substrate molecule. Massey and Curti postulated that the long -S.

-

wavelength intermediate is a biradical of the form E

which

*FADH. subsequently converted t o E, through E, * P1.Because the latter structure requires no more postulated intermediates than observable kinetic processes, we shall take E, * * * P1to be a suitable description of the long wavelength intermediate (but see Section V,A,2,b). Massey and Curti, through OHR stopped-flow measurements, showed that the oxidation of E, by 0, was too rapid to support the argument that inhibition of turnover a t high S and low 0, was caused by the accumulation of E, under these conditions (38).What appears to be the correct interpretation of the substrate inhibition phenomenon was first suggested by DeSa and Gibson, who noted that in SSK(E) experiments a t high concentrations of S much of the enzyme appeared spectrally to be fully reduced (109). They pointed out that this species (which cannot be E,) is probably E, S and that relatively slow oxidation of E, * * S by 0, would explain the observed inhibition. This explanation implies, of course, that the rate of dissociation of E, * * PI to form E, is an obligatory process when the enzyme is inhibited but is not substantially involved when the enzyme is turning over maximally. It will be noted that this explanation of the inhibition process is similar in principle to that used by Wellner and Meister (lor),but differs, based on the evidence from SSK(E) experiments (109), in the assignment of the enzyme species which is reoxidized slowly. On the basis of their studies, Massey and Curti (38)proposed a mechanism very similar to that already suggested for D-amino acid oxidase (36,466)[Eq. (20)]. However, only steps 1, 2, 3, and 6 were directly measured in rapid reaction experiments, and none of these was correlated with steady-state parameters. Furthermore, ternary complexes of the type 109. R. J. DeSa and Q. H. Gibson, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 25, 649

- - -

9

(1966).

7. FLAVOPROTEIN

O2 -

459

OXIDASES

- -

* * E, * P, were postulated to explain convergence in the steady-state double reciprocal patterns. As we have noted with D-amino acid oxidase (Section IV,A) reversibility of the reductive half-reaction (caused by values of k-, and k-, which are similar to those of k , [ S ] and k,, respectively) is also a sufficient condition for convergence. Porter undertook a systematic kinetic study of L-amino acid oxidase, using the full complement of techniques outlined in Section 111, with phenylalanine as substrate (33). This substrate was chosen for several reasons. First, the a-imino and a-keto products can be measured spectrophotometrically either directly or through their tautomers. Second, the decay rate ( k 3 ) of E, P, is easily measured in RHR experiments and can be made competitive with k5[02].Third, there is pronounced substrate inhibition in this case and the ultimate product (P,) (phenylpyruvate) is also inhibitory. Phenylalanine therefore serves to illustrate many interesting aspects of the reaction. The RHR and OHR results, when summed, established loop A of Eq. (20) and the steady-state expression of Eq. (44) :

-

1

-

Two of the rate constants, namely, k , and k,, are highly pH-dependent. Flavin reduction, controlled by k 2 , depends on a basic group with a pK, value greater than eight and has a limiting pH-independent value probably in excess of lo4 sec-'. The release of P, from E, * PI, controlled by k3, has a complex pH dependence in that it rises to a plateau value of 10 sec-l as the pH is raised (pK, 6.3) and then increases indefinitely beyond pH 9. Consequently, k , is the major rate-limiting first-order process in loop A of Eq. (20) in the accessible pH range from 5.5 to 10. Comparison of Eq. (44) with the steady-state rate equation from SSK (0,) measurements gave a quantitative term-by-term correlation only a t 0, concentrations less than lo-' M . At 0, concentrations of 5 mM and greater, the steady-state rate equation was the following [Eq. (45) 1 :

-

e

where 4"' and &' were much smaller and greater, respectively, than the first and third terms of Eq. (44). These experiments are most simply interpreted as reflecting the direct oxidation of E, * * P, [loop B, Eq. (2011 under conditions where k , [ O , ] >> k,, with +;-l = ks and +;-I = k,. It was proved by stopped-flow experiments that E, * P, (or an isomer thereof) must indeed react directly with 0,. The question of inhibition at high concentrations of S was answered

-

460

HAROLD J. BRIGHT A N D DAVID J. T. PORTER

quantitatively as follows. First, the conclusion that there accumulates a species of enzyme having a spectrum characteristic of reduced enzyme (but which is not E,) was confirmed. It was also noted that inhibition is most severe when k , is large (i.e., a t high pH) and when the ratio S/O, is large. It therefore followed that E, . S, identical spectrally to E,, was probably accumulating under inhibitory conditions as first suggested by DeSa and Gibson (109). With the scheme of Eq. (20) and its evaluated rate constants, it could be calculated that, if this were so, and if the bimolecular rate constant (k,) for oxidation of E, * S was appreciably smaller than k4,then the last 10-5 M 0, in a turnover experiment should be consumed in a first-order manner, with the value of k O b s / [ E ~extrapolating ] to k4 a t zero S and to k, a t very high S. Such experiments indeed exhibited this behavior and showed that k8 was about five times smaller than k, and about four times larger than k,. The dissociation constant for E, * * * S was about 20 mM. Phenylpyru* Pzbeing vate behaves similarly, the dissociation constant for E, about 50 mM, while this compound was oxidized about 10 times slower than E,. The reactivity of E, * * * PI can be summarized as follows [Eq. (4611:

-

- -

-

E,

* * *

P,

ks

E, X k-3

k4[0a1

= E,

It

+

6

H,O,

(46)

where k, > k6, k,, k,. Thus all ligands able to bind at the substrate site appreciably inhibit the rate of flavin oxidation. Precisely the opposite appears to be the case in the D-amino acid oxidase reaction, where k6 > k4 (44). With that enzyme, therefore, substrates utilizing loop A of Eq. (20) (possibly basic amino acids or neutral amino acids a t high pH values, where k , is large compared to k , [ O , ] ) might be anticipated to be inhibited by high 0, concentrations and to give curved, but parallel, double reciprocal steady-state patterns with 0, as variable. Rapid reaction studies with a-[ 2H]phenylalanine established that the a-C-,H substrate bond is cleaved in the rate-determining step in flavin reduction (kz) (46). Subsequently, i t was shown by Page and VanEtten (110-112) that in the case of leucine the substrate kinetic isotope effect is strikingly pH-dependent, having its maximum value a t low p H and tending to unity a t pH 9. They ascribed this to a change in rate-determin110. D. S. Page and R. L. VanEtten, BBA 191,380 (1969). 111. D. S. Page and R. L. Vadtten, BBA 191, 190 (1969). 112. D. S. Page and R. L. VanEtten, BBA 227, 10 (1971).

7. FLAVOPROTEXN

461

OXIDASES

ing step such that C-H boild cleavage is no longer rate-determining a t basic pH values and presented a reaction mechanism to account for the data. A similar pH-dependent substrate kinetic isotope effect was found in the case of phenylalanine (33) and an explanation was given based on Eq. (15). Evidence that the first product (PI) released by the enzyme is the a-imino acid was obtained using cyanide to trap this species ( 5 3 ) .These studies also explained why the scheme of Eq. (20) (with k-, = 0) was incapable of simulating the turnover behavior of E, * PIa t 550 nm, Under the conditions of stopped-flow turnover experiments, with high enzyme concentrations, the imino acid accumulates transiently a t levels approaching the dissociation constant k,/k-,. The process k-, [PI] is only effectively zero in the presence of high cyanide concentrations, which prevent the buildup of a-imino acid. Borohydride trapping experiments had also detected a-imino acid in this reaction as well as in the D-amino acid oxidase reaction (95). Finally, substituent effects with aromatic amino acid substrates have been examined (115,114).In neither case was the substituent effect traced to individual steps in the mechanism of Eq. (20). The formidable difficulties in the interpretation of such experiments have been discussed (see Section V1,A).

-

C. GLUCOSE OXIDASE Most kinetic studies of glucose oxidase have been carried out with enzyme from Aspergillus niger and from species of Penicillium. The A . niger enzyme is a dimer, of molecular weight 186,000, having two very tightly bound FAD molecules per dimer (115),while that from P . amagasakiense is very similar, the dimer having a molecular weight of 160,000 (116'). I n the latter case, each unit in the dimer is composed of two polypeptide chains connected by a disulfide bond. Swoboda carried out an extensive series of studies on the binding of FAD to the apoenzyme from A . niger (117).The latter was prepared by a modification of the classic acid ammonium sulfate resolution procedure of Warburg and Christian (118). Binding of FAD was shown to be followed by at least one, and probably two, unimolecular steps associated with protein conformational changes and it was proposed, on the basis of hydrodynamic and other measurements, that coenzyme binding 113. G. K. Radda, Nature (London) 203, 936 (1964). 114. D. S. Page and R. L. VanEtten, Bioorg. Chem. 1, 361 (1971). 115. B. E. P. Swoboda and V. Massey, JBC 240, 2209 (1965). 116. T. Yoshimura and T. Isemura, J. Biochem. (Tokyo) 69, 839 (1971). 117. B. E. P. Swoboda, BBA 175, 365 and 380 (1969). 118. 0. Warburg and W. Christian, Biochem. Z . 298, 368 (1938).

462

HAROLD J. BRIGHT A N D DAVID J . T. PORTER

converted the loose flexible coil configuration of the apoenzyme to a compact and almost spherical holoenzyme. Evidence for the sites of interaction on the FAD molecule was obtained from studies of the binding of a series of related nucleotides. The effectiveness of binding was such that it was proposed that the adenine and phosphate moieties of FAD are the first to bind, followed by the isoalloxazine nucleus. The fundamental aspects of the kinetic mechanism were incisively established for the first time by Gibson et al. (,??6) and by Nakamura and Ogura ( 4 7 ) . These studies employed all of the approaches outlined in Section 111, and the resulting mechanism has therefore required little adjustment in the intervening period and has provided a solid basis for further experimentation. The work of Gibson et al., carried out a t pH 5.5 over a range of temperatures with the A . niger enzyme, is a useful starting point for discussion (36). Three classes of sugar substrates were identified on the basis of SSK(0,) experiments. Glucose, by far the most reactive (Oo-I = 1150 sec-' at 2 7 O ) , gave the three-term steady-state rate equation [Eq. ( 5 ) ] while the equation for 2-deoxyglucose lacked +2 and those for mannose, xylose, and galactose lacked both +o and +*. SSK(E) experiments with glucose established that the steady-state parameters were independent of enzyme concentration. RHR stopped-flow measurements a t 450 nm immediately etablished the turnover mechanism as Eq. (19) for 2-deoxyglucose [with +o-l = k, and 41-1 = k , k 2 / ( k - , k,)] and as Eq. (16) = k,.). Neither of these substrate types, for the mannose group (with in contrast to glucose, is therefore sufficiently reactive in the R H R to cause the OHR to become rate limiting under the conditions used. The high reactivity of glucose causes several additional steps to become kinetically significant. First, RHR and OHR measurements yielded the identities +,-' = k, and +2-1 = k, [see Eqs. (15) and (18)]. SSK(E) experiments at Oo established that a terminal first-order process in each of the half-reactions was responsible for the maximum turnover number [&-l = k,k,/(k, k,) , see Eq. (23) 1. The general mechanism necessary to describe the behavior of all sugars under all conditions tested was, therefore, loop A of Eq. (20). In the case of glucose, k , [ S ] , k,, k 4 [ 0 2 ] , and k, are the only kinetically significant processes at 0" and p H 5.5. Formally, k, and k, were only shown to govern first-order conversions of enzyme species having the spectra of reduced and oxidized flavin, respectively. However, in view of the failure of substantial efforts to relate them to enzyme conformational changes through spectrofluorometric and spectrophotometric transients unassociated with flavin redox processes, we have taken loop A of Eq. (20) as a most probable, if not proved, hypothesis. Thus, glucose oxidase appears to be a rare example of a flavo-

+

+

7. FLAVOPROTEIN

463

OXIDASES

protein oxidase which truly oscillates with all substrates and under all conditions between fully oxidized and free fully reduced states during turnover and thus deserves the ping-pong appellation. The activation energy for k , is much higher than that for k,, and it ceases to be a significant contribution to cpo at temperatures about 1 3 O . Nakamura and Ogura, using enzyme from both P. amagasakiense (47) and A . niger (48), derived a kinetic mechanism for glucose oxidation which would satisfy most of the data of Gibson et al. (25) obtained a t 20" or above in the sense that a first-order process involving oxidized enzyme was the only contributor to 90. However, they claimed that this was flavin reduction ( k 2 ) , whereas Gibson et al. (25) showed, through SSK(E) experiments, that this process was the release of HzOz from E, . * H20z controlled by k5 (or a kinetically equivalent first-order step such as a conformational change). However, the discrepancy rests heavily on the interpretation of R H R data obtained at high glucose concentrations when the observed half-times for flavin reduction approach the mixing time of the stopped-flow apparatus. Inspection of the published data (48, Fig. 5 ) shows that the extrapolated value of kG: a t infinite glucose is probably indistinguishable from zero and therefore consistent with the results of Gibson et al. (26). Studies with [ l-2H]glucose ( 5 1 ) ,originally conceived to test whether, in fact, substrates such as glucose actually form the E, * S complex (since k , cannot be detected in these cases), gave several interesting results. First, with this substrate, the R H R clearly reached a saturating velocity (k, = 67 sec-l a t 3"). This showed that E, * * S must be formed from glucose and that k , with [l-lH]glucose must have a value not much greater than lo3 sec-' in order to be compatible with the larger experimental kinetic isotope effects reported for C-H cleavage (119). A value of lo3 sec-l for k , in the case of [l-lH]glucose a t 3 O would be consistent both with the original RHR stopped-flow experiments (25) (in which l/kz was indistinguishable from zero under conditions where half-times less than about 5 msec would not be measurable for technical reasons) and with the conclusion that +,,-' ( = 330 6ec-l at 3 O with [ l-lH] glucose) is almost entirely regulated by k, and k,, with k2 having a 20% or less contribution. Judging from the temperature dependence of k, for 2-deoxyglucose and of +"-l for [l-ZH]glucose,the contribution of k , (-4 X lo3 sec-*) to (-lo3 sec-1) a t 25O and pH 5.5 with [l-lH]glucose would remain a t about 2076, with k , being predominant and k, exerting no control because of its relatively high activation energy. With [ 1-'H] glucose a t 25", (184 sec-') is probably determined entirely by k,, as is the case at low temperature. Second, these studies were the first to

-

-

119. J. Hampton, A. Leo, and F. H. Westheimer, JACS 78, 306 (1956).

464

HAROLD J . BRIGHT A N D DAVID J. T. PORTER

demonstrate in a flavoprotein oxidase that substrate C-H bond cleavage, at least in the case of the deuterated substrate, is rate determining in flavin reduction per se, inasmuch as k , is directly measured as the spectrophotometric change which is associated with the flavin redox process. Third, the kinetic isotope effect is so large that the steady-state equation changes from Eq. (47) for [l-lH]glucose to Eq. (48) for [l-2H]glucose:

This transition serves to illustrate that, with a highly reactive h e . , “physiological”) substrate such as glucose, the numerous first-order processes in the catalytic mechanism are likely to be so closely matched energetically, perhaps as an evolutionary consequence (51),that a relatively minor perturbation in one step may easily change the distribution of rate-determining processes among the several steps in the mechanism. Poor substrates (e.g., mannose), as a corollary, tend to have one transition state whose energy is far in excess of any other. I n practical terms, Eqs. (47) and (48) emphasize that the usual assumption in kinetic isotope effect studies, namely, that introduction of the heavy isotope merely slows a step which is already rate determining with the substrate of natural isotopic abundance, needs to be carefully evaluated by rapid reaction techniques where the isotopes of hydrogen are concerned. Keilin and Hartree obtained a handsome bell-shaped initial velocitypH profile centered a t about pH 5.5 in their pioneering studies of glucose oxidase (1.20). Most research investigations and applications of the enzyme have utilized this pH value since that time. Although the marked pH dependence shows that one or more of the + coefficients must be highly pH-sensitive, neither 0, nor glucose was saturating in the original studies, and hence the origin of the pH dependence was unclear. Bright and Appleby (26)addressed this question by systematically studying the pH dependence of the individual steps of the P. notatum enzyme according to the mechanism of Gibson et al. deduced a t 25O and pH 5.5. Such studies are simplified by the unusually high stability of the enzyme over the pH range from 3 to 10. With glucose, k , ( = and k, ( = &!-l) were sigmoid functions of pH, the former being governed by a basic group (pK,, = 5.0 a t 25O and 0.2 M KCl), the latter by a n acidic group (pK,, app = 6.9 under the same conditions). The combination of these two steps accounts entirely for the overall pH dependence a t nonsaturating S and 0,. In the case of the only substrate for which saturation is 120. D. Keilin and E. F. Hartree, BJ 42,221 (1948).

7.

465

FLAVOPROTEIN OXIDASES

observed in the RHR, namely, 2-deoxyglucose, the p H dependence is observed in k-, k,/k, as expected, while k, is pH-independent. The terminal step in the OHR, governed by k , and possibly representing Eo * H,O, breakdown, was originally thought to have a bell-shaped pH dependence. This was later shown to be in error ( 2 7 ) ,in that k, depends only on an acidic group with pK,, 9. Although the latter pH studies (27) confirmed the major features of the earlier work (25,26),some new aspects of the mechanism were discovered. First, it had been noted that halides (X-) specifically bind to H'E, and hence act to raise the value of pK,, app.

+

-

-

H+E,

..

*

ka s+ E,

+ P,

(49)

Halides are therefore potent inhibitors a t low pH values, the order of effectiveness being F- >> C1- Br-. However, halides (particularly F-) were also shown, less effectively, to decrease k , to such an extent that the importance of k , in glucose turnover could be assessed. It was concluded that in the presence of halides k , was rate limiting at high S and 0, at low pH values but that k , was rate limiting a t high pH values in the presence or absence of halides. The mechanism of Gibson et al. (25) is therefore an accurate description of the rate-determining process in glucose oxidation a t pH 5.5 in the absence of halides, but to account for the reaction a t all pH values in the presence of halides, the required mechanism is a hybrid of those proposed by Gibson et al. (25) and by Nakamura and Ogura ( 4 7 # ? ) .Second, the oxidation of reduced enzyme by 0, is not, as deduced earlier ( 2 6 ) , a single process governed by an acidic group in the enzyme or in flavin itself. Rather, there is a rapid pathway governed by an acidic group of pK,, app = 7.5 (k, = 2 X lo6 M-' set+) and a slower pathway which is predominant a t pH values greater than 9 (k4' = 1.5 x lo5 M-l sec-I). The ionization involved in oxidation of E, may represent protonation a t N1 of the flavin nucleus. Rogers and Brandt carried out a series of kinetic and spectrophotometric binding measurements of the interaction of halides and of D-glucal with glucose oxidase (121).They confirmed that C1- reacts preferentially M and that glucose reacts with H'E, [Eq. (49)] with K, = 5 X obligatorily with the conjugate base E,. As a consequence, halides are simple competitive inhibitors with respect to substrates. D-Glucal is interesting in two respects. First, having a planar structure at C-1 somewhat analogous to that wihch substrates must assume as they are converted to the product lactone, it might be suspected to be a transition state analog. Despite the fact that D-glucal proved to be one of the very few 121. M. J. Rogers and K.G.Brandt, Biochemkstry 25, 4624, 4630, and 4636 (1971).

466

HAROLD J. BRIGHT AND DAVID J. T. PORTER

substrate-competitive inhibitors of glucose oxidase, its poor affinity, compared to the substrate 2-deoxyglucose, for example, indicated that it is probably not a close transition state analog. Second, D-glucal binding was found to be independent of both H+ and halide concentrations. This was shown to result from simultaneous occupancy of the active site by both D-glucal and a halide ion (each of which, separately, is substrate-competitive). This behavior is consistent with the idea proposed previously, namely, that the basic group in E, (and which regulated k, through pR,) interacts with the 1-hydroxyl group of sugar substrates or (after protonation) with a halide ion. When the 1-hydroxyl group is absent as in D-glucal, binding becomes pH-independent and, moreover, further modification of the basic group (such as halide binding following protonation) by small ligands is expected to have little effect on D-glucal binding. Keilin and Hartree concluded originally, as had others before them, that the product of the overall reaction was gluconate (120).Bentley and Neuberger subsequently showed that the first product released by the enzyme was D-glucono-S-lactone and that this was subsequently hydrolyzed nonenzymatically (122). The mechanism of hydrolysis of the lactone has been studied (123).

D. MONOAMINE OXIDASE There are two classes of monoamine oxidase, namely, the pyridoxal phosphate- and copper-containing enzymes such as plasma monoamine oxidase, spermidine oxidase, benzylamine oxidase and lysyl oxidase, and the FAD-containing monoamine oxidases. The latter will be the sole concern of this discussion. The properties of these enzymes have been reviewed by Blaschko (124). The physiological role of monoamine oxidase in the biological inactivation of catecholamine a t nerve endings is an accepted fact (125).The importance of the enzyme as a protective agent through enzymic deamination of biogenic monoamines is evident in its oxidation of noradreanaline. Thus, ingestion of cheese, which contains high amounts of tyramine, causes release of noradrenaline from storage granules. If the individual has been given a monoamine oxidase inhibitor prior to eating the cheese, death may result since the released noradrenaline is not destroyed by monoamine oxidase (12s). In addition to its protective role through en122. R. Bentley and A. Neuberger, BJ 45, 584 (1949). 123. M. A. Jermyn, BBA 37, 78 (1960). 124. R. Blaschko, “The Enzymes,” 2nd ed., Vol. 8, p. 377, 1963. 125. P. L. McGeer, Amer. Sci. 57, 221 (1971). 126. A. M. Asatoor, A. J. Levi, and M. D. Milne, Lancet 2, 733 (1963).

7.

FLAVOPROTEIN OXIDASES

467

zymic deamination of biogenic monoamines, it has been suggested that the oxidation product of the monoamine oxidase reaction may, in some cases, be a more physiologically active compound than the parent compound (127). Monoamine oxidase is located in the outer membrane of the mitochondrion (128).The major problem with monoamine oxidase purification has been that of solubilization of the membrane-bound enzyme. Although each laboratory tends to have its own preferred method of solubilization, the general pattern of purification is the preparation of mitochondria followed by solubilization, fractionation, and electrophoresis. Erwin and Hellerman (129) have purified monoamine oxidase from bovine kidney mitochondria. In this work, solubilization of the enzyme was accomplished with digitonin A and fractionation by calcium phosphate gel chromatography. Recently, Chuang (130)has modified the above procedure with a DEAE-chromatography step. The visible spectrum of the enzyme shows 450 nm absorbance, but other peaks are not well resolved. The absorbing contaminant wgs suggested to be a cytochromelike material. The spectrum of this enzyme is reminiscent of that of D-amino acid oxidase in early stages of purification. By dithionite titration, it was found to contain 115,000 g protein per mole flavin. Gomes et al. (131) and Yasunobo (132),on the other hand, have solubilized the enzyme with Triton X-100 and fractionated the enzyme with calcium phosphate, DEAE-cellulose, and hydroxylapatite column chromatography, followed by starch zone electrophoresis. The resulting enzyme preparation could be separated into three monoamine oxidases. CQmponents 1 and 2 had a molecular weight of 424,000 while component 3 had a molecular weight of 1,250,000. It was argued that the three components were not isozymes but, rather, represented states of different degrees of polymerization. Youdin and Sourkes (133) have purified monoamine oxidase from rat liver mitochondria. Instead of using a detergent, these authors have s o h bilized by sonification and have claimed that this method leads to higher-yields of soluble monoamine oxidase. The molecular weight was determined to be 290,000 by gel filtration and possibly 150,000 by centri127. V. Z.Gorkin, Pharmacol. Rev. 18, 115 (1966). 128. J. W. Greenawalt, Fed. Proc., Fed. Amer. SOC.E x p . Biol. 28, 663 (1969). 129. V. G.Erwin and L.Hellerman, JBC 242,4230 (1967). 130. H.Y. K.Chuang, D. R. Patek, and L.Hellerman, JBC 249,2381 (1974). 131. B. Gomes, I. Igaue, H. C. Kloepfer, and K. T. Yasunobu, ABB 132, 16 (1969). 132. K. T. Yasunobu, I. Igaue, and B,, Gomes, Advnn. Pharmacol. 6, Part A, 43 (1968). 133. M. B. H.Youdin and T. L. Sourkes, Can. J. Biochem. 44, 1397 (1966).

468

HAROLD J. BRIGHT AND DAVID J. T. PORTER

fugation. Tipton (154) solubilized and purified the enzyme from pig brain, replacing some of the chromatography steps of previous workers by ethanol and pH fractionation procedures. The molecular weight of the monomer was determined to be 105,000 by gel filtration, and a larger species was found with a molecular weight of 435,000. Hollunger and Oreland (135) and Oreland (136)have purified monoamine oxidase from pig liver mitochondria. The enzyme was solubilized in this case by a methyl ethyl ketone extraction procedure. The enzyme was purified by chromatographic steps on G-200 Sephadex followed by an acid precipitation and sucrose density gradient centrifugation. By gel filtration, the molecular weights of the two forms of the enzyme were 108,000-1 17,000 and 275,000-290,000. The K, for benzylamine of the pig liver enzyme purified by methyl ethyl ketone extraction was considerably lower than that for monoamine oxidase in mitochondria or monoamine oxidase solubilized by Triton X-100. This result may be explained by differences in phospholipid extracted by the two extraction methods or by preferential extraction of different isozymes by the two methods, The only metal of significance found in this preparation was iron. One mole of iron was found per 213,000 g protein, compared with one mole flavin per 115,000 g protein. The insensitivity of the enzyme activity to 1,lO-phenanthroline implies that iron does not play a part in the catalytic function of the enzyme. The importance of metals for the catalytic activity of monoamine oxidase has caused much debate, and most attention has been paid to copper. Originally, Nara et al. (137) proposed that bovine liver mitochondrial monoamine oxidase was a copper enzyme, but Erwin and Hellerman (129) concluded that copper was nonessential for the bovine kidney monoamine oxidase activity. Sourkes (138) summarized the data from several laboratories and concluded that i t is unlikely that copper is a prosthetic group of monoamine oxidase. However, all preparations of monoamine oxidase to date contain varying degrees of hemin as evidence by the absorbance seen in the Soret region. The recent modification of the Erwin and Hellerman (129) purification procedure by Chuang et al. (159)removed most of the Soret absorbing material. It is now generally agreed that metals are not cofactors for mitochondrial monoamine oxidaee catalysis. 134. K. F. Tipton, Eur. J. Biochem. 4, 103 (1968). 135. G.'Hollunger and L, Oreland, ABB 139, 320 (1970). 136. L. Oreland, ABB 148, 410 (1971). 137. S. Nara, B. Gomes, and K. T. Yasunobu, JBC 241, 2774 (1966). 138. T.L.Sourkes, Advan. Phannacol. 6A, 61 (1968). 139. H. Y . K. Chuang, D. R. Patek, and L. Hellerman, JBC 249, 2381 (1974).

7.

FLAVOPROTEIN OXIDASES

469

Purification of monoamine oxidase from mitochondria invariably results in different forms of the enzyme, raising the question of whether isozymes or artifacts of preparation are responsible for this multiplicity. Thus, the enzyme may have detergent bound to it (129),the amount of membrane material bound to a single enzyme species may vary (140), or there may be irreversible damage to the enzyme during solubilization. For a summary of these possibilities, the reader is referred to Collins (1411 * Recently, Houslay and Tipton (I@) have evaluated monoamino oxidase activity in rat liver mitochondria1 outer membranes that were prepared without the use of detergents. They concluded that there were two kinetically different monoamine oxidases by their sensitivities to clorgyline and the reversible inhibitors benzyl cyanide and 4-cyanophenol. However, they were not able to distinguish between the possibility that there was a single enzyme species in two different environments or whether the apoenzymes of the different monoamine oxidase activities were different. As with many other flavoprotein oxidases, double reciprocal plots a t fixed 0, and variable reducing substrate yield parallel lines. This result has been obtained for pig brain (149) and beef liver (144,1455)enzyme. Using benxylamine as reducing substrate, Oi et al. (I44145) have carried out a detailed study of the kinetic mechanism of monoamine oxidase by studying product inhibitor patterns. One surprising conclusion from this study was that the aldehyde and ammonia products were released a t different steps and not together, as the imino compound, which is then hydrolyzed nonenzymically to the aldehyde and ammonia. This interpretation is at variance with the results of Patek et al. (146’) who showed that the primary product released from the enzyme is the imino compound. With benzylamine as substrate, it is known that monoamine oxidase undergoes an anaerobic half-reaction as well as aerobic turnover. In the anaerobic half-reaction, the aldehyde and ammonia (presumably as the imino compound) and reduced enzyme are formed while H,O,, imino compound, and oxidized enzyme are made during aerobic turnover. From 140. K. F. Tipton, M. D. Houslay, and N. J. Garrett, Nature (London) 246, 213 (1973). 141. G. G. S. Collins, Advan. Biochem. P,sychopharmacol. 5, 129 (1972). 142. M. D. Houslay and K. F. Tipton, Biochem. J. 139,645 (1974). 143. K. F. Tipton, Eur. J. Biochem. 5, 316 (1968). 144. S. Oi, K. Shimada, M. Inamasu, and K. T. Yasunobu, ABB 139, 28 (1970). 145. S. Oi, K. T. Yasunobu, and J. Westley, ABB 145, 557 (1971). 146. D. R. Patek, H. Y. K. Chuang, and L. Hellerman, Fed. Proc., Fed. Amer. SOC.Ezp. B i d . 31, 420 (1972).

470

HAROLD J. BRIGHT AND DAVID J. T. PORTER

these results, together with the fact that double reciprocal plots of turnover data yield parallel line patterns, it has been postulated that the enzyme cycles between its oxidized and free reduced states (144). The inadequacy of this type of reasoning has been demonstrated in the case of D-amino acid oxidase (see Section IV,A) . A detailed study of the halfreactions is imperative in order to determine the importance of free fully reduced enzyme during catalysis. Oi et al. (145) have studied the pH dependence of the steady-state parameters. The pH dependence of &-* is similar to that found for other flavoprotein oxidases in that it increases with increasing pH and corresponds to an apparent pK value of 7.3. Hellerman et aZ. (147) have carried out Hammett a-p measurements with substituted benaylamines and obtained a biphasic plot of log VP,max vs. U. This result should be contrasted with those of Zeller et al. (148,149). The interpretation of such measurements, as discussed for the D-amino acid oxidase reaction (Section IV,A) , is extremely difficult. The flavin prosthetic group (FAD) is covalently bound to the apoenzyme in mitochondrial monoamine oxidase isolated from pig liver (1361, beef liver (131), and beef kidney (129). The pentapeptide segment to which the flavin is attached has been identified by Singer's group (150,151).A thioester linkage occurs between cysteine and the 8a-carbon of the flavin nucleus. Two classes of monoamine inhibitors have become increasingly important in their potential for treatment of hypertension and nervous system depression. These are hydrazino compounds and acetylenic compounds such as pargyline. In the present context, these two compounds will be discussed in relation to the mechanistic implication for monoamine oxidase action. Pargyline is known to react stoichiometrically and irreversibly with mitochondrial monoamine oxidase (152). Recently, Chuang et aZ. (139) and Oreland et al. (153), with the bovine and pig liver enzymes, respec147. L. Hellerman, H. Y. K. Chuang, and D. C. DeLuca, Advan. Biochem. Psychopharmacol. 5, 327 (1972). 148. E. A. Zeller, B. H. Babu, and W. F. March, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 29, 943 (1970). 149. E. A. Zeller, M. Hsu, and J. T. Ohesson, Fed. Proc., Fed. Amer. Soc. Exp. Biol. 32, 544 (1973). 150. E. B. Kearney, J. I. Salach, W. H. Walker, R. L. Seng, W. Kenney, E. Zeszotek, and T. P. Singer, Eur. J . Biochem. 24, 321 (1971). 151. W. H. Walker, E. B. Kearney, R. L. Seng, and T. P. Singer, Eur. J . Biochem. 24, 328 (1971). 152. L. Hellerman and V. G. Erwin, JBC 243,5234 (1968). 153. L. Oreland, H. Kinemuchi, and B. Y. Yoo, Life Sci. 13, 1533 (1973).

7.

FLAVOPROTEIN OXIDASES

47 1

tively, have demonstrated that the pargyline-inactivated enzyme contains a covalent adduct formed from the inhibitor and the FAD moiety. Thus, using pargyline labeled in the benzyl portion, it was found that the label was associated with the flavin-peptide conjugate removed by hydrolysis from the enzyme. The covalently modified flavin peptide had strong absorbance a t 398 nm (139). In addition, the pargyline-monoamine oxidase adduct lost the benzyl group upon incubation in base (165). These results are consistent with the proposed structure of the photochemically generated product of flavoquinone and pargyline reported by Zeller et al. (154). The substituted hydrazines comprise a second class of monoamine oxidase inhibitors. The mechanism of inactivation of the bovine kidney enzyme has recently been shown to occur by formation of a stable flavininhibitor adduct (166).Patek and Hellerman (165) concluded that 0, was essential for inactivation of the enzyme and that inactivation was caused by an interaction of the oxidation product of the substituted hydrazine found to be active as an enzyme inhibitor. The spectrum of the isolated flavin-peptide from inactivated monoamine oxidase was stable to 0, and had the character of reduced flavin. Inactivation may result from alkylation of oxidized flavin by the carbanion formed from breakdown of the diazene (156).Identification of this sustituted flavin would probably aid in determining the reactive center of the flavin for this enzyme. Tipton (167), studying the enzyme from pig brain, found that the irreversible inhibition by phenylhydrazine was not affected by the absence of oxygen, in contrast to the results of Patek et al. (166). I n addition, the inhibition of pig monoamine oxidase by phenethylhydrazine was attributed to the oxidation of product phenethylhydrazine and not to phenethyldiazene as proposed by Patek et al. (156). Whether these differences should be attributed to differences in sources or to other factors is not yet clear.

E. OLD YELLOWENZYME Old Yellow Enzyme catalyzes the oxidation of NADPH and NADH by 0, and other electron acceptors. Nakamura et al. (168) have studied the kinetics of oxidation of NADH and NADPH by 0, at p H 7.0.Upon 154. E. Zeller, B. Gartner, and P. Hemmerich, 2. Naturjorsch. B 27, 1050 (1972). 155. D. R. Patek and L. Hellerman, JBC 249, 2373 (1974). 156. T. Tsuji and E. M. Kosower, JACS 93, 1992 (1971). 157. K. F. Tipton, BJ 128, 913 (1972). 158. T. Nakamura, J. Yashimura, and Y . Ogura, J. Biochem. (Tokyo) 57, 554 (1965).

472

HAROLD J. BRIGHT A N D DAVID J. T. PORTER

anaerobic addition of either NADH or NADPH to the enzyme, a transient intermediate absorbing a t long wavelengths was observed. The kinetics of formation of this species were not measured but its rate of decay was found to be 0.046 sec-' and 0.5 sec-', with NADH and NADPH, respectively. The intermediate was taken to be the same as the complex formed upon the addition of reduced enzyme to either NAD or NADP. With O2 as an electron acceptor and NADH as the reductant, the accumulating enzyme species corresponded again to the complex between reduced enzyme and NAD. With NADPH as the reductant, the accumulating species did not correspond to the complex between reduced enzyme and NADP, indicating that another species, such as free reduced enzyme, accumulated during turnover. Since the turnover number for the oxidation of NADH was significantly greater than the rate of decay of the complex between NAD and reduced enzyme, it was proposed that the reduced enzyme-NAD complex reacts directly with 0,. In the case of NADPH, evidence for reactivity of the intermediate with 0, was less convincing since the turnover number of the enzyme was less than the first-order rate constant for the breakdown of the intermediate. The transient intermediate was recognized to be different from the anionic semiquinone even though there was an ESR signal associated with the complex between reduced enzyme and NADPH. Subsequently Massey et al. (159) found that the anaerobic addition of NADPH a t pH 8.5 to Old Yellow Enzyme results in three kinetic M ) was transients a t 530 nm. The first absorbance burst ( K D= 9 X unresolved in the stopped-flow apparatus and was followed by a further absorbance increase corresponding to 250 sec-l. The third phase was associated with an absorbance decrease having a rate of 1.3 sec-l. The stopped-flow spectrum of the intermediate a t the end of the second phase resembled that of oxidized enzyme having long wavelength absorbance. At the end of the third phase, the enzyme is fully reduced. The following scheme [ Eq. 501 was proposed : E-FMN -Er

FMN

250 sec-1

,FMNH*

1.3 sec-1

___t

'NADPH

E*'*NADP*

(n)

(I)

"'NADP (111)

159. V. Massey, R. G. Matthews, G. P. Foust, L. G. Howell, C. H. Williams, Jr., G. Zanetti, and S. Ronchi, in "Pyridine Nucleotide-Linked Dehydrogenases" (H. Sund, ed.), p. 394. Bpringer-Verlag, Berlin and New York, 1970; JBC 244, 1779 (1969).

7.

FLAVOPROTEIN OXIDASES

473

Because of the inferred reactivity of intermediate I1 with ferricyanide (i.e., turnover number of enzyme with ferricyanide was larger than the first-order decay of I1 to 111),it was suggested that the flavin in I1 was reduced. However, addition of NADP to E-FMNH, yielded a spectrum which differed at long wavelengths from that of I1 and a biradical structure, rather than one in which the flavin is fully reduced, was suggested for 11. I n point of fact, the spectrum of I1 and of the species resulting from the addition of NADP to E-FMNH, has a good deal of oxidized flavin character. The assignment of chemical structure to intermediates I1 and I11 might be clarified by kinetic isotope effect studies with NADP2H. These could, for example, establish whether hydrogen transfer to the flavin has occurred prior to the reaction of I1 with ferricyanide. Porter et al. (160) have found, in a model system, a spectrum of a species that is similar to that reported for intermediate I1 in the reduction of Old Yellow Enzyme. The spectrum corresponded to that of a complex between oxidized lumiflavin and N-methyl dihydronicotinamide since it occurred prior to C-H bond cleavage in the latter as demonstrated by the locus of the deuterium kinetic isotope effect. Blankenhorn (161) has measured the intramolecular electron transfer in a bisnicotinamide-flavin model reaction. The primary complex between oxidized flavin and reduced nicotinamide exhibited long wavelength absorbance but showed no sign of reactivity with ferricyanide. Old Yellow Enzyme forms long wavelength absorbing complexes with small ligands (159,162) which have been suggested t o be charge-transfer complexes (159). Enzyme isolated by the method of Matthews and Massey (159) contains a regreening factor (RGF) which can be displaced by reduction of the enzyme followed by dialysis. Regreening factor was found to be a small molecule with a pK value of 7.5. Conclusive identification of this factor has not been made, but it has been suggested to have a heteroaromatic structure that is able to undergo further reduction (159). The K , for NADPH is the same for enzyme complexed with R G F as for that free of RGF. These results suggest that RGF is bound a t a site which is entirely different from that for NADPH and which may be the site for the unknown natural receptor (163).Exploitation of the tight binding of small molecules to Old Yellow Enzyme has resulted in a one-step affinity chromatqgraphic purification of the enzyme (162). 160. D. J. T. Porter, G. Blankenhorn, and L. L. Ingraham, BBRC 52, 447 (1973). 161. G. Blankenhorn, Eur. J . Biochem. 50,351 (1975). 162. A. S. Abramovitz and V. Massey, Fed. Proc., Fed. Amer. Sac. Exp. Biol. 33, 1569 (1974). 163. R. Matthews and V. Massey, in “Flavins and Flavoproteins” (H. Kamin, ed.), p. 329. Univ. Park Press, Baltimore, Maryland, 1971.

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HAROLD J . BRIGHT AND DAVID J. T. PORTER

V. The Chemical Mechanism of Flavoprotein Oxidases

A. THECHEMICAL MECHANISM OF FLAMN REDUCTION 1. Models for Flavin Reduction The chemical mechanism of flavin reduction by amines, alcohols (or hemiacetals) , and related substrates has often been discussed in terms of one discrete pathway to the exclusion of all others. However, it is now not clear that one such discrete pathway exists, or alternatively, if it does exist, whether current experimental methods are capable of detecting this pathway and unequivocally eliminating all others. For these reasons the following scheme [Eq. (51)I, which displays most of the feasible redox and a-p-elimination mechanisms that can be expressed by formal structures, may not be so much an exercise in paper chemistry as a serious statement of plausible mechanisms which must be systematically subjected to experimental test. It emphasizes, in particular, how easily a given redox pathway could be a mixture of homolytic and heterolytic processes, as well as the inadequacy of experimental criteria used up to now in detecting and differentiating between alternative pathways. The scheme for flavin reduction is bounded by the dashed line and a-p elimination by the dotted line. Intermediates (as opposed to products) common to both pathways appear in the overlap area.

.................................................................... 1

Table I classifies the pathways of Eq. (51) according to the mode of scission of the substrate C-H bond and indicates that for the vast majority of physiological substrates, C-H scission is the rate-determining step (measured experimentally as k,) between E, * * S and E, * PI.

-

-

7.

475

FLAVOPROTEIN OXIDASES

TABLE I CLASSIFICATION OF PATHWAYS SHOWNIN EQ. (51) Steps in the reaction

E o . . . S -kl+ E r .

*

PI

Rate determining

Rapid

1. Hydride abstraction

5

-

2. Proton abstraction a. Followed by one 2-electron transfer b. Followed by two l-electron transfers c. Followed by adduct formation and rearrangement d. Synchronous with adduct formation followed by rearrangement of adduct

8 8 8 4

C-H

scission

3. Hydrogen atom abstraction 8. Followed by electron transfer

or b. Followed by adduct formation or

11 11 11 11

14 12 and 6 3 and 10 10

6 12 and 14 7 and 10 12, 3, and 10

The resulting nine (and possibly more) pathways are experimentally indistinguishable on the basis of the evidence obtained thus far with physiological substrates. Studies with model substrates (analogs and inhibitors) and flavins, as well as evidence from nonenzymic model systems, tend to favor several of these pathways. 2. Evidence f r o m Enzyme Studies: Native Coenzyme and Physiological

8UbStTclteS ' P I and E , Formation from E, Sin a. The Kinetics of E , Flavoprotein Oxidases. The flavoprotein oxidase reductive half-reaction in-

-

- -

volves, a t most, four detectable intermediates [see Eqs. (19) and (20) ], namely, E,, E, * * S, E, * * P,, and E,. Since flavin reduction is initiated within E, * S and is either entirely completed a t the stage of E, * P, or a t least half-completed (see Section V,A,P,b), the search S+ for reaction intermediates is focused on the process E, E, * * * P,. Rapid reaction techniques have failed to detect any spectral intermedi* SandE, P,in the aminoacidoxidasereacates between E, tions. Since most of the plausible intermediates of Eq. (51) (such as a covalent adduct or a flavin-substrate biradical) are known or expected from model studies (see Section V,A,B,b) to possess distinctive absorption spec-

-

-

- -

-

- - -

- -

- -

476

HAROLD J . BRIGHT AND DAVID J . T. PORTER

tra, this negative evidence shows either that intermediates do not, in fact, exist [ e.g., a hydride transfer mechanism by step 5 of Eq. (51) ] or, more probably, that scission of the substrate C-H bond is entirely rate determining in E, * P, formation and that intermediates resulting therefrom are converted very rapidly to E, * P. We assume for this discussion that E, * PI is a complex between fully reduced enzyme and the a-imino acid product (see Section V1A,2,b). Although the search for spectral intermediates lying between E, * * - S and E, * * PI (or E,) is often difficult because of the inability to saturate E, in the RHR, results from the glucose oxidase reaction show rather clearly that the spectrum of E, * S is indistinguishable from that of E, (27,48). Furthermore, it is probably implicit in reports of the pointby-point steady-state spectrum of E, * PI in the D- and L-amino acid oxidase reactions; where the kinetics and amplitudes of spectrophotometric transients were measured over a wide wavelength range (36,38,44) that the kinetics of E, * . * P, formation were independent of wavelength. This conclusion was specifically stated in spectrophotometric and fluorescence-monitored turnover studies of glucose oxidase, indicating that no spectral intermediates other than species corresponding to fully oxidized and fully reduced enzymes were detected (26). We would assume that such stopped-flow measurements would detect a novel transient spectrophotometric species which comprised at least 10% of the total enzyme. An alternative statement of this conclusion is that in no case has the kinetics of E, PI appearance (e.g., a t 550 nm) been reported to differ from those of E, * * * S disappearance (e.g., at 450 nm). This would rule out intermediates between E, * * S and E, * * PI of unique spectral properties having lifetimes exceeding about 5 msec. The steadyS in the D-amino acid oxidase state spectrum attributed to E, * reaction (164) is probably that of E, . * PI, as noted by Yagi (44). Despite the failure to detect such intermediates in the case of physiological substrates by rapid reaction techniques, such negative evidence in no way argues against either their existence or their obligatory role in catalysis. This is well illustrated by the oxidation of nitroethane anion by D-amino acid oxidase, where the conversion of E, * * S to E, * * P, behaved as a single kinetic process by rapid reaction criteria and yet was subsequently proved to consist of the interconversions of a t least three substrate-flavin adduct species (166). Several independent lines of evidence point to the conclusion that isomerization of the initial E, S complex must occur before the redox mechanism is initiated. First, the kinetics of binding of the substrate

- - -

--

- -

- -

- -

--

-

-

-

164. H. Watari, A. Isomoto, H. Oda, and M. Kuroda, BBA 167, 184 (1968). 165. D. J. T. Porter, J. G. Voet, and H. J. Bright, JBC 248, 4400 (1973).

7.

477

FLAVOPROTEIN OXIDASES

analogs benzoate and anthranilate to D-amino acid oxidase clearly gives evidence for a two-step process (166). It is interesting that lactate, a poor substrate for this enzyme, gives an E, S complex whose spectrum is vibrationally resolved in the 450-nm band, as is the case with the E,-benzoate complex (44). This is thought to arise from changes in the environment of the flavin induced by ligand binding (13,44) and thus may represent, in the case of lactate, as is certainly the case with benzoate, the spectrum of E,’ * S after isomerization of E, S has occurred [see Eq. (52) 1. Second, the substrate norvaline shows saturation kinetics in the RHR but fails to give a kinetic isotope effect when deuterated at the a-carbon (83). Since all other substrates which have been so examined give such an isotope effect, i t is not likely that some special feature in flavin reduction results in equal rates of scission of C-’H and C-,H in the rate-determining transition state. Rather, this result most likely reflects rate-determining isomerization of the initial E, * S complex. The electronic spectrum of the rate-determining intermediate in the norvaline reductive half-reaction would probably show whether this interpretation is correct or whether C-H scission occurs before the ratedetermining step. Third, we have emphasized that many flavoprotein oxidase substrates fail to show saturation kinetics in the R H R (i.e., k , is unresolved) and yet the reaction traces are characterized by a single exponential with no indication of an initial lag. This behavior can be formally explained by the condition k-l << k, >> k , [S], with the result that k, is taken to be the rate-determining process in flavin reduction. However, the values of k , so determined ( 2 5 ) , which range from 3 M-’ sec-1 (xylose and glucose oxidase) to a high value of lo5 M-l sec-’ (phenylalanine and L-amino acid oxidase) , are anomalously small when compared with the diffusion-controlled values ( lo8 M-I sec-l) , which are expected on theoretical grounds for such reactant pairs and which are often obtained experimentally by rapid reaction measurements where (unlike the case of flavoprotein oxidases) the initial enzyme-ligand complex can be monitored directly and uniquely (167). Moreover, even when Eq. (15) is taken to describe such cases, it is not possible to fit the data with a diffusion-controlled value for k , , when, as frequently occurs, substrate deuteration affects both the slope and the abscissa intercept of double reciprocal R H R plots ( 5 1 ) . Such anomalies, as well as the behavior of norvaline, benzoate, and anthranilate, are readily explained by isomerization of the initial enzyme-substrate complex as follows :

-- -

- --

- -

- -

N

ki

ki‘

kr

E,+St-IE,...S~E,‘...S~E....Pi k-i

k-i

(52)

166. M.Nishikimi, M.Osamura, and K. Yagi, J . Biochem. (Tokyo) 70, 457 (1971). 167. G. G. Hammes and P. R. Schimmel, “The Enzymes,” Vol. 2, p. 67, 1970.

478

HAROLD J . BRIGHT A N D DAVID J. T. PORTER

The observed pseudo-first-order formation of E, will be given by the following:

*

*

*

PIin the R H R

- -

Having introduced an additional intermediate (E X) it is, of course, a trivial matter to accurately fit Eq. (53) to RHR data which do not strictly require such an intermediate. However, the significance of Eq. (52) is that it allows both for the features of the binding of norvaline, benzoate, and anthranilate, as already noted, and for the assignment of the large values of k , and k-, (including the case of results from kinetic isotope studies with glucose oxidase), which are now held to be characteristic for diffusion-controlled formation and dissociation of weakly bonded noncovalent complexes. It is believed that Eqs. (52) and (53) may be generally applicable to flavoprotein oxidase catalysis ; for example, a rapidly reversible, but thermodynamically unfavorable, isomerization of E, * S to E,’ * S will result in the expression of kl,Bpp = k2/K,Kl‘, instead of k , , to describe the bimolecular interaction of S with enzyme. Other special cases of Eq. (53) are readily apparent. The molecular processes underlying the isomerization of E, S are obscure, as the label implies, although such isomerizations appear to be very common in all classes of enzymes. One may speculate that it represents an example of the concept of “togetherness” as reviewed by Jencks and Page recently (168). I n this case, the optimization of snugness of fit, which can be expected to give very large rate enhancements (compared to the nonenzyme-catalyzed reaction) from entropic factors alone, will refer to the precise alignment of substrate, flavin, and acid-base residues * S and which is driven which takes place during isomerization of E, by the free energy of binding of S to E,. Alternatively, the isomerization may represent an acidification of the carbon-bonded hydrogen of the substrate achieved through a distortion of the tetrahedral sp3 carbon configuration to one closer to trigonal sp2 geometry, which would also be driven by energy acquired from the initial binding of S to E,. The formation of a discrete carbanion from such weakly acidic physiological substrates as glucose [pathways 9[14], 8[12,6], or 8[3,10] of Eq. (51)] would certainly require an acidification mechanism of some kind in order to allow for the rate of proton abstraction by an enzyme base to be compatible with observed turnover numbers of up to los sec-’. Although the increase of s-orbital character in C-H, through bond distortion, is known to increase the acidity of alkanes (169), uncertainties concerning the pK.

--

- -

-- -

-

-

168. W. P. Jencks and M. I. Page, FEBS Symp. 29, 96 (1972). 169. G. L. Closs and L. E. Closs, JACS 85,2022 (1963).

7.

479

FLAVOPROTEIN OXIDASES

values of carbon-bound hydrogen in free or enzyme-bound flavoprotein oxidase substrates and the actual amount of energy available from the binding of typical substrates make this a highly speculative proposal. Furthermore, the acid strengthening effect of an electrophilic region of the flavin nucleus (such as N-5) which might come into play during concerted proton removal and adduct formation [pathway 4[10] of Eq. (51) ] is entirely unknown. In summary therefore, isomerization of E, * * * S in flavoprotein oxidase reactions is a reasonable hypothesis, but the molecular details and function of this process can only be a matter of speculation. b. T h e Chemical Identity of E , * P I , the Long Wavelength (Purple) Intermediate. A number of long wavelength-absorbing flavoprotein intermediates were reported for the first time by Beinert in 1957 shortly after he' had observed similar spectral changes during the oxidation of FMNH, by 0, (84,170). An incisive interpretation of the free flavin spectra was given by Massey and Gibson (36) who confirmed Beinert's assignment of 570 nm absorbance as resulting from the flavin semiquinone and proposed that the 900-nm species was a charge-transfer complex between F M N and FMNH,. Since the work of Beinert almost 30 papers have appeared which describe transient and stable complexes of flavoproteins of all kinds, involving both oxidized and fully reduced forms of the enzymes, and which share the common property of having weak and broad absorbance bands (or, at least, weak absorbance) a t wavelengths greater than 450 nm (the longest wavelength absorption maximum of free or enzyme-bound flavin). A useful compilation of such examples is given by Massey and Ghisla (171). Our concern here is the chemical identity of E, * P, in the amino acid oxidase reactions since this has a great bearing on chemical mechanism. It should be recalled that E, P, is not detected in the glucose oxidase reaction (although it might still be kinetically important). The following facts are established in the amino acid oxidase reactions:

-

- -

- - -

-

1. The substrate a-hydrogen has been removed prior to E, * * PI formation (44,46) as evidenced by large deuterium kinetic isotope effects S to E, * * PI. attending the conversion of E, * 2. The E, * P, intermediate contains one mole equivalent of substrate per flavin. Thus, Yagi and Ozawa (172) have analyzed the crystalline purple complex (E, * PI) prepared anerobically from D-amino acid oxidase and have found levels of pyruvate, after extensive washing

- -

- -

-

- -

170. H . Beinert, JACS 78, 3532 (1956). 171. V. Massey and S. Ghisla, Ann. N.Y. Acad. Sci. 227, 446 (1974). 172. K. Yagi and T. Ozawa, BBA 81, 29 (1964).

480

HAROLD J. BRIGHT AND DAVID J. T. PORTER

of the crystals, which approach one mole per mole FAD. A more direct method was used which measured the amplitude at 370 nm (an approximate isosbestic point for E, * PI and E,) in the stopped-flow spectrophotmeter after anaerobic addition of alanine (39). This titration method is feasible because k , is very small [see Eq. (20)] and the forma* tion E, P, (because of the smallness of k-, and k-J is irreversible A plot of AaTOvs. alanine consisted of two straight line segments, the second (of zero slope) intercepting the first in an alanine concentration precisely equal to the concentration of bound flavin. 3. The electronic spectrum of E, * P1depends on the R group of the substrate (36,38,.44) and shows much greater similarity to the spectrum of fully reduced flavin than to that of oxidized flavin. 4. The physical and chemical properties of a large number of covalent adducts between various ligands and positions of the oxidized and reduced flavin nucleus are well known, chiefly through the work of Hemmerich and his colleagues (6). Certain cationic adducts a t N-5 (7,173) and hydroxyl substitutions a t C-6 and C-9 (174,175) having flavin in the oxidized state, possess long-wavelength absorbance bands. However, their spectra are entirely different from that of E, * P, and, in the absence of any evidence for the presence of a second two-electron-accepting prosthetic group in the amino acid oxidases, these cannot be considered as models for E, * * P,. Adducts involving reduced flavin and substituted at N-1, N-3, N-5, C-2, C-4, C-4a1 C-6, C-8a, or 10-a possess no long-wavelength bands and do not resemble E, * P1 closely (7,l1 ,I 73,175-177). 5. Anaerobic stopped-flow experiments employing phenol red as a p H indicator showed that one proton is lost to solvent during the conversion E, + E, * * * S, but no proton is released or taken up when E, * * S is converted to E, * PI (39). 6. During turnover, the product is released as the a-imino, rather than P, is not generated a-keto, acid (30,44,53,95). Furthermore, E, * when E, is mixed anaerobically with the corresponding a-keto acid in the absence of ammonia.

-

- -

--

-

-

-

-

173. K. H. Dudley, A. Ehrenberg, P. Hemmerich, and F. Miiller, Helv. Chim. Acta. 47, 1354 (1984). 174. G.Schollnhammer and P. Hemmerich, 2.Naturforsch. B 27, 1030 (1972). 175. S. G. Mayhew, C. D. Whitfield, S. Ghisla, and M. Schuman-Joms, Eur. J Biochem. 44, 579 (1974). 176. F. Miiller, in “Flavins and FIavoproteins” (H. Kamin, ed.), p. 363. Univ. Park Press, Baltimore, Maryland, 1971. 177. M. Brustlein, W. R. Knappe, and P. Hemmerich, Angezu. Chena., Znt. Ed. Engl. 10, 804 (1971).

7. FLAVOPROTEIN 7. E,

*

*

-

481

OXIDASES

P, is diamagnetic (S6,46).

These facts unequivocally establish the empirical formula of E, * . PI as (E, 2Hf 2e R-C-C02-) and leave four formal isomers of this

+

+ +

II

NH net oxidation state to be considered [Eq. (54)]. R E-FAD&

*

I .F=NH

(4)

The carbanion complex (structure 3) is extremely unlikely to make a very large contribution to E, P, for two reasons. First, it is inconceivable that such a strongly basic species would have a half-life of about 1 min a t pH 8.5, as is observed for proline and D-amino acid oxidase ( 3 6 ) . Second, the transient complex between E, and the carbanion of nitroethane in the case of D-amino acid oxidase (165), although having long wavelength absorbance has strong bands a t 450 and 375 nm which are little changed from those of E, and which are entirely different from those of E, * P,. Whether E, . P, is best represented by structure 1 (36) or by a rapidly equilibrating mixture of two (178),or more of the species of Eq. (54) is an entirely open question. Although E, * PIis diamagnetic (S6,45), Kosower and others have convincingly demonstrated the existence of radical dimers in model chemical systems which are experimentally diamagnetic (17’9). The weight of opinion, if not experimental evidence, has now shifted

- - -

- -

--

-

-

in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 146. Univ. Park Press, Baltimore, Maryland, 1968. 179. E. M. Kosower, in “Flavins and Flavoproteins” (K. Yagi, ed.), p. 149. Univ. Park Press, Baltimore, Maryland, 1968. 178. V. Massey,

482

HAROLD J. BRIGHT AND DAVID J . T. PORTER

from the biradical structure 2 of Eq. (54) to that of structure 1 of Eq. (54).Massey and Ghisla (171)have argued recently that many, if not all, of the stable and transient long-wavelength absorbing complexes formed by flavoprotein oxidases including E, PIof the amino acid oxidases) and other flavoenzymes are charge-transfer complexes with the oxidized or reduced flavin acting as a 7 acceptor or 7 donor depending on the nature of the ligand. Hemmerich and Schuman Jorns (6) contended that only a few of these complexes conform to the Mulliken criteria (180) of ‘II charge transfer and offered four alternative explanations of the weak long-wavelength absorbance, none of which, however, explicitly embraces the biradical concept of structure 2 of Eq. (54). In the realization that a choice between structures 1 and 2 of Eq. (54) cannot now, or in the foreseeable future, be experimentally achieved with certainty, structure 1 of Eq. (54)will hereafter merely represent an adePI when differenquate working hypothesis for the structure of E, tiation from structure 2 is not a critical issue. When differentiation between structures 1 and 2 of Eq. (54) is critical, it shall so be indicated. c. Deuterium Kinetic Isotope E j e c t on Flavin Reduction. I n all cases tested, with the exception of norvaline and D-amino acid oxidase as noted (82), the spectrophotometric change in the R H R which monitors flavin reduction is associated with a large primary kinetic isotope effect when deuterium is substituted for hydrogen in the C-H bond of the physiological substrate (44,46,51). This result clearly shows that scission of the substrate C-H bond is entirely rate determining in the conversion of E, * S to E, * * P, in these cases. The breaking of this bond, which may be preceded in certain mechanisms of Eq. (51) by an activation brought about by rapid enzyme-catalyzed deprotonation of the substrate ammonium or hydroxyl group, can be taken to represent the initiation of the redox process (as well as its virtual completion if hydride transfer is involved). Hence, the mechanism of flavin reduction by physiological substrates has the kinetic characteristics of one of only four possible steps [see Eq. (51)],namely, hydride abstraction (step 5 ) , hydrogen atom abstraction (step ll), or proton abstractions (steps 4 or 8). It follows that S and E, * . P,, if they exist, will be intermediates between E, * undetectable kinetically [but susceptible, a t least in principle, to chemical trapping as was demonstrated in the case of nitroalkanes (165)1, Consequently, all of the hypothetical redox pathways leading from initial C-H bond scission will be homeomorphic kinetically, and hence indistinguishable, in the case of physiological substrates and native coenzymes. This provides the reason why so much effort is now concerned with substrate analogs, inhibitors, and model (noneneymic) studies.

-

- -

-

-

- -

180. R. S. Mulliken, JACS 58,801 (1952).

7

7. FLAVOPROTEIN OXIDASES

483

d. The p H Dependence of Flavin Reduction. In all cases where the pH dependence of flavin reduction has been studied by rapid reaction techniques, the rate of the redox process is governed by a basic residue in the enzyme. In the case of glucose oxidase, this residue must be in its ionized state in order for substrate binding to occur (26,27,121), whereas in the amino acid oxidase reactions the pH dependence occurs in the flavin reduction step itself (k2)(,?3,44,112). The kinetically determined pKa values vary from 3 to 4 in the case of glucose oxidase, depending upon the halide concentration, to values of 7 or greater in the amino acid oxidase reactions. Being kinetically determined, these apparent pK, values are probably lower limits of the true pKa values. This is particularly obvious in the L-amino acid oxidase reaction (33). Although the catalytic function of the basic residue in the oxidation of physiological substrates is not known with certainty, observations on the oxidation of nitroalkane anions and their conjugate acids by glucose oxidase strongly suggest that it functions in these cases to abstract the carbon-bound substrate hydrogen as a proton. Thus, neutral nitromethane, albeit a highly unreactive substrate, shows the same pH dependence for flavin reduction as does glucose and the other physiological substrates, namely, a sigmoid profile with an apparent pK value between 3 and 5, depending on the halide concentration (26,27,?9,181) . On the other hand, nitroethane anion, from which the carbon-bound proton is removed prior to the kinetic experiments, shows precisely the opposite dependence in that flavin reduction is governed by the conjugate acid state of the same ionizable residue. These results were interpreted to mean that in the case of the anionic substrate, unfavorable charge interactions between the substrate and the conjugate base (which may itself be negatively charged) prevent, or a t least severely weaken, the binding of substrate to the enzyme (181,189). Thus when the carbon-bound proton is absent in the substrate, the basic residue of the enzyme no longer functions in the catalytic process. This shows, at least with the nitroalkanes, that the function of the basic residue which controls flavin reduction in the case of all physiological substrates examined is to abstract the substrate hydrogen as a proton. As with all kinetic evidence obtained from studies of the physiological substrates, the interpretation of the general base control of flavin reduction cannot be uniquely attributed to one of the four rate-determining initiation reactions of Eq. (51). In general, the electron flow from substrate to flavin will be favored by proton abstraction from the former 181. D. J. T. Porter, J. G. Voet, and H. J. Bright, Z. Naturforsch. B 27, 1052 (1972). 182. D. J. T. Porter, J. G. Voet, and H. J. Bright, Fed. Proc., Fed. Amer. Sac. Ezp. Biol. 31, 447 (1972).

484

HAROLD J. BRIGHT AND DAVID J. T. PORTER

and proton addition to the latter and a plausible function for a general base can be visualized for most of these four processes. The proposal that the basic residue abstracts the C-H hydrogen as a proton may be entirely reasonable for the concerted mechanism of step 4 of Eq. (51) since it obviates the problem of high basicity of a discrete carbanion and utilizes the electron-deficient flavin nucleus to assist in the formation and stabilization of the substrate carbanion. 3. Evidence from Enzyme Studies: Native Coenzyme and

Model Substrates The intransigence of the physiological substrates, as previously noted, has forced investigators to study substrate analogs and derivatives in order to obtain evidence for the chemical mechanism of flavin reduction. Two independent lines of investigation with such model substrates have pointed to the probability of abstraction of carbon-bound substrate hydrogen as a proton as an obligatory process in fiavoprotein oxidase catalysis [steps 4 and 8 in Eq. (51) 1, and one of these has demonstrated obligatory covalent adducts derived from attack of the substrate carbanion on N-5 of the flavin nucleus (equivalent to pathway 8[3,10] of Eq. (51) for a physiological substrate). a. p-Chloro-a-amino Acids. The first model substrate to yield any mechanistic evidence whatsoever was p-chloroalanine in the case of D-amino acid oxidase. Miyake et al. (183,184) appear to have been the first to have investigated this interesting substrate. They noted the great dissimilarity between its behavior and that of alanine, namely, that P, was never observed under the characteristic spectrum of E, * * aerobic or anaerobic conditions and that the anaerobic formation of E, was at least 100 times slower than observed with alanine. The steady-state spectrum of the enzyme, whether recorded 6 sec after initiation of the reaction under anaerobic conditions or while the aerobic reaction was in progress, was remarkably similar to that of En,except that a weak longwavelength band extended to beyond 600 nm. The similarity of this spectrum to those of En * * o-aminobenzoate and of E, * A-l-piperidine2-carboxylate (19) was noted. These workers also showed that pyruvate was probably a major product under aerobic conditions, but apparently failed to make the crucial determination as to whether this product was also obtained anaerobically (185). Consequently, they did not clearly

-

-

-

183. Y. Miyake, T. Abe, and T. Yamano, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), p. 209 (and following discussion). Univ. Park Press, Baltimore, Maryland, 1973. 184. Y. Miyake, T. Abe, and T. Yamano, J. Biochem. ( T o k y o ) 73, 1 (1973).

7.

485

FLAVOPROTEIN OXIDASES

demonstrate that the enzyme catalyzes the a-p elimination of HCI from p-chloroalanine in the absence of 0,. In their initial studies, Walsh et al. (31)made the important observation that under anaerobic conditions the enzyme catalyzed the formation of pyruvate and HCl, while a t high concentrations of O2 the product was that expected of the normal oxidative pathway, namely, chloropyruvate. They noted that mixtures of the two products were produced a t intermediate 0, concentrations but that the rate of formation of total a-keto acid (i.e., pyruvate plus chloropyruvate) was approximately independent of the 0, concentration. This was interpreted to mean that a rate-determining intermediate, common to both the elimination and oxidative pathways and reacting directly or indirectly in a fast step with 0,, was responsible for the kinetic control of the type of product obtained. The rate-determining intermediate, on the basis of the deuterium kinetic isotope effect of 1.7-1.9-fold on each of the two pathways, was suggested to be an enzyme-bound substrate a-carbanion or a carbanion-flavin adduct. Schematically, therefore, the results and interpretations of Walsh et aE. (Sl) can be expressed as follows [Eq. (55)], where PIand P, refer, respectively, to the p-chloro-a-imino acid and pyruvate.

E,

-.- a-carbanion

These experiments raise two important questions concerned with kinetics and chemical mechanism. First, is the kinetic scheme of Eq. ( 5 5 ) , as written, correct? Second, regardless of whether or not the scheme of Eq. ( 5 5 ) is correct, what is the validity of extrapolation from the existence and properties of the elimination pathway to the mechanism of flavin reduction? Stopped-flow anaerobic measurements of the interaction of P-chloroalanine with D-amino acid oxidase showed that the scheme of Eq. ( 5 5 ), insofar as the assignment of rate-determining step is concerned, cannot be correct (186). The reaction trace a t 550 nm showed four phases, only the first two of which were rapid enough to represent obligatory turnover processes. Steady-state measurements, whether obtained aerobically through 0, consumption or total keto acid formation, or anaerobically 185. J. G. Voet, D. J.

T.Porter, and H. J. Bright, 2. Naturforsch. B 27,

1054 (1972).

486

HAROLD J. BRIGHT AND DAVID J . T. PORTER

through pyruvate formation (by coupling to lactate dehydrogenase or by direct monitoring of the a-carbonyl a t 322 nm), gave a value for of about 5 sec-', whereas the kinetic constants associated with the last two phases of the stopped-flow experiment were appreciably smaller than 5 sec-'. The first phase of the 550-nm reaction trace was substrate-saturable with an apparent maximum value of a t least 200 sec-' and gave a deuterium kinetic isotope effect (with IY- [ 2H]-p-chloroalanine) of about 4-fold. Clearly, the a-hydrogen is abstracted in the first rapid phase a t a rate 2 200 sec-', whereas the scheme of Eq. ( 5 5 ) requires a-hydrogen abstraction to occur approximately forty times slower than this. It was suggested that the stopped-flow results could be reconciled with $he steadystate parameters if it were assumed that a very large fraction of the enzyme was diverted to an unreactive (or slowly reacting) intermediate during, or soon after, the first turnover. The subsequent work of Walsh et al. with a-amino-p-chlorobutyrate (186) strongly suggests that this intermediate is the enzyme-bound enamine resulting from a-p elimination of HC1. Walsh et al. (186) showed that D-amino acid oxidase also catalyzes the a-p elimination of HC1 from a-amino-p-chlorobutyrate but did not oxidize this substrate a t a detectable rate to a-chloro-p-ketobutyrate. Other reactivity trends should be noted. Thus, for alanine, p-chloroalanine, and a-amino-p-chlorobutyrate, the (p,,-l values are 10, 5, and 0.1 sec-l, respectively; while the +z-l values are 1.3 X lo5, 1.8 x lo4, and not measurable, respectively (39,186). As with p-chloroalanine, the transients observed in stopped-flow experiments with p-chloro-a-aminobutyrate did not correlate with steady-state parameters (186).The first transient was substrate-saturable with a maximum value of 33 sec-1 (and a deuterium kinetic isotope effect of 5-fold, which should be compared with a kinetic isotope effect of about 2-fold on &,-l), the second transient had a value of 0.67 sec-', while the third had a value of about 0.017 sec-l. As with p-chloroalanine, it seems highly unlikely that cleavage of the C-H bond is the rate-determining first-order process in turnover. The answer to these dilemmas in the p-chloroalanine and p-chloro-cu aminobutyrate reactions may well lie in the formation and subsequent tautomerization of enzyme-bound enamine, a species which is formed only in the a-p-elimination reaction. Walsh et al. (186) showed, rather surprisingly, that only 0.5 atom of deuterium was incorporated into the p positions of pyruvate and a-ketobutyrate (racemically in the latter case) from 2Hz0during a-p elimination. The corollary of this result, namely, the demonstration of intramolecular transfer of isotopic hydrogen from the a- to p-carbon, was also achieved. On the reasonable (but as yet 186. C. T. Walsh, E. Krodel, V. Massey, and R.H. Abeles, JBC 248, 1946 (1973).

+,-'

7. FLAVOPROTEIN

487

OXIDASES

untested) assumption that racemic deuteration of 0.5 of the p-carbon in 2H,0 indicates that no more than approximately half of the enamine produced by a-p elimination is released from the enzyme as such, these results show that the conjugate acid of an enzyme base, bearing the proton which originated a t the a-carbon of the substrate, is used by the enzyme to tautomerize the enamine product to the a-imino product while the former is still enzyme-bound. The fact that the "unphysiological" enzyme-catalyzed protonation of the p-carbon of the enamine is able to compete so effectively with the rate of dissociation of enamine from the E, * * enamine complex suggests in turn that the latter process might be extremely slow. The following mechanism [Eq. (56) ] would then apply to both p-chloro-a-amino substrates, although the evaluated rate constants (a-deuterated substrate values in brackets) refer only to p-chloroalanine. We assume the same oxidative pathway (k,and k,) as * X unwas demonstrated for alanine (see Section IV) and leave E * defined, except that it bears the proton originating a t the a-carbon of the p-chloro-a-amino acid substrate.

-

1.7 X los M-l sec-I (1.7 x 109

This mechanism was tested for p-chloroalanine by anaerobic double stopped-flow experiments in which E, was rapidly and irreversibly trapped after one turnover by either anthranilate or benzoate. When the anthranilate (E, I) formation are measured kinetics of E, a t 550 nm after the second mix, two phenomena are observed. First,

--

- - -

488

HAROLD J. BRIGHT AND DAVID J. T. PORTER

E,

* * * I formation is first order (17 sec-l) and exhibits no deuterium kinetic isotope effect with a-[ 2H] -p-chloroalanine. This transient is assigned to k,, the dissociation of imino acid from oxidized enzyme. Second, the total amplitude of this process is dependent on the time a t which anthranilate is added in the second mix. The total amplitude decreases with increasing delay of anthranilate addition and corresponds to a process with no deuterium isotope effect associated with it. This process is assigned as k,, describing deprotonation of the conjugate acid bearing the proton derived from the a-carbon of the substrate by a solvent species X: , with ka = 6.5 sec-1 (after eventual correction for the steady-state mole fraction of E * X). This is the same process as is observed in the third phase of the single mix stopped-flow experiment (186). When benzoate is added in the second mix, on the other hand, E, * I is nonabsorbing, and the disappearance of enzyme intermediates having 550 nm absorbance will be monitored. I n this experiment, the reaction trace was biphasic, the fast phase occurring a t 91 sec-' (and associated with a deuterium kinetic isotope effect of 2.4-fold) ,and the slow phase corresponding to 0.8 sec-' (with no kinetic isotope effect). The fast phase is assigned to k,, the intramolecular trautomerization of enzyme-bound enamine utilizing the proton derived from the substrate a-carbon, while the slow phase is attributed to the very slow dissociation of the enzymeenamine complex. The scheme of Eq. (56) explains several features of the a-@-elimination pathway. First, it predicts for p-chloroalanine a +o-l value of 6.5 sec-l and a kinetic isotope effect of 2-fold on this maximum turnover number under anaerobic conditions. These values are extremely close to the corresponding values determined independently from steady-state kinetic measurements (S1,$9). Second, it explains the apparent discrepancy between the results of stopped-flow experiments and steady-state measurements (186).Thus, the origin of the kinetic isotope effect on +o-l is almost exclusively in k, rather than k,. The major reason for the apparent discrepancy between the transient- and steady-state kinetic results is that as soon as 2 sec into turnover, 50% or more of the enzyme is trapped as the extremely unreactive E, enamine species, as had been originally suggested. Third, the mechanism of Eq. (56) explains why the value of +z-l for p-chloroalanine is approximately 10 times smaller than that for alanine ($9). In the latter case, &-l = k,. However, in the case of p-chloroalanine

-

- -

-

+2-1

kakskio

=

kskm

Thus, the Oz reactivity of E,

+ ksks + krkio = 0.08 *

*

k6

- PI derived from p-chloroalanine is

7.

489

FLAVOPROTEIN OXIDASES

approximately the same as that derived from alanine. The oxidative pathway for p-chloroalanine appears, from the steady-state parameter d2-', enato be inhibited because of the very slow breakdown (klo) of E, mine. This may well be the reason both for the apparent inability of D-amino acid oxidase to oxidize p-chloro-a-aminobutyrate (186) and for the very small maximum turnover number ( c $ ~ - ~for ) that substrate. Both and +2-1, according to Eq. (56),are directly proportional to klo, and one would predict, in the case of p-chloro-a-aminobutyrate, that the enzyme would be almost totally converted to Eo * * enamine after 10 sec or so into turnover [see Fig. 4 of Walsh et al. (18671. Fourth, the relative flux ( k , / h ) through the two anaerobic loops of Eq. (56)with p-chloro-a[2H]-alanine as substrate in 'H,O will give a maximum value for the fraction of intramolecular ((Y+ p ) deuterium transfer [because the pathway involving solvent exchange, as opposed to deprotonation, of the conjugate acid species has been ignored in Eq. (56)l.This fraction is predicted to be 0.85, which can be compared with an experimental value of 0.2 obtained under similar conditions with a-tritiated substrate (f 86). Fifth, compounds having an enaminelike structure, such as anthranilate and piperidine-2-carboxylate, from very stable complexes with EDwhich have absorption spectra remarkably similar to the steady-state spectra obtained during elimination of HCl from the p-chloro-substituted amino acids (f7f).This supports the mechanism of Eq. (56) which, spectro* enascopically, would be dominated by the slowly dissociating E, mine complex after steady state is achieved. Lastly, the right-hand anerobic loop of the scheme of Eq. (56) involves the rate-determining P1, which is dissociation ( k , ) of imino acid product from ED characteristically rate determining in the oxidation of physiological substrates by this enzyme. Moreover, the imino acid derived from p-chloroalanine by a-P HC1 elimination is identical to that derived from oxidation of alanine. Therefore, if the mechanism of Eq. (56) is correct, k8 (17 sec-') should be numerically equal to the corresponding rate constant in alanine oxidation ( k , of Eq. (41), equal to 10 sec-l). Considering the greater technical difficulties associated with the double stopped-flow measurements, these values are in reasonably good agreement. Moreover, it should be noted that during the elimination process the imino acid must dissociate from the conjugate base form of E, - B * P,, whereas during oxidation the imino acid may dissociate, at least in part, from

-

--

-

-

- -

+

ED- B H

--

* P,. These differences in ionization state would be expected to affect the dissociation rates of P,. The importance of the scheme of Eq. (56) is that it shows that the maximum velocity of the elimination reaction is equal in magnitude to

490

HAROLD J . BRIGHT AND DAVID J. T. PORTER

that of the oxidation pathway (ka) for fortuitous reasons (namely, the particular weighting of k,, k,, Ic,, and Ic,, in the +,-l parameter for elimination) and not because they share a common rate-determining intermediate (E * * X) as originally supposed (31). Because of the great significance of the original interpretation of E X as an enzymebound a-carbanion of some kind ( H ) ,the evidence bearing upon the inclusion of E * X as a branch-point species must be carefully scrutinized. Three schemes for oxidation and elimination are compatible with the evidence thus far discussed. These are given in simplified form in Eq. (57), where scheme C is identical to that of Eq. (56). The slow elimination pathway of Eq. (56) has been omitted from Eqs. (57a), (57b), and (57c) to simplify matters.

-

- - -

- -

p~+HclyyJp~+ nation

E, oxidation

0 2

(57%)

S

HC1+ P,

Mechanism (57c) [a simplified version of Eq. (56)] is the only one of the three in which the enzyme-bound substrate in the branch-point species has undergone chemical change (i.e., C-H bond cleavage a t minimum). In schemes (57a) and (57b) the oxidation and elimination pathways are not chemically linked but merely share E, or E, and E, * S. The only evidence which favors scheme (57c) over (57a) and values [+1 = (k1 k , ) / k , k , for scheme (57b) is the fact that the (57c)l for oxidation and elimination are very similar, and each of these values shows approximately the same kinetic isotope effect (which would originate in lc, in this case) with a- [*HI-p-chloroalanine (39).

-

-

+

7.

491

FLAVOPROTEIN OXIDASES

Such equalities would not hold for schemes (57a) and (57b) except through fortuitous relationships between certain rate constants in the two pathways. Thus we are forced to conclude, albeit through limited evidence, that scheme (57c), and hence Eq. (56) , is indeed the best hypothesis for the p-chloroalanine reaction. The second, and perhaps more crucial issue, is the chemical identity X [Eq. (56)] in the p-chloro-aof the branch-point species E * * amino acid reactions because discussions of the chemical mechanism of flavin reduction depend critically on the structure of E * X. The only experimental criterion that needs to be satisfied is that the substrate C-H bond be cleaved in E * X, as evidenced by the deuterium kinetic isotope effect on k, (39,186). There are three possibilities for E * X, namely, E, * * enamine, E, * p-chloro-a-imino acid (E, * PI) or a complex between enzyme and substrate a carbanion (including a carbanion-flavin adduct) or a radical. However, we consider the first possibility to be the most unlikely because it would require that the enzyme sequester C1- and would, moreover, set the P-chloro-a-amino acids as completely unique amino acid substrates. The second possibility, p-chloro-a-imino acid (E, * * P,) could arise by a namely, E, * variety of pathways, as shown in Eq. (51), initiated by rate-determining steps 4, 5, 8, or 11. The third possibility, namely, a.complex of enzyme with substrate a carbanion or a radical, would be formed by the same set of rate-determining steps and would consist of one, two, or all of the complexes in the cyclic pathway of steps 3, 7, and 12 in Eq. (51). The purpose of the preceding discussion has been to emphasize that the demonstration of an a-p-elimination reaction catalyzed by a flavoprotein oxidase according to Eq. (56) cannot be considered as necessary and sufficient evidence for obligatory a-proton abstraction in the oxidative mechanism. This is particularly evident if, as appears to be the case in a recent discussion ( l r l ) ,the branch-point species in Eq. (56) is consid* P, (E, * p-chloro-a-imino acid). The latter is dered t o be E, the final product of, and not a chemical intermediate in, the oxidation pathway and could, in principle, be derived from any one of a number of pathways involving either a-proton, a-hydrogen atom, or P-hydride abstraction [Eq. (51) 1. It might then eliminate chloride as follows:

-

-

-

-

-

- -

-

- -

- -

-

-

- -

492

HAROLD J. BRIGHT AND DAVID J. T. PORTER

Therefore, just as in the case of Eqs. (57a) and 57b), the mechanism of Eq. (57c) does not unequivocally establish a-proton removal as an obligatory step in the oxidative pathway. b. Nitroalkanes. The second line of investigation with nonphysiological substrates has been concerned with nitroalkanes. Nitromethane was the first nitroalkane to be established as a substrate for a flavoprotein oxidase (187).Subsequently, they were found to be good substrates for glucose oxidase and both amino acid oxidases, particularly as their anions. The overall reaction stoichiometry for D-amino acid oxidase is formally analogous to that of amino acid substrates, Eq. (59) (166) : RCHzNO,

+ 0%+ OH-

4

RCHO

+ Hi02 + NO2-

(59)

The conjtigate acids and bases of the nitroalkanes interconvert extremely slowly and thereby afford a unique opportunity to test the effect of proton removal on their reactivity as flavoprotein oxidase substrates. I n all cases, the anions were much more reactive than the Carent acids. I n the case of glucose oxidase, where a valid comparison of pH-independent parameters could be made, nitroethane anion was 10 times more reactive than glucose and loe times more reactive than neutral nitroethane, as measured by and corroborated by stopped-flow measurements (181). The kinetic mechanism for oxidation of nitroethane anion by D-amino acid oxidase was established, through a combination of SSK(02) and stopped-flow studies, to be formally identical to that established for nonbasic physiological substrates [loop B of Eq. (20)]. The analogy extended even to the fact that the release of acetaldehyde from E, * * * PI ' was the principal rate-determining process in turnover. In the original studies with nitromethane, it was noted that the enzyme slowly became inhibited during turnover (187'). The spectrum of the inhibited enzyme resembled that expected for a N-5-alkylated flavin derivative (7).It was reasoned that a covalent flavin-substrate intermediate was being slowly attacked by a second (ionized) nitromethane molecule to form an inactive flavin adduct. As a result of a search for other inhibitory anions, cyanide was found to be a rapid and irreversible inhibitor of nitroalkane oxidation. Further studies were conducted with nitroethane (166) anion because this substrate is far less active than nitromethane anion in inhibiting its own turnover (187). The locus of cyanide inhibition in loop B of Eq. (20) was determined kinetically as follows. First, a t saturating cyanide, both fhe rate of enzyme inhibition and the rate of formation of a flavin-substrate adduct were found by rapid reaction techniques to be precisely equal to the rate 187. D.

J. T. Porter, J. G . Voet, and H. J. Bright, JBC 247, 1951 (1971).

7.

493

FLAVOPROTEIN OXIDASES

of flavin reduction by nitroethane anion ( k , ) in Eq. (60). Second, 0, was not required for inhibition to occur. These two observations limit the locus of cyanide interaction to intermediates lying between E, S and Er * PI including the latter but excluding the former. E, P, was eliminated as a candidate by two observations. First, E, * * P, reacts with 0,, and if it were able to react with cyanide also then competition between cyanide and 0, should result. Instead, increasing 0, concentrations markedly increased the rate of cyanide inhibition during turnover, a result to be expected if the intermediate EX which is attacked by cyanide is located between E, * S and E, * * PI. Second, cyanide was not inhibitory when it was added to solutions of E, * PI which had been prepared from E, and acetaldehyde. These results established the following scheme for cyanide inhibition of the RHR where EX is the intermediate attacked by cyanide and EI is the cyanide-inactivated holoenzyme.

-

-

- -

-

-

.S

EI

It should be emphasized that EX, although possessing a spectrum highly - * P, (this conclusion distinct from those of E,, E, * - * S, and E,

-

being based on the subsequent identification of E X as a cationic imine adduct between substrate and N-5 of the flavin nucleus) is not detected in stopped-flow R H R experiments because the rate-determining step (k,) of the complete RHR sequence precedes EX. The electronic spectrum of EI (in the wavelength region where aromatic protein residues do not interfere) consisted of a single peak with A, = 332 nm and z = 5.4 X lo3 M-1 cm-l. This is quite unlike the spectrum of any flavoprotein oxidase intermediate discovered heretofore and is, in fact, strongly indicative of alkylation a t the N-5 position of the flavin nucleus ( 7 ) . E I was found to contain one substrate and one cyanide equivalent per flavin but did not contain the nitro group. EI could be resolved by methanol treatment into apoenzyme and substrate-FAD adduct. Upon resolution of the holoenzyme, small spectral changes occurred, but these wer$ shown to be fully reversible by combining the modified FAD with native apoenzyme. The free substrate-FAD adduct showed a pK, of 6.4, which is characteristic of N-1 ionization and indicates that this position is unmodified. The adduot was assigned as 5-cyanoethyl-1,5dihydroFAD on the basis of comparison of its spectral and ionization

494

HAROLD J . BRIGHT AND DAVID J. T. PORTER

properties with those of a series of model flavin derivatives (7,188-190). This assignment was subsequently confirmed by synthesis of 5-cyanomethyl-l15-dihydroflavinby reductive cyanomethylation (6).It should be noted that molecular orbital calculations suggest that position 5 of the oxidized flavin nucleus (nitrogen or carbon) will be preferentially reactive with an incoming nucleophile ( 2 2 ) . The structure of the isolated flavin adduct, together with the kinetic mechanism for cyanide inhibition, clearly leads to the following chemical mechanism for nitroalkane anion oxidation and cyanide inhibition [ Eq. (61)l. Flavin reduction is initiated within E, * * S by nucleo9

---

(E,

S)

R

-

~ o , -I c- ...FOEk2 Kc k

NO,-

R I C-FADH-E I

H

NO;

I

CN- fast R I

CN-C-FADH-E I

H (EI)

philic attack of the substrate carbanion on N-5 of enzyme-bound FAD, forming 5-nitroethyl-l15-dihydroFAD. This process, characterized by k, [Eqs. (60) and (Sl)], is entirely rate determining in the sequence E, S -+ E, * * PI. Nitrite is rapidly eliminated to give a N-5 cationic imine. This species [EX in Eqs. (60) and (611 would be expected to be highly susceptible to attack by a variety of nucleophiles. In the normal course of catalysis, EX is rapidly hydrated to form N-5-carbinolamine which then eliminates E-FADH, to form enzyme-bound acetaldehyde (E, * * PI).In the presence of nucleophiles other than H,O, such as nitromethane anion (187)or cyanide (166),the irreversible formation of EI [Eqs. (60) and (61)], which is controlled by k,, becomes competitive with the hydration process and the enzyme becomes irreversibly inhibited

--

-

5. Ghisla, U. Hartmann, P. Hemmerich, and F. Muller, Justus Liebigs, Ann. Chem. 73, 1388 (1973). 189. G. Blankenhorn, S. Ghisla, and P. Hemmerich, 2.Naturforsch. B 27, 1038 (1972). 190. W. R. Knappe and P. Hemmerich, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 13, 188.

293 (1971).

7.

495

FLAVOPROTEIN OXIDASES

as EI [Eqs. (60) and (61)]. Nucleophiles other than cyanide (166)and nitromethane (187)anion, such as NH,, mercaptoethanol, ethylmercaptan, hydroxylamine, and hydride (as BH,-) , also react with EX to form N-5 adducts analogous to EI and the carbinolamine of Eq. (61) (39). Irreversible inhibition is observed only in the case of hydride, which forms an adduct having the properties expected of N-B-propyl-1,5-dihydroFAD when 2-nitropropane anion (which is unreactive with EX) is used as substrate. The other nucleophiles (except for ethylmercaptan) behave like H,O in that they add to the cationic imine (EX) and form a tetrahydral N-5 adduct which, through rearrangement, forms E-FADH, and the corresponding enzyme-bound product. In the case of ethylmercaptan, the tetrahedral N-5 adduct reacts directly with 0,, representing the first case (other than 1,5-dihydroFAD) in which such direct oxidation has been demonstrated in an enzymic reaction. These results are summarized in Eq. (62). The factor which determines whether the nucleophile R Z:

I

Z-C-FADH-E

4

I R

(Z = CN-,

H-,CH,NO,)

R, @ ,C =F ADH- E

R

Y:

=

(Y:= H,O, NH,, NH,OH, CH,CH,SH, HOCH,CH,SH)

(inactive) (EI)

7

1

Y- C-FADH-E I

R (active)

L (Y= H,O, NH,, NH,OH

El. *

. .P,

HOCH,CH,SH)

forms an N-5 adduct which is catalytically inactive (nucleophile = Z : ) or active (nucleophile = Y : ) appears to be the absence or presence of a nonbonded electron pair on the atom which is two bonds removed from the N-5 atom in the tetrahedral adduct. As predicted from Eq. (61), nucleophiles of type Y protect against CN- inactivation (39). The demonstration that catalytically active enzyme-bound 2-aminopropyl-1,5-dihydroFAD can be formed in the D-amino acid oxidase reaction from the addition of, Y = NH, during oxidative turnover of 2-nitropropane anion is particularly significant because this adduct is strictly analogous to that which would be formed from a physiological amino acid substrate if the latter were oxidized through an obligatory N-5 flavin

496

HAROLD J . BRIGHT AND DAVID J. T. PORTER

adduct [see Eq. (51)]. When Y = H,O, the N-5-carbinolamine is analogous to the adduct anticipated in the glucose oxidase reaction. Although nucleophiles of type Z [Eq. (62)] form covalent adducts [EI, Eq. ( S l ) ] which are catalytically unreactive under the conditions of ensymological studies, they can become reactive a t higher temperatures. Thus, free 5-cyano-ethyl-1,5-dihydroFAD, formed as a result of cyanide inhibition of nitroethane anion turnover and resolved from the apoprotein by methanol treatment, was found to produce FADH, and acetaldehyde (and cyanide, presumably) when heated anaerobically a t 70° (166). FAD and acetaldehyde (together with cyanide and H,O,) were produced aerobically a t 70°. The free modified flavin, once cyanide is eliminated at elevated temperatures, is therefore capable of undergoing the same net reaction as is normally catalyzed by the apoprotein. This nonenzymic reaction presumably involves formation of the cationic imine, hydration of this species to form the carbinolamine and then elimination of acetaldehyde to form FADH,, as shown in Eq. (61). I n the presence of 02, the latter will rapidly form FAD and H,O,. Model reactions have demonstrated the reversible formation of N-5-carbinolamines from reduced flavin and aldehydes (189,191.). The oxidation of nitroalkanes catalyzed by glucose oxidase differs in some interesting respects from the corresponding n-amino acid oxidase reaction. First, the overall stoichiometry of nitroethane oxidation is nonintegral (and, as yet, incomplete), suggesting the presence of two or more reaction pathways whose chemistry is distinctly different (59,181).Distinctly less than one equivalent of H,O, is produced, together with traces of nitrate and dinitroethane. Second, although SSK (0,) measurements conform to the usual three-term expression [Eq. (5) 1, stopped-flow spectrophotometric measurements of the half -reactions and of the enzyme in turnover clearly showed that two pathways for glucose oxidation exist, both of which are quantitatively significant. Thus, stopped-flow measurements showed that anaerobic reduction of the enzyme produced the fully reduced and semiquinone flavin enzyme species in the relative amounts of 0.65 and 0.35. The kinetic data do not differentiate between a common branch-point intermediate (such as the N-5-nitroalkyl-1,5-dihydroFAD adduct formed in the D-amino acid oxidase reaction) and a reaction * S. Both of these species scheme which branches at the level of E, * undergo turnover in the presence of 0,, by separate pathways, to regenerate E,. This appears to be the first authentic example of the involvement of kinetically competent free radical intermediate in a simple flavoprotein oxidase reaction. The present state of cur knowledge concerning the chemical mechanism 191. S. Shinkai and T.C.Bruice, JACS 95, 7526 (1973).

-

7.

FLAVOPROTEIN OXIDASES

497

of flavin, insofar as this derived from studies of p-chloroamino acids and nitroalkanes, may be summarized as follows. The fact that a variety of flavoproteins catalyze the a-p elimination of HC1 from p-chloro-substituted substrates is not, ips0 facto, clear evidence for a-proton abstraction as the obligatory first step in the chemical mechanism of flavin reduction. Similarly, ambiguities exist in the nitroalkane oxidation mechanisms. Whereas carbanions and N-5 flavin-carbanion adducts are quite clearly, by all experimental criteria, competent kinetic intermediates in the oxidation of nitroethane anion by D-amino acid oxidase, a free radical pathway is equally clearly involved in part in the oxidation of nitroethane anion by glucose oxidase. The latter result raises the serious question as to whether the mechanism of flavoenzyme reduction by nitroalkane anions ought not to be viewed,, according to Eq. (51), as one in which the initial E, * * carbanion complex has access to two favorable kinetic pathways, namely, rate-determining adduct formation through step 3 [ as shown in Eq. (51) J or one-electron transfer to flavin [step 12 of Eq. (51)] followed either by step 6 or steps 7 and 10. Whereas AF$ for the transition states of pathways 3 and 12 in the glucose oxidation reaction would be almost identical according to this viewpoint, even the corresponding hF$ values in the D-amino acid oxidase reaction need not differ greatly (in favor of AF$ for step 3) to be reconciled with spectrokinetic and chemical trapping data (165) which, by their nature, would not detect a competing free radical pathway comprising up to 5% of the total turnover flux. This matter will be returned to in the discussion of model flavin redox systems in Section V,A,5.

-

4. Evidence f r o m Enzyme Studies: Inhibitors and Coenzyme Analogs

Several studies have been published recently which utilize the carbon analog of isoalloxazine, namely, 5-deazaflavin. The rationale for the use of this model flavin is that carbon-bonded substrate hydrogen which might be transferred to C-5 of deaza-flavin would not equilibrate with solvent protons. The first of these studies demonstrated that the 4-C-H of NADH was transferred directly to the C-5 position of 5-deaza-Aavin in aqueous solution (19%’).Bruice now suggests (19s)that this reaction in particular, and oxidations of dihydropyridines in general, occur via electron transfer followed by hydrogen atom abstraction [i.e., reverse of Eq. (66) with dihydropyridine in place of methanol] as first suggested by Kosower (14). Subsequently, NADH-FMN oxidoreductase was shown 192. M. Briistlein and T. C . Bruice, JACS 94,6548 (1972). 193. R. F. Williams, S. Shinkai, and T. C. Bruice, Proc. Nut. Acud. Sci. U. S . 72, (1975).

498

HAROLD J. BRIGHT AND DAVID J. T. PORTER

to catalyze direct hydrogen transfer from NADH to 5-deaza-riboflavin by Fisher and Walsh (194)and N-methylglutamate synthetase, containing 5-deaza-FMN, in place of FMN, was demonstrated by Schuman Jorns and Hersh to incorpoate the a-hydrogen of glutamate into the enzymebound deaza-FMNH, and to subsequently transfer this hydrogen to the product, N-methylglutamate (195).The results of all these studies are, a priori, consistent with any mode of C-H bond scission shown in Table I, and other experimental approaches will be required to distinguish between the major possible pathways. Acetylenic substrates and inhibitors of flavoproteins are promising tools for the investigation of flavoenzyme reduction mechanisms. Walsh et al. (196)have shown that the substrate 2-hydroxy-3-butynoate inactivates lactate oxidase by forming a substrate-flavin adduct in which the C-2 substrate hydrogen is missing. The structure of this adduct may suggest the pathway for its formation. The inhibition of monoamine oxidase by acetylenic amines such as pargyline results in the formation of an undissociable enzyme-inhibitor complex which has been shown to be a covalent, flavin-inhibitor adduct (139,153).Cycloaddition products of flavin and acetylenic compounds have been prepared photochemically and suggested to be models for the enzymic counterparts (154).Whether this is so will require full structural characterization of the enzyme adducts. 3-Bromoallylamine has also been shown to be a potent inhibitor of monoamine oxidase (197).Whether this involves covalent addition to the coenzyme or apoeneyme, or t o both, requires much more structural information than is currently available.

5. Evidence from Chemical Model Systems Until quite recently, the mechanistic aspects of both flavoenzyme and model flavin systems were so obscure that neither line of inquiry was of much assistance to the other. Recently, however, owing in no small measure to the influence of Hemmerich and his colleagues in emphasizing the highly electrophilic nature of N-5 and C-4a of the flavin nucleus (6), the two lines of inquiry have advanced enormously and have converged to the extent that a highly productive interplay now exists between them. This discussion shall be confined rather strictly to model redox reactions involving the isoalloxazine nucleus and compounds (hydroxy derivatives and nitroalkanes) which closely resemble the substrates of the simple 194. J. Fisher and C. Walsh, JACS 96, 4345 (1974). 105. M. Schuman Joms and L. B. Hersh, JACS 96, 4012 (1974). 196. C. T. Walsh, A. Schonbrunn, T. Lockridge, V. Massey, and R. H. Abeles, JBC 247, 6004 (1972). 197. R. R. Rando, JACS 95,4438 (1974).

7.

499

FLAVOPROTEIN OXIDASES

flavoprotein oxidases. A broader review of mechanistic aspects of model flavin studies has appeared recently (198). We should first emphasize the highly significant difference between the thermodynamics of the model half-reaction [Eq. (63)] and that of the enzymic reductive half-reaction [e.g., Eq. (11)1. Taking E,’ = -0.21 F,

+

I I

- C - X H c

F,

+

\

,C=X

(63)

V for a typical free flavin such as riboflavin or lumiflavin (199) and computing E,’ = -0.32 V for gluconolactone/glucose from the data given by Strecker and Korkes (200), the reaction is exergonic as written a t pH 7 to the extent of - 5 kcal while for the oxidation of methanol to formaldehyde [E,’ = -0.190 V ( 2 0 1 ) ] , reaction (63) is endergonic by 1.5 kcal. The flavoprotein oxidases, however, are far better oxidizing agents than free flavin (202,203).Thus, if either glucose oxidase (203) or D-amino acid oxidase (202) (both of which give E,’ = 0.01 V a t pH 7) were to catalyze the reaction of Eq. ( 6 3 ) ,it would be highly exergonic in both cases, amounting t o -14 kcal for glucose and -8 kcal for methanol. The major consequence of these thermodynamic differences has been that whereas the enzymic reaction of Eq. (63) can be reversed only with great difficulty (204), most model reactions involving noncyclic carbonyl/alcohol pairs (e.g., formaldehyde/methanol) have been carried out in the right to left direction of Eq. (63). As a result, the rate-determining steps in the model and enzymic pathways, even if these pathways were identical, are on opposite sides of the highest transition state and therefore cannot easily be compared. This would not be such a serious problem were it not for the strong possibility [Eq. (51)] that experimental differentiation between heterolytic and homolytic pathways following the ratedetermining step will prove t o be very difficult. However, this problem has recently been solved in model studies through the use of the highly electron-withdrawing cyano substituent on the flavin nucleus (2051, and

-

198. T. C. Bruice, Progr. Bioorg. Cheni. (in press). 199. R. D. Draper and L. L. Ingraham, ABB 125, 802 (1968). 200. H. J. Strecker and S. Korkes, JBC 196, 769 (1952). 201. W. M. Latimer, “Oxidation Potentials,” 2nd ed., p. 130. Prentice-Hall, Englewood Cliffs, N. J., 1952. 202. M. Brunori, G. Rotilio, E. Antonini, B. Curti, U. Branzoli, and V. Massey, JBC 246, 3140 (1971). 203. F. R. Duke, R. N. Kust, and L. A. King, J . Electrochem. SOC. 116,32 (1969). 204. A. N. Radhakrishnan and A. Meister, JBC 233, 444 (1958). 205. I. Yokoe and T. C. Bruice, JACS 97,450 (1975).

500

HAROLD J . BRIGHT AND DAVID J. T. PORTER

it is to be hoped that the model and enzymic reactions can at least be studied in the same sense in the future. The large difference in redox potential between free and enzyme-bound flavin must originate from much stronger interactions between the enzyme and reduced flavin as compared with those between enzyme and oxidized flavin. This factor is difficult to measure through equilibrium binding experiments because of the very great affinity even of oxidized f l a h However, the dissociation constant of E-FADH, has been tentatively estimated to be lo6 times smaller than that for E-FAD in the case of redox potential measurements with D-amino acid oxidase (202). In the case of the binding of F M N and FMNH, to old yellow enzyme, the corresponding factor is 2 X lo2 in favor of FMNH? binding (206). One of the first discrete mechanisms proposed for flavoprotein dehydrogenations on the basis of model studies was that of Brown and Hamilton (207,208).These authors studied the anaerobic oxidation of C6H5CH(XH)--CO,CH, (X = 0 or N H ) and other substrates by 10-phenylisoalloxazine under strongly basic and anhydrous conditions and isolated the products expected for a two-electron oxidation process. Although kinetic studies were not carried out and intermediates were not detected, a general mechanism for flavoenzyme-catalyzed dehydrogenation was proposed in which the electronegative substrate group -XH adds to C-4a of the flavin nucleus to form a covalent adduct. The substrate a-hydrogen is then removed as a proton in concert with the elimination of reduced flavin. Although the mechanism of Hamilton (207,208)has had the admirable effect of spurring a renewal of interest and experimental effort in the problem of flavin reduction and, indeed, has been invoked often in discussions of the enzymic reactions (sometimes, mistakenly, as a mechanism which stabilizes a substrate carbanion), it is not likely to be a correct description of model or enzymic reactions which conform to Eq. (63) for the following reasons. First, the reversal of Eq. (63), which occurs a t appreciable rates in both the model and enzymic systems, would have to ,

be initiated through nucleophilic attack of C-4a on atom X in

\ C=X /

(X = 0 or NH) and proton addition to carbon. Such a reactionis probably without precedence in bioorganic chemistry (191). Second, Rynd and Gibian showed a t almost the same time (209) that enediolates (and other 206. C. S.Vestling, Acta Chem. Scand. 9, 1600 (1955). 207. L. E. Brown and G. A. Hamilton, JACS g.2, 7225 (1970). 208. G. A. Hamilton, Progr. Bioorg. Chem. 1, 83 (1971). 209. J. A. Rynd and M. J. Gibian, BBRC 41, 1097 (1970).

7.

FLAVOPROTEIN OXIDASES

501

carbanions) are readily oxidized by flavins to a-diketones in aprotic basic solutions and pointed out that substrate carbanion formation might initiate oxidation in flavoenzyme systems. However, they observed rapid production of flavin anion radical as well as fully reduced flavin and were unable to determine whether the overall oxidation proceeded in one two-electron or two one-electron steps. As was subsequently pointed out (6) the mandelic ester studied by Brown and Hamilton (207), which is an a-ketol, might well react with flavin as a n enediolate rather than through C-4a adduct formation. Third, these questions appear to have been clearly resolved by the recent work of Shinkai et al. (210) which shows that a-ketols such as furoin and benzoin are indeed oxidized by a variety of oxidizing agents (including flavin) through rate-determining formation of the corresponding enediolate. The C-4a mechanism (207,208)therefore appears to be an unlikely possibility for simple flavoprotein oxidases and, consequently, is not included in the scheme of Eq. (51). Whether the processes following substrate (carb-)anion formation in the model reactions should be regarded as hydride transfer (in the case of a-/I unsaturation), one two-electron transfer, or two one-electron transfers is another matter. Bruice and his colleagues favor the latter possibility (210). However, the concept of a C-4a adduct (207) may be valid for redox reactions which do not conform to Eq. (63), such as 2 RSH + RS-S-R. Thus, sulfite can add either to N-5 or C-4a (211)and the oxidation of thiophenol is attributed to C-4a adduct formation (2U5). Furthermore, migration of certain cations between N-5 and C-4s has been demonstrated (612-214). These results seem to indicate the feasibility of enzymic control of either N-5 or C-4a adduct formation, depending on the type of reaction to be catalyzed. Model studies of nitroalkane anion oxidation by flavin would be of great interest because they should afford a detailed comparison with the chemical mechanism of flavin reduction by these substrates in the flavoprotein oxidase reactions (165). This, in turn, would show whether the enzymes merely improve the nonenzymic pathway or follow an entirely different one. Such model studies have not been feasible heretofore because the E,’ values are not favorably matched. Recently, however, Yokoe and Bruice (205) have solved this problem through the synthesis of an electron-deficient flavin, namely, 3,10-dimethyl-8-cyanoisoalloxa210. 211. 212. 213. 214.

S. Shinkai, T. Kunitake, and T. C. Bruice, JACS 96, 7140 (1974). L. Main, G. Kasperek, and T. C. Bruice, Biochemistry 11, 3991 (1972). W. Haas and P. Hemmerich, 2.Naturforsch. B 27, 1035 (1972). W. H. Walker, P. Hemmerich, and V. Massey, Eur. J . Biochem. 13,258 (1970). D. Clerin and T. C. Bruice, JACS 96,5571 (1974).

502

HAROLD J. BRIGHT AND DAVID J . T. PORTER

zine. This flavin oxidizes nitroalkanes, in aqueous solution and in the pH range used for enzymic studies, according to the stoichiometry of the enzymic reaction [Eq. (63)]. The mechanism of the model reaction, on the basis of its kinetic order and lack of general acid-base catalysis [the latter criterion rests only on studies with sulfite (215) and ought to be verified with other nucleophiles] was suggested as proceeding either through a N-5 adduct, as demonstrated in the case of D-amino acid oxidase (165), or via a free radical process, as shown in the glucose oxidase reaction (182) (see discussion in Section V,A,3,b). As Yokoe and Bruice pointed out (205) and as emphasized many times in the discussion here, carbanion oxidation pathways involving N-5 alkylation (covalent adduct formation) on the one hand, and free radical processes on the other, need not be mutually exclusive. Quite apart from questions of kinetic indistinguishability, there exists chemical precedence for a free radical pathway in alkylation by carbanions (or enediolates) derived from nitroalkanes and a-hydroxycarbonyl compounds (616,217).Thus, Kerber et al. (216) have proposed the following mechanism for the alkylation of p-nitrobenzyl chloride (R-CH,Cl) by 2-nitropropyl anion [ (R') &NO,-]. The

electron pair which is lost from the radical anion as chloride in Eqs. (64) would, in a flavoprotein oxidase system, be accommodated intramolecularly by N-1 of the flavin nucleus. If a mechanism similar to Eqs. (64) were to apply to the model (205) and enzymic (165) nitroalkane oxidations, then (64a) would be rate determining and one would have a "hidden radical" as proposed in slightly different contexts (193,213). Perhaps the most important contribution of model studies to date concerns the anaerobic reduction of formaldehyde to methanol by 1,5-dihydroflavin. This reaction was first studied by Blankenhorn et al. (189) and was postulated to occur via a N-5-carbinolamine (5-CA) as shown by steps 10 and 4 in Eq. (65). The presence of 5-CA was later confirmed, but its role as an obligatory intermediate was ruled out by transient kinetic studies of the formation of oxidized flavin (191). The kinetics conformed instead to the mechanism of Eq. (66) in which the 5-CA spe-

c.

215. T. Bruice, L. Hevesi, and S. Shinkai, Biochemistry 12, 2083 (1973). 216. R. C. Kerber, G. W. Urry, and N. Kornblum, JACS 87, 4520 (1965). 217. G. A. Russell and R. K. Norris, R e v . Reactive Org. React. 1, 65 (1972).

7.

503

FLAVOPROTEIN OXIDASES

cies is nonproductive. At face value, this kinetic mechanism appears as hydride transfer, controlled by k3‘. 5

-

C

A

c F,

+

\

,C=O-

k’,

Fo

I

f H-C-OH

(65)

I

Very recently, Williams et al. (193) have constructed a reaction coordinate diagram for the kinetic scheme of Eq. (66) (using formaldehyde, pyruvate, and ethyl pyruvate in aqueous solution, p H 5-9) by assuming that the chemical mechanism underlying k,’ is entirely free radical in nature with k,’ = k,k,/k-,

Fo

+

CH,OH

Reiative ground state energies of thc intermediates and final and initial states were computed from AFrO’ and Eo’values and from ratios of measured rate constants. AFS values were computed by assigning charand for acteristic values for electron transfer from a radical anion (L) diffusion-controlled proton transfer (k3) and then computing lcz from the experimental Ic,’ value. The value of k4 was estimated from H abstraction data in the literature. The important result of this proposal, which appears for the most part to be based on sound assumptions, is that the AFO’ values for radical intermediates do not exceed, in any instance, the computed A F t values. Williams et al. therefore concluded that “it is difficult to imagine how radical mechanisms cannot be involved” (193).The predictive value of Eq. (66) will of course ultimately determine whether such confidence is warranted. However, initial tests, in which the addition of ethyl pyruvate to N-5-methyl-l,5-dihydroflavin was reported to generate N-5methyl-FA, appear very promising (193).

-

B. THEMECHANISM OF OXIDATIONOF REDUCED FLAVIN BY 0, I n all flavoprotein oxidases examined, the oxidation of E, by 0, yields E, and H,O, in a strictly bimolecular reaction characterized by k , = 104-106M-’ sec-*. Examination of half-times for the oxidation of FMNH, by 0, (218) clearly shows that the overall rate of the model 218. Q . H. Gibson and J. W. Hastings, BJ 83, 368 (1962).

504

HAROLD J. BRIGHT A N D DAVID J. T. PORTER

reaction is of the same order of magnitude as that of the enzymic reactions. By this rough comparison, therefore, it is evident that the flavoprotein oxidases offer little or no catalytic assistance to this process and that, in principle a t least, studies of model reaction mechanisms should be capable of direct extrapolation to flavoprotein oxidases. Furthermore, knowledge of the site of 0, attack on the flavin nucleus would then suggest the mechanism by which flavoprotein dehydrogenases are able to largely inhibit the interaction of 0, with reduced flavin. However, the model reactions are distinctly autocatalytic and 0z1 is a major transient product (218,219). The major cause of autocatalysis appears to be the formation of highly reactive semiquinone through disproportionation of the fully oxidized and fully reduced flavin. Disproportionation of E, and E, in the case of flavoprotein oxidases takes place on a time scale of hours rather than milliseconds (38)and is therefore not a factor in their oxidation by 0 2 . The observed difference in reduction products of O2is quite generally thought to center around the fate of an initial covalent adduct between reduced flavin and O2 ( 1 1,219) [Eq. (67). The formation of this adduct is regarded as activation of 02.

-/

flavoprotein dehydrogenases

H,F’

+

0; t H’

model reactions

i

f lavoprotein hydroxylases H+

[OH+]+ HF,

+

H,O

I n the uncatalyzed model reaction, as well as the “unphysiological” oxidation of flavoprotein dehydrogenases, homolytic cleavage of the adduct is kinetically most favorable. The flavoprotein oxidases, presumably through direction by general acid-base catalysis, cleave the adduct heterolytically between flavin and oxygen. The flavoprotein hydroxylases are presumed to generate OH+or its chemical equivalent through heterolytic cleavage of the bonds between the oxygen atoms in the adduct. The structure of the flavin-0, adduct is not known. Massey et al. have suggested either 10a or 4a adducts (219; see also Chapter 4 by Massey 219. V. Massey, G. Palmer, and D. Ballou, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), p. 25. Univ. Park Press. Baltimore, Maryland, 1973.

7.

505

FLAVOPROTEIN OXIDASES

and Hemmerich this volume). The former seems unlikely in view of the rapid oxidation of reduced flavin derivatives which are sterically blocked at the l a position (198). Hemmerich and Miiller appear to favor an adduct a t C-6 (11). Entsch et al. (220) have detected a spectrophotometric species in a flavoprotein hydroxylase reaction which may correspond to a flavin-0, adduct. Whether such an adduct was formed with singlet 0, as was claimed in other studies (921) is obscured by the fact that i t is not evident from the data presented that the starting flavin was actually N-5-benayl-1,5-dihydroflavin.

ACKNOWLEDGMENTS Preparation of the chapter and experimental studies in the authors’ laboratory were supported in part by the National Institutes of Health, Grant GM 11040. The authors wish to thank Dr. Thomas C. Bruice for discussions of flavin chemistry and for sending them manuscripts prior to publication. 220. €3. Entsch, V. Maeaey, and D. P. Ballou, BBRC 57, 1018 (1974). 221. M. Yamasaki and T. Yamano, BBRC 51,612 (1973).

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Cof$er-Containing Oxidases and Superoxide Dismutase B . G. MALMSTROM L.-E. ANDREASSON B . REINHAMMAR I. Introduction . . . . . . . . . . . . . . . . I1. Enzymes Reducing Dioxygen to Hydrogen Peroxide . . . . A . Introduction . . . . . . . . . . . . . . . B Amine Oxidases . . . . . . . . . . . . . . C . Galactose Oxidase . . . . . . . . . . . . . TI1. Superoxide Dismutase . . . . . . . . . . . . . A . Introduction . . . . . . . . . . . . . . . B . Purification and Assay Methods . . . . . . . . . C . Molecular Properties . . . . . . . . . . . . D . The Catalytic Mechanism . . . . . . . . . . . IV. The Blue Copper-Containing Oxidases . . . . . . . . A . Introduction . . . . . . . . . . . . . . . B . Purification and Some Molecular Properties . . . . . C . The Forms of Copper: Magnetic and Spectroscopic Properties D . Oxidation-Reduction Properties . . . . . . . . . E . Catalytic Properties . . . . . . . . . . . .

.

. . . . .

. .

. . . . . .

. .

.

507 511 511 511 527 533 533 538 542 552 557 557 560 563 571 574

.

1 Introduction

Many areas of what might be called metallobiochemistry were initially approached in pioneering explorations by D. Keilin in Cambridge. The important role that we now know copper to play in biological oxidations was first surmised from his work in the late 1930’s. Thus, in 1938. Keilin and Hartree ( I ) summarized evidence for cytochrome oxidase being a 1 . D. Keilin and E . F. Hartree. Nature (London) 141. 870 (1938) . 507

508

B. G. MALMSTROM, L.-E. ANDREASSON, AND B. REINHAMMAR

copper-protein compound. In the same year Keilin and Mann ( 2 ) showed that copper is a component of tyrosinase, and a year later they reported (3) the same for laccase. Of course, the enzymes involved had been known much longer ; and, indeed, the very term “oxidase” had been introduced by G. Bertrand in the last decade of the nineteenth century in connection with his work on laccase and tyrosinase. I n fact, Bertrand (4) had also suggested that laccase is a metalloprotein, although he incorrectly identified the metal involved as manganese, and he was thus the first to introduce the concept of a metal as an essential constituent of oxidizing enzymes [for a fuller account of these historical aspects, see, for example, Keilin (6)and Fruton ( 6 )1. The key biological function of copper is undoubtedly its involvement in the cytochrome c oxidase of the mitochondria1 respiratory chain. Because of its importance this enzyme is the subject of Chapter 5, Volume XIII, and will consequently not be treated here. Copper is, however, now known to be an essential component in several oxidases that play important roles in more peripheral parts of the metabolism of microorganisms, plants, and animals. The copper-containing oxidases known today are listed in Table I. They can be divided into two main groups. Most of the enzymes do not utilize the full oxidizing power of dioxygen, which in these cases is reduced to hydrogen peroxide only. On the other hand, a few enzymes can, like cytochrome c oxidase, catalyze reactions in which both atoms of dioxygen are reduced to water. This group is often referred to as the “blue oxidases” because of the beautiful, strong blue color associated with one class of copper ions of the enzymes (see Section IV). Many readers may be surprised not to find tyrosinase included in Table I (7). This enzyme is a special case, however. It is true that it can act on certain diphenols, in which cases dioxygen is reduced to water, but 2. 3. 4. 5.

D. Keilin and T. Mann, Proc. Roy. Soc., Ser. B 125, 187 (1938). D. Keilin and T. Mann, Nature (London) 143, 23 (1939). G. Bertrand, C. R . Acad. Sci. 118, 1215 (1894). D. Keilin, “The History of Cell Respiration and Cytochrome.” Cambridge Univ.

Press, London and New York, 1966. 6. J. S. Fruton, “Molecules and Life.” Wiley (Interscience), New York, 1972. 7. International Union of Biochemistry Commission, “Enzyme Nomenclature. Recommendations of the Commission on Biochemical Nomenclature.” Elsevier, Amsterdam, 1973. The classification and nomenclature of copper-containing oxidases admittedly preHent difficult problems, but the recommendations would appear inexcusably confusing (cf. Table I). The earlier (1965) classification was somewhat better since it recognized the welldocumented difference between tyrosinase and laccase and did not put ceruloplasmin in a category separate from the other blue oxidases. The recommended names were, however, misleading since they implied a high specificity. Here the well-understood and established names, such as laccase, will be used exclusively.

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

509

TABLE I THECOPPER-CONTAINING OXIDASES Enzyme0 Producing H202 Amine oxidases Galactose oxidase Producing HsO (blue oxidases) Ascorbate oxidase Ceruloplasmin Laccase

Enzyme Commission recommended name and numbeP

Amine oxidase (pyridoxal-containing)c; EC 1.4.3.6 Same; E C 1.1.3.9 Same; E C 1.10.3.3 Ferroxidased; EC 1.16.3.1 Not includede

,. Commonly used name, employed also here (7).

~

See reference 7. Not all copper-containing amine oxidases have pyridoxal (see Section 11). See Section IV,E,2,b. Except, erroneously, as a trivial name synonym _for monophenol monooxygenase (tyrosinase), EC 1.14.18.1. I t was included in the earlier (1965) recommendations as EC 1.10. 3.2 under the name p-diphenol: oxygen oxidoreductase. b

its physiological substrates are believed to be monophenols. When monophenols are oxidized, one of the atoms of dioxygen is incorporated into the substrate, so that in these reactions tyrosinase functions as a monooxygenase (“mixed-function oxidase”) . Consequently it is discussed in Chapter 5. Table I includes only those enzymes which have quite unambiguously been shown to be copper proteins. For some enzymes often found in tabulations of copper-containing oxidases (for example, uricase) , there are, however, conflicting reports, as will be briefly discussed in Section 11. Two symposia have dealt specifically with the biochemistry of copper (8,9).Copper-containing proteins, particularly the blue oxidases, have been the subject of recent reviews (10-12’). Some relevant articles can also be found in the proceedings from an oxidase symposium (IS) and 8. W. D. McElroy and B. Clam, eds., “Copper Metabolism.” Johns Hopkins Press, Baltimore, Maryland, 1960. 9. J. Peisach, P. Aisen, and W. E. Blumberg, eds., “The Biochemistry of Copper.” Academic Press, New York, 1966. 10. R. Malkin and B. G. Malmstrom, Advan. Enzymol. 33, 177 (1970). 11. T. Vannglrd, in “Biological Applications of EPR” (H. M. Swartz, J. Bolton, and D. Borg, eds.), p. 411. Wiley (Interscience), New York, 1972. 12. R. Malkin, in “Inorganic Biochemistry” (G. L. Eichhorn, ed.), Vol. 2, p. 689. Elsevier, Amsterdam, 1973. 13. T. E. King, H. S. Mason, and M. Morrison, eds., “Oxidases and Related Redox Systems.” Univ. Park Press, Baltimore, Maryland, 1973.

510

B. G. MALMSTROM, L.-E. ANDR~ASSON, AND B. REINHAMMAR

in a current book on oxygen activation (14). I n the following, reference to these surveys will often be made in place of a detailed documentation of early literature, both in the interest of space and to allow an emphasis on current problems of the field. I n the same year that the presence of copper was demonstrated in tyrosinase, Mann and Keilin (15) isolated two copper proteins from bovine erythrocytes and liver, respectively. For a long time these were thought to have a storage function. Thirty years after the discovery of hemocuprein, or erythrocuprein, as the protein from red cells was called, McCord and Fridovich (16) showed, however, that this protein can catalyze the dismutation of superoxide radicals (17) to hydrogen peroxide and dioxygen. They suggested that this catalytic activity represents its physiological function, and that as an enzyme the protein should be named superoxide dismutase (17,19)in the future. Their report kindled a tremendous interest in erythrocuprein and related proteins from other sources, and the ensuing literature is voluminous. The amount of detailed knowledge on the molecular level is, however, still not sufficient for this protein to get a chapter of its own, in terms of the general criteria used in planning “The Enzymes.” Instead, an extensive section (Section 111) on superoxide dismutase is included in this chapter. Since the dismutase is not an oxidase, it may appear that the only logic behind this is that the protein contains copper. Superoxide is, however, the primary reduction product of dioxygen with some oxidases, for example, xanthine oxidase ( 2 0 ) ,and its formation in some reactions involving copper-containing oxidases yielding hydrogen peroxide has been suggested (see Sections I1 and 111).I n addition to the survey in Section 111,other recent reviews of superoxide dismutase are available (21-23). 14. 0. Hayaishi, ed., “Molecular Mechanisms of Oxygen Activation.” Academic Press, New York, 1973. 15. T. Mann and D. Keilin, Proc. Roy. Soc., Ser. B 126, 303 (1938). 16. J. M. McCord and I. Fridovich, JBC 244, 6049 (1909). 17. According to the recommendations of the International Union of Pure and

Applied Chemistry (IUPAC) the name “hyperoxide” should be used to designate the 0; ion (18). The names “hyperoxide” and “hyperoxide dismutase” (19) have not, however, so far appeared in the literature reviewed. In order to avoid confusion the name “superoxide” has therefore been retained throughout this article. 18. “Nomenclature of Inorganic Chemistry,” 2nd ed. Butterworth, London, 1971. 19. Superoxide dismutase or superoxide :superoxide oxidoreductase, E C 1.15.1.1 (7). 20. R. C. Bray, Chapter 6, this volume. 21. I. Fridovich, in “Molecular Mechanisms of Oxygen Activation” (0. Hayaishi, ed.), p. 453. Academic Press, New York, 1973. 22. I. Fridovich, Annu. Rev. Biochem. 44, 147 (1975). 23. U. Weser, Struct. Bonding (Berlin) 17, 1 (1973).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

511

The emphasis in this chapter will be on the molecular properties of the enzymes, particularly on the relationship between structure and function, but brief presentations of the biological distribution and physiological functions will, in general, be given. None of the enzymes treated has had its three-dimensional structure determined by X-ray crystallography. In many cases, however, it has been possible to use copper as a built-in molecular probe of the active site, and much structural information has been derived from the application of spectroscopic methods, such as EPR. The organization of the chapter will to some extent not be around individual enzymes since it is often of great interest to compare the same property for a group of related enzymes. The blue oxidases, in particular, are treated in this manner since progress during the last decade has shown that all members of this group show great similarities in some respects (Section IV) . II. Enzymes Reducing Dioxygen to Hydrogen Peroxide

A. INTRODUCTION The copper-containing oxidases which utilize only half the oxidizing power of dioxygen (see Table I) are the amine oxidases (Section II,B) and galactose oxidase (Section I1,C). Hydrogen peroxide is also formed in the oxidation of urate by the pig liver enzyme uricase, which has been reported to contain copper (24). Uricases obtained from other sources, e.g., from bovine liver ( 6 5 ) , Candida utilis ( 2 6 ) , and Arthrobacter pascens (27,28), contain no copper, however, although the inhibition and pH dependence are similar to those of the pig liver enzyme. Since there appears to be no reports on copper in the pig liver enzyme except those earlier reviewed by Mahler (24), this enzyme will not be considered in this review (cf. Section I).

B. AMINE OXIDASES 1. Definition and Classification Amine oxidases are enzymes which catalyze the oxidative deamination of mono-, di-, and polyamines with the formation of stoichiometric amounts of aldehyde, hydrogen peroxide, and ammonia according t o the 24. 25. 26. 27. 28.

H. R. Mahler, “The Enzymes,” 2nd ed., Vol. 8, p. 285, 1963. R. Truscore and V. Williams. BBA 105, 292 (1965). K. Itaya, J. Fukumoto, and T . Yamamoto, Agr. B i d . Chem. 35,813 (1971). K. Arima and K . Nose, BBA 151, 54 (1968). K. Nose and K . Arima, BBA 151, 63 (1968).

512

B. G. MALMSTROM, L.-E. ANDB~ASSON,AND B. REINHAMMAR

following equation: RCHZNHZ

+ O*+

RCHO

+ H,Oa + N H I

(1) Equation (1) was established in 1938 by Zeller (as) and has been confirmed many times since. Methods for the measurements of the substrates and products in this reaction have been reviewed by Zeller ( S O ) . The amine oxidases are usually divided into two groups, mono- and diamine oxidases. This classification was initially based on the substrate specificities of the enzymes ( 3 1 ) .Blaschko and Duthie (32,33), however, reported later that some monoamine oxidases oxidize long-chain aliphatic diamines which are not degraded by diamine oxidases. These and many other observations on substrate and inhibitor reactions [for reviews, see Zeller ( 3 4 ) and Kapeller-Adler (36)]led to the classification used today by most workers in this field: Diamine oxidases, in contrast to monoamine oxidases, do not oxidize secondary amines and are not inhibited by the potent monoamine oxidase inhibitor 2-phenylcyclopropylamine. Monoamine oxidases are not inhibited by hydrazine and unsubstituted acylhydrazides while the diamine oxidases are strongly inhibited by hydrazine and semicarbazide (34). Much of the earlier work on amine oxidases has already been the subject of several review articles [see, for example, Zeller (34), KapellerAdler ( S b ) , and Blaschko ( 3 6 ) ] and reference to these articles will often be made instead of extensive quotations of old literature. Only a few amine oxidases seem to contain copper and only these enzymes will be dealt with in this article. With the exception of ii pig liver amine oxidase, which is not inhibited by semicarbazide, they are all diamine oxidases according to the classification given. Highly purified and extensively studied enzymes have been obtained from the fungus Aspergillus niger, pea seedlings, bovine blood plasma, pig plasma, and pig kidney cortex. The main part of this section will be devoted to these enzymes. Recently, a few other amine oxidases reported to contain copper have been prepared from pig liver and various connective tissues. Since 29. E. A. Zeller, Helv. Chim. Acta 21, 880 (1938). 30. E. A. Zeller, Advan. Enzymol. 2,93 (1942). 31. E. A. Zeller, R. Stern, and M. Wenk, Helv. Chim. Acta 23, 3 (1940). 32. H. Blaschko, Pharmacol. Rev. 4, 415 (1952). 33. H. Blaschko and R. Duthie, BJ 39, 478 (1945). 34. E. A. Zeller, “The Enzymes,” 2nd ed., Vol. 8, p. 313, 1963. 35. R. Kapeller-Adler, “Amine Oxidases and Methods for Their Study.” Wiley (Interscience), New York, 1970. 36. H. Blaschko, “The Enzymes,” 2nd ed., Vol. 8, p. 337, 1963.

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

513

the knowledge of these enzymes is still very limited, they will only get brief attention (Section II,B,8). The copper-containing amine oxidases have many properties in common. They are, however, distinctly different enzymes both with respect to molecular properties and substrate specificity. Therefore, each enzyme will first be treated separately (Section II,B,3) but in most sections a comparative presentation is used.

2. Metabolic Function The physiological function of the amine oxidases is the breakdown of a number of biologically active amines. Since diamines and polyamines are widely distributed in the living world, the diamine oxidases may act as regulators of the concentration of amines and, therefore, participate in a great number of biological processes (34-36). 3. Purification, Molecular Weight, and Substrate Specificity a. Aspergillus niger Amine Oxidase. In 1941, Werle (37), and later Roulet and Zeller (58), reported that some bacteria are able to catalyze the oxidative deamination of histamine, agmatine, aliphatic diamines, and spermidine. Since then many microorganisms have been found to produce diamine oxidases [for reviews, see Kapeller-Adler (35) and Yamada et al. ( 3 9 ) ] .Thus, in 1965, Yamada and his associates (40) detected amine oxidase in the mycelia of various fungi if these were grown with mono- and diamines as sole nitrogen sources. Later on, a copper-containing amine oxidase was prepared in high yield and purity from the mycelial extract of Aspergillus niger (41,42). The preparation method involves fractionations with ammonium sulfate, chromatography on DEAE-cellulose and DEAE-Sephadex, and, finally, crystallization from an ammonium sulfate solution. The enzyme appears fairly homogeneous in ultracentrifugation analysis. The molecular weight estimated according to the approach-to-equilibrium method is 252,000 (43),while sedimentation-diffusion analysis gives a value of 273,000 (4.2). If the enzyme is treated with guanidinium chloride containing mercaptoethanol it dissociates into subunits having a molecular weight of 85,000 (41). 37. E. Werle, Biochem. 2.309, 61 (19411. 38. F.Roulet and E. A. Zeller, Helu. Chim. Actu 28, 1326 (1945). 39. H.Yamada, H.Suzuki, and Y. Ogura, Advan. Biochem. Psychopharmacol. 5, 185 (1972). 40. H.Yamada, 0.Adachi, and K. Ogata, Agr. Biol. Chem. 29, 117 (1965). 41. 0.Adachi and H. Yamada, Agr. Biol. Chem. 33, 1707 (1969). 42. H.Yamada and 0. Adachi, “Methods in Enzymology,” Vol. 17B, p. 705, 1971. 43. H.Yamada, 0.Adachi, and K. Ogata, Agr. Biol. Chem. 29, 864 (1965).

514

B. G . MALMSTROM,

L.-E.

ANDREASSON, AND

B. REINHAMMAR

The enzyme oxidizes both mono- and diamines but not polyamines and secondary amines (43). The amines most rapidIy oxidized are aliphatic monoamines with chain lengths of C,-C,, phenethylamine, benzylamine, histamine, and agmatine. Tyramine, tryptamine, norepinephrine and serotonin are oxidized somewhat slower. The aliphatic diamines with a chain length of C,-C, are also oxidized but a t considerably lower rates. b. Pea Seedling Amine Oxidase. The studies of amine oxidase from pea seedlings go back to 1948 when Werle and his associates (44,45) reported that extracts of some leguminous plants are able to catalyze the oxidation of the diamines putrescine and cadaverine and also histamine. In 1955, Mann (46) reported a method for the preparation of an amine oxidase from pea seedling, and later a modified method which resulted in a highly purified enzyme was published (47). The preparation method consists of fractionating a crude extract from seedlings with a mixture of ethanol and chloroform, with ammonium sulfate and repeated precipitations a t pH 5. The product is then subjected to chromatography on columns of hydroxylapatite and DEAE-cellulose. A modification of this method has been reported by Werle et al. (48). Both methods give enzyme with high specific activity. The enzyme appears homogeneous a t low concentrations according to ultracentrifugation analysis. At high concentrations a small amount of faster sedimenting material was also detected (49). A molecular weight of 96,000 was estimated by electron microscopy (@). The minimum molecular weight obtained from the copper content is 53,000 (49) to 73,000 (@), however. The amino acid composition of electrophoretically pure enzyme has been determined (60).It shows an unusually high content of ornithine (5.8%) and, since no halfcystine was detected, disulfide bridges seem to be absent in the enzyme (see Table 11). Pea seedling amine oxidase catalyzes the oxidative deamination of mono-, di-, and polyamines. The diamines putrescine and cadaverine are most readily oxidized while aliphatic diamines with shorter chains are not oxidized. Other relatively good substrates are spermidine, agmatine, n-propyl- to heptylamine, benzylamine, tyramine, tryptamine, histamine, and L-lysine (49). c. Bovine Plasma Spermine Oxidase. Spermine oxidase was first found 44. 45. 46. 47. 48.

E. Werle and A. Raub, Biochem. 2.318,538 (1948). E. Werle and A. Zabel, Biochem. 2.318,554 (1948). P. J. G. Mann, BJ 59, 609 (1955). P. J. ,G.Mann, BJ 79, 623 (1961). E. Werle, I. Trautschold, and D. Aures, Happe-Seyler'e 2. Physiol. Ghem. 326,

200 (1961). 49. J. M. Hill and P. J. G . Mann, BJ 91, 171 (1964). 50. U.Nylkn and P. Ssybek, Actu Chem. Scund. B28, 1153 (1974).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

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TABLE I1 AMINOACID COMPOSITION OF THREE AMINE OXIDASES

Amino acid

Pea seedling amine oxidase"

Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Lyaine Histidine Arginine Amide ammonia Tryptophan Half-cystine Ornithine

17 9 27 24 22 11 8 1 7 8 4 5

7 5 A

Pig plasma benzylamine oxidaseb

Bovine plasma spermine oxidaseCsd

117 65 101 143 112 113 121 102 12 38 120 48 80 34 54 72 163 21 16

114 74 92 158 114 115 101 104 24 38 113 45 77 36 41 63 (162) 23 12

6

From NyEn and Szybek (60).

* From Blaschko and Buffoni (74). From Yamada et al. (67). Recalculated using a molecular weight of 170,000 (see 66). The reported figures are given as nearest integral values.

in the blood plasma of sheep by Hirsch (51) and subsequently Blaschko and his associates (56) reported the presence of this enzyme in the plasma of many ruminants. Partial purification of the bovine enzyme was first reported by Tabor e t al. ( 5 3 ) , and later highly purified crystalline enzyme was prepared by Yamada and Yasunobu ( 5 4 ) .The latter preparation method is based on fractionation with ammonium sulfate, column chromatography on DEAE-cellulose and hydroxylapatite, and, finally, crystallization in ammonium sulfate. The preparation is homogeneous according to electrophoretic analysis while results from ultracentrifuga51. 52. 53. 54.

J. G. Hirsch, J . Ezp. Med. 97, 435 (1953). H. Blaschko, Advan. Comp. Phgsiol. Biochem. 1, 67 (1962). C. W. Tabor, H. Tabor, and S. M. Rosenthal, JBC 208, 645 (1954). H. Yamada and K. T. Yasunobu, JBC 237, 1511 (1962).

516

B. G. MALMSTROM, L.-E. ANDREASSON, AND B. REINHAMMAR

tion studies show the presence of a lighter component which amounts to about 10% (64).The molecular weight, as determined by the methods of gel filtration and equilibrium centrifugation, is 170,000 (66). The enzyme molecule apparently consists of two identical subunits which are covalently bound. Dissociation into subunits with a molecular weight of 87,000 was accomplished when the enzyme was treated with 5 M guanidinium chloride which contained 0.1 M mercaptoethanol (66). It is a glycoprotein containing 4.6% carbohydrates (66). The amino acid composition of the crystalline enzyme has been determined (67) (see Table 11). The enzyme oxidizes some primary monoamines, diamines, and polyamines. Spermine and spermidine are most readily oxidized. Then benzylamine, heptylamine, amylamine, kynuramine, and butylamine are oxidized with decreasing rates. This enzyme does not oxidize tyramine, mescaline, norepinephrine, serotonin, and agmatine (63,64).Tabor et al. (68) have demonstrated that spermine is oxidized in both terminal positions with the formation of a dialdehyde while spermidine is converted to a monoaldehyde. Yamada et al. (69) have reported that also N,N‘bis (3-aminopropyl) -1,2-diaminoethane and N,N’-bis (3-aminopropyl) -1,6diaminohexane are oxidized. d. Pig Kidney Diamine Oxidase. The studies of pig kidney diamine oxidase go back to the 1930’s, and several workers have presented methods for partial purification from kidney cortex extracts [for a review, see Zeller (344)l.More recently] several methods for the preparation of apparently homogeneous enzyme have been reported (60-64). Molecular weight determinations with the sedimentation-diffusion technique have given values of 185,000 (61,66), 129,600, and 135,000 55. F. M. Achee, C. H. Chervenka, R. A. Smith, and K . T. Yasunobu, Biochemistry 7, 4329 (1968). 56. K. Watanabe and K. T. Yasunobu, JBC 245, 4612 (1970). 57. H. Yamada, P. Gee, M. Ebata, and K. T. Yasunobu, BBA 81, 165 (1964). 58. C. W.Tabor, H. Tabor, and U.Bachrach, JBC 239,2194 (1964). 59. H.Yamada, H.Kawasaki, T. Oki, I. Tomida, H. Fukami, and K. Ogata, Mem. Res. Inst. Food Sci., Kyoto Univ. 29, 11 (1968). 60. E. V. Goryachenkova, L. I. Scherbatyuk, and C. I. Zamaraev, in “Pyridoxal Catalysis: Enzymes and Model Systems” (E. E. Snell et al., eds.), p. 391. Wiley (Interscience), New York, 1968. 61. H.Yamada, H.Kumagai, H. Kawasaki, H. Matsui; and K. Ogata, BBRC 29, 723 (1967). 62. B. Mondovi, G.Rotilio, M. T. Costa, and A. Finazzi Agrb, “Methods in Enzymology,” Vol. 17,Part B, p. 735, 1971. 63. W.G. Bardsley, J. S. Ashford, and C. M. Hill, BJ 122, 557 (1971). 64. R.Kapeller-Adler and H. MacFarlane, BJ 82, 49P (1962). 65. J. M. Pionetti, BBRC 58, 495 (19x4).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

517

(66),and a value of 119,500 was obtained by the method of approach to equilibrium (66). The lower values are not a multiple of the minimum molecular weight of 85,000 (61) or 87,000 (66) obtained by copper analysis. It has been suggested that the protein molecule may undergo association-dissociation processes (66) and that the active unit has a molecular weight of 185,000 (66), Further association of the unit with the molecular weight of 185,000 to a tetramer of 725,000 has been observed (65). This process is dependent on the oxygen concentration. I n air-saturated solutions the monomer exists while the tetramer is formed a t low oxygen tensions. The substrate specificity has been extensively investigated. Early results with less pure enzyme preparations have been reviewed ( S 4 ) , and later investigations with more pure enzyme have given similar results with a few exceptions (61,63,67). The most rapidly oxidized amines are the alkyl diamines which have their amino groups separated by 6-9 A. Thus cadaverine and putrescine are oxidized fastest, and the rate of oxidation decreases when longer or shorter diamines are tested. Of the alkyl monoamines examined only propylamine and butylamine are oxidized a t appreciable rates. Other good substrates are histamine, agmatine, and p-bis(aminoethy1) benzene while the ortho and meta isomers are strong inhibitors. e. Pig Plasma Benzylamine Oxidase. In 1957, Blaschko and his associates (68) discovered an enzyme in horse serum which rapidly oxidizes benzylamine, and the name “benzylamine oxidase” was proposed for this enzyme. A benzylamine oxidase from pig plasma was described by Bergeret and Blaschko (69),and this enzyme was subsequently crystallized by Buffoni and Blaschko (YO) and by Taylor et at. (71).Both methods involve fractionations with ammonium sulfate, chromatography on DEAE-cellulose, DEAE-Sephadex, and hydroxylapatite, and, finally, crystallization with ammonium sulfate. A somewhat modified method has been reported by Lindstrom and Pettersson (72). The preparation is homogeneous according to starch gel electrophoresis at four different pH values and in ultracentrifugation analysis (70).Sedimentation-diff usion and approach-to-equilibrium analysis of the enzyme gives a molecular 66. B. Mondovi, G . Rotilio, M. T. Costa, A. Finazzi Agrb, E. Chiancone, R. E. Hansen, and H. Beinert, JBC 242, 1160 (1967). 67. W. G. Bardsley and C. M. Hill, BJ 117, 169 (1970). 68. B. Bergeret, H. Blaschko, and R. Hawes, Nature (London) 180, 1127 (1957). 69. B. Bergeret and H. Blaschko, Brit. J. Pharmacol. Chemother. 12, 513 (1957). 70. F. Buffoni and H. Blaschko, Proc. R o y . Soc, Ser. B 161, 153 (1964). 71. C. E. Taylor, R. S. Taylor, C. Rasmussen, and P. F. Knowles, BJ 130, 713 (1972). 72. A. Lindstriim and G. Pettersson, Eur. J . Biochem. 34, 564 (1973).

518

B. G . M A L M S T R ~ M , L.-E. ANDR~~ASSON,AND B. REINHAMMAR

weight of 196,000 (70) or 190,000 ( 7 3 ) . The amino acid composition has been determined, and the enzyme also contains about 10% carbohydrates (74) (see Table 11).In 6 M guanidinium chloride, containing 0.1 M mercaptoethanol, and in 1% sodium dodecyl sulfate, the enzyme is dissociated into subunits of molecular weight approximately 95,000 ( 7 3 ) . Benzylamine oxidase acts on many monoamines and also histamine but not on diamines and polyamines ( 7 5 ) .The most readily oxidized substance is benzylamine. After it come mescaline, 4-picolylamine1histamine, P-phenethylamine, tyramine, dopamine, tryptamine, cystamine, and serotonin. Kynuramine is also a good substrate. 4. Spectral Properties Highly purified preparations of amine oxidases in the oxidized state are pink or pinkish yellow in color. The color results from a broad absorption band with a maximum between 420 and 490 nm for the Aspergillus enzyme (39), at 505 nm for the pea seedling enzyme (49), between 470 and 500 nm in the pig kidney enzyme (60,61,66), and at 470 nm in the pig plasma benzylamine oxidase (70,76).The bovine plasma enzyme shows two absorption maxima in the visible region which are pH-dependent (77). Thus, a t pH 5.7 there is a maximum a t 480 nm, which is shifted to 410 nm at pH 9.8 with an isosbestic point at 466 nm. In some preparations of the Aspergillus enzyme (41,43) and the pig kidney enzyme (60,66) a shoulder at about 410 nm is also observed. This band is not reduced by amines and is thought to result from impurities, most likely heme compounds (60,63). The absorption bands disappear on reduction with amines or dithionite under anaerobic conditions and reappear on oxidation with oxygen (39,43,49,64,60,61,66,70).On the addition of various amines under anaerobic conditions new bands are formed a t 470 and 440 nm with the Aspergillus enzyme ( 3 9 ) , and a t 466, 437, and 350 nm with the pea seedling enzyme (49). These new bands have been attributed to an enzyme-amino group complex (39) or an enzyme-substrate complex ( 4 9 ) . With the pea seedling enzyme the formation of the 466-nm band requires the presence of copper ( 4 9 ) .When copper is removed the absorption maximum a t 505 nm is shifted to 480 nm in the pea seedling enzyme (49) while in the bovine plasma and the pig plasma enzymes the corresponding absorption bands disappear (75,77).Addition of various inhibi73. N. Boden, S. C. Charlton, M. C. Holmes, and P. F. Knowles, Biochem. SOC. Trans. 1, 1008 (1973). 74. H. Blaschko and F. Buffoni, Proc. R o y . Soc., Ser. B 163, 45 (1965). 75. F. Buffoni and L. Della Corte, Advan. Biochem. Psychopharmacol. 5, 133 (1972). 76. A. Lindstrom, B. Olsson, and G . Pettersson, Eur. J . Biochem. 35, 70 (1973). 77. H.Yamada and K. T. Yasunobu, JBC 238, 2669 (1963).

8.

COPPER-CONTAINING

OXIDASES AND S U P E R O X I D E DISMUTASE

519

tors also causes changes in the optical absorption spectrum; for example, phenylhydrazine, hydroxylamine, and hydrazine form derivatives with the native bovine plasma enzyme with maxima a t 447, 370, and 310 nm, respectively. The copper-free enzyme forms complexes which have maxima a t 410, 330, and 320 nm with these inhibitors (77). Similarly, addition of phenylhydrazine to the pea seedling enzyme (50),the AspergilZus enzyme ( S 9 ) ,and the pig plasma enzymes (72) results in the formation of strong absorption bands centered at 430-440 nm. 5. Prosthetic Groups

a. Metal Content. Copper is the only metal which has been detected in significant amounts in the amine oxidases. Thus, the Aspergillus enzyme has a copper content corresponding to three copper ions per protein molecule (78,79).The pea seedling enzyme contains 0.085-0.12% (48,49). These values correspond to one copper ion per 73,000-53,000 and should be compared to the estimated molecular weight of 96,000 (see Section II,B13,b). The bovine plasma enzyme has 1-2 copper ions per protein molecule ( 8 0 ) .The copper content of the pig kidney enzyme, as prepared by two different methods, corresponds to two copper ions per molecule (61,66), but a value of 3.3 copper ions has also been reported (60). With the pig plasma enzyme different laboratories report different copper contents. Buffoni and Blaschko (70) found 3 copper ions per protein molecule. Other laboratories reported 1.9-2.3 (73,81) or 2.5-2.7 (72) according to chemical determinations and integrations of EPR spectra. The copper ions in the amine oxidases are firmly bound but can be a t least partially removed by treating the enzymes with diethyldithiocarbamate (49,60,66,78,82-84) or acidic buffers (82). On removal of copper the enzymic activity is lost. Reactivation is obtained by the addition of suitable amounts of Cu2+(49,66,78,82-84) or Cu2+plus pyridoxal phosphate (60). Other metal ions do not reactivate the copper-free enzyme (49,82,83). At least part of the copper present in the amine oxidases is detected by EPR and thus present as Cu2+;for example, of the three copper ions 78. H. Yamada and K. T. Yasunobu, JBC 29, 912 (1965). 79. H. Yamada, 0. Adachi, and T. Yamano, BBA 191,751 (1969). 80. F. Achee, C. Chervenka, T. M. Wang, and K. T. Yasunobu, in “International Symposium of Pyridoxal Enzymes” (K. Yamada et at., eds.), p. 139. Maruzen, Tokyo, 1968. 81. F. Buffoni, L. Della Corte, and P. F. Knowles, BJ 106,575 (1968). 82. H. Yamada and K. T. Yasunobu, JBC 237, 3077 (1962). 83. H. Yamada, K. T. Yasunobu, H. Yamano, and H. S. Mason, Nature (London) 198, 1092 (1963). 84. E. Buffoni, Pharmacol. Rev. 18, 1163 (1966).

520

3. G. MALMSTROM,

L.-E. A N D R ~ S S O N , AND B. REINHAMMAR

in the Aspergillus enzyme, only about two are responsible for the signal (79) . With the bovine plasma enzyme about 70% of the total copper is detected by EPR (83). For the pig kidney enzyme two laboratories reported that between 78 and 100% of the copper ions are seen by this technique (60,SS). Electron paramagnetic resonanck indicates that Cuz+ is also present in the pea seedling enzyme ( 5 0 ) , but no quantitation of the signal has been made. Only in the case of the pig plasma enzyme does the amount of EPR-detectable copper correspond to the total copper present. Thus, Boden e t al. (73) and Buffoni et al. (81) found 1.9-2.3 copper ions, and Lindstrom and Pettersson (72) reported 2.5-2.7 copper ions in both cases according to chemical analysis as well as to integration of EPR spectra. The EPR parameters of amine oxidases (see Table 111) are quite similar and indicate that the Cu2+ coordination has tetragonal symmetry (66,73,79).The two Cuz+ions in the pig plasma enzyme are in different chemical environments as reported by Boden et at. (73), while the EPRdetectable copper ions in the pig kidney enzyme seem to be in equivalent sites (66). To characterize the ligand environment of the copper ions in the pig plasma enzyme further, Boden et al. (73) measured the proton relaxation enhancement caused by the protein. Their interpretation of these experiments is that water coordinated axially to the Cu2+ions is rapidly exchanging with water in the bulk aqueous phase. Electron paramagnetic resonance studies of the native and amine-reduced enzymes indicate that there is no significant reduction of the EPR signal intensity when amines are added under anaerobic conditions (50,76,79,81,83).However, for the pig kidney enzyme conflicting results have been reported. While in two cases only a small reduction of the TABLE I11 EPR PARAMETERS OF COPPER-CONTAINING AMINEOXIOASES Source

9max

QII

AII(G)

Ref.

Aspergillus niger

2.07 2.053 2.063 2.05 2.060 2.1

2.31

162 155 149 171 144 141

79 83 66

Bovine plasma Pig kidney Pig kidney Pig plasma Pea seedling

2.294 2.25 2.226 2.35

60 81 60"

The parameters were estimated from a published spectrum in Nyl6n and Szybek (60).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

521

EPR intensity was found (60,66), Mondovi et al. (85) reported that there were two types of changes in the E P R intensity on anaerobic addition of amines. First, there was an increase of about 20-30% of the EPR-detectable copper, followed after several minutes by a decrease of 2 5 3 0 % of the original copper signal. Whether these changes reflect oxidation and reduction of copper during the catalytic degradation of amines has, however, not yet been established. Although the addition of amines does not generally seem to reduce the EPR-detectable Cu2+in amine oxidases, small changes in the shape of the spectra have been reported (60,66,76,79,81) except for the bovine plasma enzyme ; for example, in the native pig kidney enzyme several superhyperfine lines with a splitting of about 14 G are observed in the g L region, indicating the involvement of nitrogen ligands in the binding of copper (66). Small changes in the superhyperfine pattern have been observed when substrates are added, and these changes depend on the substrate used. However, since the same superhyperfine pattern is obtained with [ 14N]putrescine as with [ 15N]putrescine, the possibility that the amine nitrogen of the substrate is a ligand to Cuz+ seems to be ruled out. On addition of dithionite under anaerobic conditions the EPR signal is reduced (81,83). The function of copper in amine oxidases has not yet been established. Since the metal is necessary for the activity, it has been suggested that copper is involved in the electron transfer from substrate-reduced protein to molecular oxygen [see, for example, Zeller (86)3. b. Pyridoxal Phosphate. The presence of pyridoxal phosphate in some amine oxidases was first suggested on the basis of several lines of evidence. Thus, the Aspergillus enzyme appears to contain two aldehyde groups according to hydrazine titrations (39) (see Section II,B,6) and a component which supports growth of Saccharomyces carlsbergensis and has fluorescence properties similar to those of pyridoxal phosphate was obtained from the hydrolyzed protein. Furthermore, a component with properties similar to those of pyridoxethylamine has been isolated from the enzyme after reaction with ethylamine followed by reduction with borohydride (41): On the basis of the phosphorous content and the spectral properties it has been suggested that the enzyme molecule contains two pyridoxal phosphates (77,87). The presence of pyridoxal phosphate in the pig kidney enzyme was reported by Kapeller-Adler and MacFarlane in 1963 (88). This sugges85. B. Mondovi, G. Rotilio, A. Finazzi Agrb, M. P. Vallogini, B. G. Malmstrom,

and E. Antonini, FEBS (Fed. Eur. Biochem. SOC.)Lett. 2,182 (1969). 86. E. A. Zeller, Advan. Biochem. Psychopharmacol. 5, 167 (1972). 87. H.Yamada and K. T. Yasunobu, BBRC 8,387 (1962). 88. R.Kapeller-Adler and H. MacFarlane, BBA 67, 542 (1963).

522

B. G. MALMSTROM,

L.-E. ANDR~ASSON, AND B. REINHAMMAR

tion was later supported by various experimental approaches. Thus, Mondovi et al. (89) reported that pronase hydrolysis of enzyme treated with semicarbazide and diethyldithiocarbamate liberates a substance which migrates as pyridoxal semicarbazone in paper chromatography. Furthermore, by alternately freezing and thawing the enzyme in the presence of phenylhydrazine and diethyldithiocarbamate, a substance which activates apoaspartate aminotransferase is liberated. Kumagai et al. (90) reduced the enzyme with sodium borohydride during reaction with [ 14C]histamine, After acid hydrolysis a substance which was identified as pyridoxal histamine was isolated. Acid hydrolysis also yields substance with the same electrophoretic mobility and fluorescence properties as pyridoxal phosphate, and this substance can reactivate apoaspartate transaminase (60). Each enzyme molecule appears to contain two pyridoxal phosphates. The pig plasma enzyme has also been reported to contain pyridoxal phosphate. Pronase digestion and acid hydrolysis of the crystalline enzyme yields a substance which shows optical absorption and fluorescence properties of pyridoxal (74,91),and this substance can reactivate apodecarboxylase ( 7 4 ) . On the basis of these results and the finding that the enzyme contains 4 moles of phosphate per mole of protein, it has been suggested that the enzyme contains three to four strongly bound pyridoxal phosphates ( 7 4 ) .However, by treating the enzyme with phenylhydrazine, catran, or cuprizone, Pettersson and his associates (72,96) found that there is only one inhibitor reactive group in the native or urea-denatured enzyme. If this group is blocked, the enzyme becomes inactive. Early results with the bovine plasma enzyme suggested the presence of one pyridoxal phosphate per enzyme molecule ( 7 7 ) .This is supported by the demonstration that the enzyme has about one hydrazine-reactive group, which is necessary for the activity (93).However, by isolation of phenylhydrazine derivatives from hydrolyzed inhibitor-treated enzyme, Watanabe et al. (94) found that the isolated compounds contain neither pyridoxal phosphate nor phosphate, and these authors therefore suggested that another cofactor might be present. With the pea seedling enzyme spectrophotometric titrations and the inhibitory effects on the enzyme of various hydrazines also indicate that 89. B. Mondovi, M. T. Costa, A,. Finazzi Agrb, and G. Rotilio, ABB 119, 373 (1967). 90.H. Kumagai, T. Nagate, H. Yamada, and H. Fukami, BBA 185,242 (1969). 91. E. H. Fischer, A. B. Kent, E. R. Snyder, and E. G. Krebs, JACS 80, 2906 (1958). 92. A. Lfndstr.om and G. Pettersson, Eur. J. Biochem. 48, 229 (1974). 93. J. E. H6cko-Haas and D. J. Reed, BBRC 39, 396 (1970). 94. K. Watanabe, R. A. Smith, M. Inamasu, and K. T. Yasunobu, Advan. Biochem. Psychopharmacol. 5, 107 (1972).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

523

there is one react,ive aldehyde group necessary for the activity (50,95). The nature of this aldehyde group is not known, but since the enzyme does not seem to contain pyridoxal phosphate (50,96), another cofactor may also be present in this case.

6. Inhibitor Reactions The inhibition of amine oxidases has been studied extensively. Out of hundreds of compounds tested as inhibitors, only a few will be mentioned here to give an idea of the inhibitor action. For a more detailed review the reader is referred to an article by Zeller (S4). All diamine oxidases are strongly inhibited by various carbonyl reagents like hydrazines, hydroxylamine, semicarbazide, aminoguanidine, isonicotinic acid hydrazine, and sodium cyanide (50,72,74,77,78,89,95,97). These agents are generally inhibiting a t to lo-' M concentrations. The inhibitors seem to form covalent complexes with carbonyl groups of pyridoxal phosphate in the enzyme having this cofactor, or of an unknown cofactor in the pea seedling and bovine plasma enzymes (see Section II,B,5,b) ; for example, the reaction between Aspergillus enzyme and phenylhydrazine, as proposed by Yamada et al. (39), is ,CHO E

'CHO

+

2@-NH-NH2

* E,

, CH=N-

NH-@

CH= N-NH-@

+ 2H20

(2)

in which equation +-NH-NH, is phenylhydrazine. Since 2 moles of phenylhydrazine must be added to inhibit the enzyme and to develop fully the new absorption band a t 442 nm, it appears that there are two carbonyl groups necessary for activity (39) (cf. Section II,B,5,b)'. I n some cases reversibility of inhibition has been demonstrated (34,93) ; for example, complexes between various hydrazines and the bovine plasma enzyme slowly decompose to active enzyme and noninhibitory products indicating that the inhibitors are altered by the reaction with the enzyme (93). The amine oxidases are also strongly inhibited by a number of metal chelating agents. The most used chelators are diethyldithiocarbamate, 1,lO-phenanthroline, 2,2'-bipyridyl, 8-hydroxyquinoline, cuprizone, CN-, and N,' (34,46,48,7.4,78,82,91,95,98,99) ; for example, micromolar concentrations of the first four compounds completely inhibit the pea seedling enzyme (98). Except for diethyldithiocarbamate, which is known to re95. E. F. Yamasaki, R. Swindell, and D. J. Reed, Biochemistry 9, 1206 (1970). 96. J. M. Hill, BJ 104, 1048 (1967). 97. W. G. Bardsley and C. M. Hill, BBRC 41, 1068 (1970). 98. J. M. Hill and P. J. G. Mann, BJ 85, 198 (1962). 99. W. G. Bardsley, R. E. Childs, and M. J. C. Crabbe, BJ 137, 61 (1974).

524

B. G. MALMSTR~M, L.-E. ANDRI~ASSON,AND B. REINHAMMAR

move copper in all these enzymes (see Section II,B,5,a), the inhibition was reversed upon dialysis or addition of various divalent metal ions. It was therefore sugested that these chelators inhibit by forming complexes with the enzyme-bound copper while only diethyldithiocarbamate actually removed the copper from the protein. The inhibitory action of cuprizone on the pig plasma enzyme has recently been investigated (96). This substance does not seem to inhibit by chelating the copper but rather by forming an irreversible complex with the pyridoxal phosphate in a similar way as the hydrazine compounds. 7. Interactions of the Substrates with the Active Site

Based on specificity results, Zeller and his associates (100) and Bardsley et al. ((i7,lOl) have proposed a scheme for the interaction between substrates and the active site in pig kidney amine oxidase. A recent analysis by Zeller is found in reference (86) and only a brief presentation will be given here. The scheme is as follows: D C B A NHz(CHz)s CHzNHz D' C' B' A'

Substrate Active site

a. Amino Group ( A ) and Pyridoxal Phosphate (A'). The group A must be a primary amine. Schiff-base formation between this amino group and A', which is the aldehyde group of pyridoxal phosphate, probably occurs, followed by an oxidation to aldehyde. The amino group A is to a large extent protonated and thus positively charged. Whether the group A' carries a negative charge or not is a matter of dispute. Bardsley et al. (67,101) suggested that there is a negative charge close to the prosthetic groups, pyridoxal phosphate and copper in the region of A', and that, there is an electrostatic interaction between A and A', but this idea is contested by Zeller (86). b. The a-Methglene Group. The amino group A must be attached to an unmodified a-methylene residue to permit degradation of amines by the pig kidney amine oxidase (100). A release of one of the a-hydrogens as a proton appears to be the rate-limiting step, and the mechanism for proton transfer seems to involve a base of the protein (66,86). c. Hydrophobic Interactions between Groups C and C'. Bardsley et al. (63) have reported that all substrates which are readily oxidized, are also characterized by low Michaelis constants. Consequently, good binding seems to affect the catalytic efficiency. In a calculation of the approx-

100. E. A. Zeller, J. R. Fouts, J. A. Carbon, J. C. Lazanas, and W. Yoegtli, Helv. Chim. Acta 39, 1632 (1966). 101. W. Bardsley, C. M. Hill, and R. W. Lobley, BJ 117, 169 (1970).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

525

imate binding energy between C and C’, Zeller (86) reported that about one-third to two-thirds of the total binding energy was derived from the interaction between C and C’. The hydrophobic binding site C’ is suggested to bind methylene groups more strongly than aromatic rings (101). d. Interaction between D and the Biding Site D’. The structure of D is not limited to an amino group. It can be replaced by imidazole and other heterocycles, by guanidine, dimethylamine, dimethyl sulfonium or isothiuronium residues (63,67,lOO). On the basis of these observations, Bardsley et al. (63) proposed that there is a negatively charged residue D’ surrounded by a hydrophobic region on the enzyme surface and that the binding between D and D’ is electrostatic. However, Zeller et al. (100) propose that D is uncharged and acts as a nucleophile toward an electron-accepting site (D’) which is suggested to be a carbonyl group. The groups A’ and D’ are suggested to be separated by 6-9 A since this corresponds to the internitrogen separation of the best substrates (101). For the interaction between substrates and the pea seedling enzyme, NylCn and Szybek (50) propose that for good binding the substrate amine to be oxidized must be uncharged, while the other amino group should be positively charged to bind to a carboxyl group of the enzyme.

8. Catalytic Mechanism Little is known about the mechanism of electron transfer from substrates to oxygen. The formation of a Schiff base between the amino group of the substrate and the activity-linked aldehyde groups is probably the first step in the reaction sequence. The available chemical and kinetic data indicate a ping-pong mechanism (102) for all these enzymes (39,50,71,75,82,95,96,l0%112). Except for the pig plasma enzyme, the 102. W.W.Cleland, “The Enzymes,” 3rd ed., Vol. 2, p. 1, 1970. 103. S. Oi, M. Inamasu, and K . T. Yasunobu, Biochemistry 9, 3378 (1970). 104. D. J. Reed and R. Swindell, Fed. Proc., Fed. Amer. SOC.Exp. Biol. 28, 891 (1969). 105. W. G.Bardsley and J. S. Ashford, BJ 128,253 (1972). 106. A. Finazzi Agrb, G. Rotilio, M. T. Costa, and B. Mondovi, FEBS ( F e d . Eur. Biochem. Soc.) Lett. 4, 31 (1969). 107. F. Buffoni, in “Pyridoxal Catalysis: Enzymes and Model Systems” (E. E. Snell et al., eds.), p. 363.Wiley (Interscience), New York, 1968. 108. F. Buffoni, L. Della Corte, and G. Ignesti, Pharmacol. Res. Commun. 4, 99 (1972). 109. A. Lindstrom, B. Olsson, G. Pettersson, and J. Szymanska, Eur. J . Biochem. 47, 99 (1974). 110. A. Lindstrom, B. Olsson, and G. Petterson, Eur. J. Biochem. 42, 177 (1974). 111. W. G. Bardsley, M. J. C. Crabbe, J. S. Shindler, and J. S. Ashford, BJ 127, 875 (1972). 112. W.G. Bardsley, M. J. C. Crabbe, and J. Shindler, BJ 131, 459 (1973).

526

B. G. MALMSTROM, L.-E. ANDR~ASSON, AND B. REINHAMMAR

first product formed in the reaction of amines in the absence of oxygen is an aldehyde. Thus, 2 moles of aldehyde per moIe of enzyme are formed with the Aspergillus enzyme (SQ),while about 1 mole of aldehyde is formed with the pea seedling ( 9 5 ) , the bovine plasma (103,104), and the pig kidney enzymes (106). After oxygen is introduced, hydrogen peroxide and ammonia are released. Which of these products is released first has been studied by Bardsley and Ashford (105) with the pig kidney enzyme. They found that ammonia gave competitive inhibition with diamine, which was taken as evidence that ammonia was the last product to be liberated. Furthermore, hydrogen peroxide gives uncompetitive inhibition with diamine indicating that it is released after the second substrate has been added. On the basis of these observations they propose the following reaction scheme for this enzyme: P

A

E

(EA) (FP)

Q T

B

t

1

1 F

(FB) (EQW

R

t ER

E

In this, E and F are two enzyme forms, A is a diamine, B oxygen, P aminoaldehyde, Q hydrogen peroxide, and R ammonia. For the bovine plasma enzyme Oi et al. (103) propose a random release of ammonia and hydrogen peroxide. In the scheme proposed by Taylor et al. (71) the first product formed is ammonia and not aldehyde as with the other amine oxidases. The detection of ammonia but not aldehyde under highly anaerobic conditions supports this order. According to this scheme, aldehyde is the last product released and, although aldehyde is split off before oxygen reoxidiaes the enzyme, it is bound to a hydrophobic site in the protein and not released until ammonia and hydrogen peroxide are released. However, Pettersson and his associates (110) report that, in the presence of 0.01-0.25 mM oxygen, 1 mole of aldehyde is formed in a first-order burst in stopped-flow experiments. The aldehyde is released prior to the rate-limiting interaction between oxygen and the reduced form of the enzyme. As suggested by them, these two conflicting results might depend on the difference in the oxygen concentration used in the two different experiments, Taylor et al. using 0.01 p M oxygen (71). 9. Other Amine Oxidases T h a t M a y Contain Copper a. Pig Liver Monoamine Oxidase. Monoamine oxidase from pig liver

has been purified about 200-fold according to the increase in specific activity (113).It was reported to have a molecular weight of 1,200,000 and to contain eight subunits, with a molecular weight of 146,000, and 113. W. R. Carper, D. D. Stoddard, and D. F. Martin, BBA 334, 287 (1974).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

527

eight copper ions. By dialysis against diethyldithiocarbamate the activity was lost and partial reactivation was obtained by the addition of Cu2+, Coz+, Znz+, and Ni2+. That other metals can replace Cuz+ is not found for the other copper-containing oxidases (see Section II,B,5,a). The enzyme also differs from the other amine oxidases in its inhibitor reactions. Thus, it is not inhibited by semicarbazide and sodium cyanide. However, several chelators, e.g., neocuproine, o-phenanthroline, diethyldithiocarbamate, and N3-,are relatively good inhibitors. b. Connective Tissue Ainine Oxidase. Amine oxidases have also been prepared from various connective tissues. Thus, Nakano et a2. ( 1 1 4 ) prepared an oxidase from bovine dental pulp, and Rucker et al. (115,116) isolated one from bone tissue. Amine oxidases have also been observed in bovine and chick aorta (117,118). These enzymes have only been partially purified and have not been well-characterized. In most cases a copper analysis has not been made, but the chick aorta enzyme is reported to contain 1 g-atom of copper per 45,000 g of protein (118) Metal ions and pyridoxal phosphate, however, seem t o be necessary for the activity of the other connective tissue oxidases (114-117). The enzymes are inhibited by carbonyl reagents, e.g., phenylhydrazine, hydroxylamine and semicarbazide (116,118), isoniazide (114,116,117), and chelating agents (116,118). The properties of these amine oxidases seem to be similar to the bovine plasma amine oxidase (86). They deaminate benzylamine and polyamines such as spermine and spermidine. The amine oxidases in the aorta also deaminate peptidyllysine in lysine-vasopressin (116), indicating a possible deamination of peptidyllysine in collagen or elastin to produce cross-linking reactions.

C. GALACTOSE OXIDASE 1. Discovery and Purification Galactose oxidase was discovered in 1959 by Cooper et a2. (119) in the extracellular culture medium of the fungus Polyporus circinatus (120). 114. C. Nakano, M. Hasada, and T . Nagatsu, BBA 341,366 (1974). 115, R. B. Rucker, J. C . Rogler, and H . E. Parker, Proc. SOC.Exp. Biol. M e d . 130, 1150 (1969). 116. R. B. Rucker and B. L. O’Dell, BBA 235,32 (1971). 117. R. B. Rucker and B. L. O’Dell, Fed. Proc., Fed. Amer. SOC. Exp. Biol. 29, 668 (1970). 118. E. D. Harris, W. A. Gonnerma, J. E. Savage, and B. L. O’Dell, BBA 341, 332 (1974). 119. J. A. D. Cooper, W. Smith, M. Bacila, and H. Medina, JBC 234,445 (1959). 120. Some confusion remains whether the organism producing this enzyme is Polyporus circinatus or Dactylium dendroides (121).

528

B. G. MALMSTROM, L.-E.

ANDREASSON, AND

B. REINHAMMAR

It catalyzes the oxidation of galactosides a t the C-6 position (122), as shown in Eq. (3). In subsequent publications, Horecker and co-workers CH,OH

HOQ

H,OR

+

0 , -

OH

H

O

O H,OR

+

H,O,

(3)

OH

(1%-124) reported methods for the cultivation of the fungus and the preparation of crystalline enzyme. The preparation method (121) involves ammonium sulfate fractionations of the culture filtrate in the presence of powdered cellulose, chromatography on DEAE-cellulose, another ammonium sulfate precipitation step, and, finally, crystallization from an ammonium sulfate solution. Other available purification methods (126,126) involve only slight modifications of this procedure. The crystalline enzyme appears homogeneous in gradient electrophoresis, sucrose gradient centrifugations (121),and in equilibrium centrifugation (124).

2. Chemical and Physical Properties

The molecular weight of galactose oxidase, as determined by equilibrium centrifugation, is 42,400 f 4,000 (124). Values of 50,000 (126), 55,000 (l27),and 68,000 (128) have, however, also been reported. The amino acid composition has been determined and the number of residues per molecule is given in Table IV (124).There appear to be three disulfide bridges and one thiol group in the enzyme molecule. Reduction with p-mercaptoethanol leads to the rupture of one disulfide bond and complete loss of enzymic activity. The reduced protein can be oxidized by air with complete restoration of activity. The enzyme remains fully active after treatment for 1 hour in 8 M urea (124))and the native enzyme is very stable a t room temperature (123). 121. D. Amaral, F. Kelly-Falcos, and B. L. Horecker, “Methods in Enzymology,” Vol. 9, p. 87, 1966. 122. G. Avigad, D. Amaral, C . Arsenio, and B. L. Horecker, JBC 237, 2736 (1962). 123. D. Amaral, L. Bernstein, D. Morse, and B. L. Horecker, JBC 238, 2281 (1963). 124. F. Kelly-Falcor, H. Greenberg, and B. L. Horecker, JBC 240, 2966 (1965). 125. S. Bauer, G. Blauer, and G. Avigad, Isr. J. Chem. 5, 126p (1967). 126. G. A. Hamilton, J. DeJersey, and P. K. Adolf, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), p. 103. Univ. Park Press, Baltimore, Maryland, 1973. 127. G. A. Hamilton, R. D. Libby, and R. C. Hartsell, BBRC 55, 333 (1973). 128. R. S. Giordano and R. D. Bereman, JACS 98, 1019 (1974).

8.

COPPER-CONTAINING

OXIDASES AND SUPEROXIDE DISMUTASE

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TABLE I V AMINOACID COMPOSITION OF GALACTOSE OXIDASEO-~ Amino acid

No. of residuesc

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

27 8 22 6 42 28 33 29 23 55 34 18 8 13 19

5 14

From Kelly-Falcoa (124). The number of residues in moles per mole of enzyme was calculated assuming a molecular weight of 47,000. The reported values have been converted to the nearest integral number.

3. T h e Copper of Galactose Oxidase The only nonprotein constituent of galactose oxidase appears to be copper. The copper content determined in different laboratories corresponds to one atom per molecule on the basis of molecular weights of 40,000-48,000 (121), 55,000 ( l 2 7 ) ,60,000 ( 1 2 6 ) ,or 68,000 (128).This copper ion appears necessary for the enzymic activity (123,124). It is firmly bound and cannot be removed by dialysis against NaCN, EDTA, or Chelex 100 (123).Dialysis against diethyldithiocarbamate, isoamyl alcohol, or H,S, on the other hand, removes the copper with a concomitant loss of activity (l,%?,lZ4). The apoenzyme is stable even a t room temperature (121). Restoration of the native enzyme can be obtained by the addition of either Cu+ or Cu2+.Other metals tested are completely inactive (123,124). At least part of the copper in the native protein is in the Cu2+state according to integrations of EPR spectra. Thus, in the first EPR analysis of this enzyme, Blumberg et al. (129) found that about 70% of the total copper is detected by EPR. Hamilton et d. 129. W. E. Blumberg, B. L. Horecker, F. Kelly-Falcoz, and J. Peisach, BBA 96, 336 (1965).

530

B. G. MALMSTROM,

L.-E. ANDRI~ASSON, AND B. REINHAMMAR

TABLE V ELECTRON PARAMAGNETIC RESONANCE PARAMETERS FOR GALACTOSE OXIDASEO

2.058

2.048 2.04

4

2.273 2.28

28.8

30.1

176.5 175.5

130 189

Absolute values of the hyperfine couplings are given in gauss.

(197) also report that the resting enzyme usually contains 75% Cu2+. The appearance of the E P R spectrum indicates that the Cu2+ ion is in a pseudo-square planar environment (130) with the estimated parameters presented in Table V. The addition of galactose under aerobic or anaerobic conditions does not lead to any changes in the shape or intensity of the E P R spectrum (12?',129,130). Furthermore, the presence of 0.3 M galactohexodialdose, the reaction product of galactose (122),or catalase and superoxide dismutase, has no effect on the EPR spectrum (130). On the other hand, Hamilton et al. (127) have reported that there is a 20-30% increase in the E P R intensity and an inhibition of the enzyme-catalyzed reaction when superoxide dismutase is added to the reaction medium. They also found that addition of ferricyanide or superoxide ion resulted in a marked decrease in the EPR signal and a great increase in activity. Various other additions have also been found to induce changes in the E P R spectrum ( 1 9 1 ) ; for instance, stoichiometric amounts of CN- or SCN- cause changes in the EPR parameters, and higher concentrations of the ligands cause no further alterations in the spectra. On the other hand, N3-, F-, Br- and H,02 give full changes only when added in molar excess. From these observations i t has been inferred that only a single coordination site of the copper ion is readily accessible to these ligands.

4. Optical Properties Galactose oxidase is colorless even in concentrations around 0.1 mM (121).Circular dichroism (CD) spectra of the native enzyme show bands at 314, 400, 485, and 600 nm (1.92). The addition of galactose under 130. D. J. Kosman, R. D. Bereman, M. J. Ettinger, and R. S. Giordano, BBRC 54, 856 (1973). 131. R. S. Giordano, R. D. Bereman, D. J. Kosman, and M. J. Ettinger, JACS 96, 1023 (1974). 132. M. J. Ettinger and D. M. Kosman, Fed. Proc., (Fed. Amer. SOC.Ezp. Biol.) 32, 543 Abstr. (1973).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

531

anaerobic conditions results in a 50% decrease in the 314- and 600-nm bands. 5. Inhibitor Reactions NaCN in a concentration of 0.1 miM completely inhibits galactose oxidase (123,133).Since reactivation is obtained by the addition of Ni2+, CO", or Ag+, it was suggested that CN- complexes with, but does not remove, the copper ion (123).Complete inhibition is also caused by 0.1-1 mM concentration of N3-, formate, hydrazine, and hydroxylamine (123,133).The enzyme is furthermore inhibited by H,Oz (119, 126,134), C1-, phosphate (126,133), acetate, ethylmorpholine, and Tris (196). 6. Specificity Galactose oxidase catalyzes the oxidation of D-galactose and a number of substances related to it (161,1%,1~6).Thus, the monosaccharides 1, 5-anhydrogalactitol, 2-deoxy-~-galactose,n-talose, D-galactosamine, and N-acetyl-D-galactosamine are readily oxidized. The galactose configuration a t position 4 is essential; for example, the enzyme does not oxidize D-fructose, D-glucose, or D-mannose. Furthermore, derivatives of D-galactose having substituents on the hydroxyl group a t C-4 are not oxidized. Although 2-amino-2-deoxy-~-galactose is a relatively good substrate, derivatives of this substance having glycosyl substituents a t C-3 are not oxidized. The C-1 position does not need to be free since the galactosides methyl-2-~-galactopyranoside, methyl-p-D-galactopyranoside, melibiose, and lactose are oxidized. Oligosaccharides and polysaccharides containing galactose, for instance, raffinose, stachyose, planteose and guran, are among the best substrates. The much simpler molecule, dihydroxyacetone, has also been reported to be a good substrate (134). Galactose oxidase has been used in a number of analytical estimations of galactose in biological samples and in mixtures of carbohydrates as well as in structural studies of galactose-containing polysaccharides (121,133,136,136). The extent of oxidation in these analyses has in most cases been estimated by a method in which H202produced in the reaction is coupled, through peroxidase, to a chromogen (121,133,135). Manometric (119) and polarographic (IS?+)techniques have also been used for measurements of the rate of 0, consumption during reaction. 133. R. A. Schlegel, C. M. Gerbeck, and R. Montgomery, Carbohyd. Res. 7, 193 (1968). 134. G. A. Hamilton, Advan. Enzymol. 32, 55 (1969). 135. G. G. Guilbault, P. J. Brignac, and M. Juneau, Anal. Chem. 40, 1256 (1968). 136. S. M. Rosen, M. J. Osborne, and B. L. Horecker, JBC 239, 3196 (1964). 137. G. T. Zancan and D. Amaral, BBA 198,146 (1970).

532

B. G . MALMSTROM, L.-E. ANDR~ASSON, AND B. REINHAMMAR

7 . Mechanism Only very limited data concerning the catalytic mechanism of galactose oxidase are available. Hamilton et al. (126') have suggested that the reaction follows a ping-pong mechanism. Ettinger and his associates (132,138) have also formulated a sequential mechanism on the basis of changes in the CD spectrum on addition of galactose and on kinetic data. Recently, Hamilton et al. (127) have proposed a scheme for the involvement of the copper in a redox cycle during catalysis. They suggested that in the active, oxidized protein the copper is tervalent and becomes monovalent during the oxidation of substrate. During reoxidation by oxygen Cut is first converted to C u z t ~ 0 2and - then further oxidized to Cu3+with the formation of H,O,. They also suggested that 0,- sometimes dissociates from the CuZf.O,- intermediate to give a Cu2+-enzyme which is catalytically inactive, thus accounting for the strong EPR signal observed in the resting enzyme (see Section II,C,3). In the presence of superoxide dismutase the liberated 0,- would be removed, which would explain the inhibitory action of this enzyme. An important basis for the proposed scheme was the observation that on the addition of 0,- or ferricyanide there is a large increase in the catalytic activity concomitant with a decrease in the EPR intensity. Here, 0,- was thought to drive the equilibrium in Eq. (4) to the right while the role of ferricyanide would be to oxidize Cu2+to Cus+: 02-

+ cu2+ s CU'+. 02-

(4)

Kwiatkowski and Kosman (139) have also observed strong inhibition of the catalytic rate when superoxide dismutase is incubated with galactose oxidase prior to the addition of the substrate mixture. However, the addition of even greater amounts of the dismutase after the reaction has begun has no effect on the reaction rate. Furthermore, the inactivation caused by superoxide dismutase is abolished if peroxidase or bovine serum albumin is present in either the enzyme or the substrate solution. These observations cast some doubts on at least part of the experimental basis for the proposed detailed mechanism of electron transport (127).I n some respects the proposal raises more questions than it answers; for example, Cu3+has never been observed in an enzyme and would not be expected to be a stable species. Furthermore, the oxidation-reduction potential of the Cuz+/Cu+couple must' be unusually low. Since the potential of the 02/0,-couple is about -0.3 V ( I @ ) , the scheme of Hamilton e t al. (127) 138. D. J. Kosman, L. Kwiatkowski, M. J. Ettinger, and J. D. Broide, Fed. Proc., Fed. Amer. SOC.Exp. B i d . 32, 550 Abstr. (1973). 139. L. D. Kwiatkowski and D. J. Kosman, BBRC 53,715 (1973). 140. P.Wood, FEBS (Fed Eur. Biochem. Soc.) Lett. 44,22 (1974).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

533

requires a similar value for the copper couple. Such a low oxidation-reduction potential has never been found for any other protein-bound Cu2+. Obviously, further experiments are necessary to establish whether the copper actually changes its valence state during catalysis. 111. Superoxide Dismutase

A. INTRODUCTION 1. A Brief Historical Review

The discovery by Mann and Keilin of the copper-containing proteins hemocuprein and hepatocuprein in bovine erythrocytes and liver (15) was followed by the isolation of a similar protein in human brain tissue (141). This was consequently named “cerebrocuprein.” Immunological investigations (14S,14S) led to the suspicion that the three cuproproteins were identical but the limited purity of the investigated material made de,finitive conclusions impossible. Eventually, Carrico and Deutsch (144) could demonstrate the identity of human erythrocuprein, hepatocuprein, and cerebrocuprein by conducting extensive studies of the physical, chemical, and immunological properties of the highly purified proteins. They suggested that the metalloprotein should be given the name cytocuprein. Furthermore, they were able to show that zinc is firmly bound t o the protein in addition to copper (145).I n 1965, McCord and Fridovich (16) purified a protein from bovine blood with superoxide dismutase activity which was found to be identical with erythrocuprein. Copper-zinc proteins with superoxide dismutase activity have now been isolated from cells of a wide variety of eukaryotic organisms so that it is reasonable to assume that they have a vital function in oxygen metabolism. A detailed historical account of the discovery of superoxide dismutase is given by Fridovich ( 2 1 ) .

2. T h e Physiological Role of Superoxide Dismutase a. Dismutation of Superoxide. A number of biological reactions in aerobic organisms have been proposed to involve the generation of superoxide, for example, reactions involving nonheme iron proteins (146-150), rubredoxin- or NADPH-dependent oxidation of epinephrine (151,152), or 141. H. Porter and J. Folch, A M A Arch. Neurol. Psychiat. 77, 8 (1950). 142. G. S. Shields, H. Markowitz, W. H. Klassen, G. E. Cartwright, and M. M. Wintrobe, J. Clin. Invest. 40, 2007 (1961). 143. M. J. Stansell and H. F. Deutsch, JBC 241,2509 (1966). 144. R. J. Carrico and H. F. Deutsch, JBC 244, 6087 (1969). 145. R. J. Carrico and H. F. Deutsch, JBC 245, 723 (1970).

534

B. G. MALMSTR~M, L.-E.

ANDREASSON,

AND B. REINHAMMAR

the autoxidation of epinephrine (153).Some reactions involving flavoproteins are other sources of superoxide (164,155).The molybdenum-containing flavoenzyme, xanthine oxidase, generates superoxide as a product in catalysis (156)which has been demonstrated by the use of rapid-freeze EPR (157).Evidence has also been found that superoxide is formed during autoxidation of hemoglobin (168-160). The 0,- radical is potentially hazardous to living matter. There are indications that 0,- reacts with thiol groups and with tryptophan residues (16'1,162), reactions which may well be lethal. Lavelle et ul. (163) have reported that 0,- produced by photooxidation of reduced flavin or in the xanthine oxidase reaction kills bacteria and renders viruses noninfective (and presumably induces mutations). In addition, ribonuclease is inactivated. Thus, since there is evidence that superoxide is produced by living organisms, these must be expected to have developed means to dispose of this dkgerous species. Evidence for a protective action of superoxide dismutase has been found in a number of cases. There seems to exist a clear correlation in microorganisms between the intracellular concentration of superoxide on the one hand and oxygen tolerance and survival rate on the other (163-168). I n many cases the protective effect of superoxide dismutase 146. P. Handler, K. V. Rajagopalan, and V. Aleman, Fed. Proc., Fed. Amer. SOC. Ezp. Biol. 23, 30 (1964). 147. R. C. Bray, G. Palmer, and H. Beinert, JBC 239, 2667 (1964). 148. H. P. Misra and I. Fridovich, JBC 246,6886 (1971). 149. H. P. Misra and I. Fridovich, JBC 247, 188 (1972). 150. W. H. OrmeJohnson and H. Beinert, BBRC 38,905 (1969). 151. S. D. Aust, D. L. Roerig, and T. C. Peterson, BBRC 47, 1133 (1972). 152. S. W. May, B. J. Abbott, and A. Felix, BBRC 54, 1540 (1973). 153. H. P. Misra and I. Fridovich, JBC 247, 3170 (1972). 154. R. P. Kumar, S. D. Ravindranath, C. S. Vaidyanathan, and N. A. Rao, BBRC 48, 1049 (1972). 155. V. Massey, S. Strickland, S. G. Mayhew, L. G. Howell, P. C. Engel, R. G. Mabthews, M. Schuman, and P. A. Sullivan, BBRC 38, 891 (1969). 156. J. M. McCord and I. Fridovich, JBC 243,6753 (1968). 157. P. F. Knowles, J. F. Gibson, F. M. Pick, and R. C. Bray, BJ 111, 53 (1969). 158. H. P. Misra and I. Fridovich, JBC 247, G960 (1972). 159. R. Wever, B. Ondega, and B. F. van Gelder, BBA 302, 475 (1973). 160. W. J. Wallace, J. C. Maxwell, and W. S. Caughey, BBRC 57, 1104 (1974). 161. A. M. Michelson, Biochimie 55, 465 (1973). 162. A. R. Green and G. Curzon, Nature (London) 220, 1095 (1968). 163. F. Lavelle, A. M. Michelson, and L. Dimitrijevic, BBRC 55, 350 (1973). 164. E. M. Gregory, F. J. Yost, Jr., and I. Fridovich, J. Bactem'ol. 115,987 (1973). 165. E. M. Gregory and I. Fridovich, J . Bacteriol. 114, 643 (1973). 166. E. M. Gregory, S. A. Goscin, and I. Fridovich, J. Bacteriol. 117, 456 (1974). 167. E. M. Gregory and I. Fridovich, J. Bacteriol. 114, 1193 (1973). 168. F. J. Yost, Jr., and I. Fridovich, ABB 161,395 (1974).

8.

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OXIDASES AND SUPEROXIDE DISMUTASE

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is enhanced by the presence of catalase, and in aerobic microorganisms both enzymes are present in most cases (169).There exist a few aerobic bacteria which contain only superoxide dismutase but lack catalase completely ( 17O,171), which indicates that superoxide is more toxic than hydrogen peroxide. An explanation of the effect of catalase in improving the protection against superoxide would be that it prevents the production of reactive hydroxyl radicals which may otherwise be formed via the reaction :

+

0%- Hz02

+ H+

+

HO'

+ HZ0 + 02

(5)

It has recently been demonstrated that superoxide dismutase is localized to those parts of rat liver cells where production of superoxide is known to take place, for example, the cytosol (172).Evidence has been found showing that superoxide dismutase protects membrane systems by preventing peroxidative degradation of lipids (173,174). Also, degradative autoxidation of lysine-tRNA ligase (a lipoprotein complex) is inhibited by superoxide dismutase (163).The presence of catalase in these systems enhances protection. The ability of superoxide dismutase to scavenge 0,- radicals has been used to probe the mechanism of a number of chemical reactions, e.g., sulfite oxidation (175),ethylene production in fruit ( l 7 6 ) , and nonenzymic hydroxylation (177). b. Scavenging of Singlet 0,. While the ability of superoxide dismutase rapidly to disproportionate 0,- radicals appears to have been unambiguously demonstrated there exists no general agreement concerning the true biological function of the enzyme. It has been suggested that the main function would be to scavenge singlet oxygen rather than the disproportianation of superoxide radicals (178,179). Singlet oxygen (l&+ or 'A,) can be expected to be deleterious in biological systems, mainly by causing 169. J. M. McCord, B. B. Keele, Jr., and I. Fridovich, Proc. N u t . Acud. Sci. U S . 68, 1027 (lfll). 170. R.N. Costilov and B. B. Keele, Jr., J . Bacterial. 11,628 (1972). 171. A. A. Yousten, L. A. Bulla, and J. M. McCord, J. Bucteriol. 113, 524 (1973). 172. G.Rotilio, L.Calabrese, A.. Finazzi Agrb, M. P. Argento-Ceru, F. Autuorio, and €3. Mondovi, BBA 321,98 (1973). 173. J. A. Fee and H. D. Teitelbaum, BBRC 49, 150 (1972). 174. R. Zimmermann, L. FlohC, U. Weser, and H. J. Hartmann, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 29, 117 (1973). 175. J. M. McCord and I. Fridovich, JBC 244, GO56 (1969). 176. C. 0.Beauchamp and I. Fridovich, JBC 245, 4641 (1970). 177. S. A. Goscin and I. Fridovich, ABB 153, 778 (1973). 178. A. Finazzi Agrit, C. Giovagnoli, P. Del Sole, L. Calabrese, G. Rotilio, and B. Mondovi, FEBS (Fed. Eur. Bwchem. Soc.) L e t t . 21, 183 (1972). 179. W.Paschen and U.Weser, BBA 327,217 (1973).

536

B. G. MALMSTBOM,

L.-E. A N D R ~ S S O N ,AND B. REINHAMMAR

oxygenation of unsaturated compounds (180-182). Kahn (183) pointed out a few years ago that the spontaneous dismutation of 02gives rise to the singlet state (lz0+O2)which is excited relakive to the triplet ground state. The singlet state can be revealed by its ability to induce chemiluminiscence in certain organic compounds (184). I n several cases superoxide dismutase has been observed to quench chemiluminescence generated in systems where singlet oxygen of either type is believed to be or actually has been shown to be formed (178,179,186-189). Paschen and Weser (179). showed that the quenching of lho oxygen is dependent on the enzyme concentration and much more specific to the enzyme than the dismutating ability. Thus, model copper chelates were found to be almost inactive in quenching luminiscence whereas they have been shown to have a significant superoxide dismutase activity (190). Contradictory to these results, Goda et al. (191) found that superoxide dismutase did in fact enhance the chemiluminescence produced from decaying singlet oxygen (see 192) when l-phospho-2,8,9-trioxadamantaneozonide was used as a singlet oxygen source (193). Also, the enzyme did not affect the reaction of singlet oxygen with a-lipoic acid and 9,lO-diphenylanthracene-2,3-dicarboxylic acid, which led Goda et al. to suggest that superoxide dismutase does not quench singlet oxygen. In this context it should be pointed out that chemiluminescence may result from the direct action of superoxide on certain organic compounds, for example, luminol (194-197), which has often been used in investigations of the reaction 180. C. S. Foote, S. Wexler, W. Ando, and R. Higgins, JACS 90, 975 (1968). 181. D. R. Kearns, R. A. Hollins, A. U. Kahn, R. W. Chambers, and P. Radlick, JACS 89, 5455 (1968). 182. D. R. Kearns, R. A. Hollins, A. U. Knhn, R. W. Chambers, and P. Radlick, JACS 89, 5456 (1968). 183. A. U. Kahn, Science 168, 476 (1970). 184. A. U. Kahn and M. Kasha, JACS 88, 1574 (1966). 185. R. M. Arneson, ABB 136,352 (1970). 186. U. Weser and W. Paschen, FEBS (Fed. Eur. Biochem. Soc.) Lett. 27, 248 (1972). 187. H. W. S. Chan, JACS 93, 2357 (1971). 188. A. Finazzi Agrb, L. Avigliano, G. A. Veldink, J. F. G. Vliegenthart, and J. Boldingh, BBA 326, 462 (1973). 189. 0. M. M. Faria Oliviera, D. L. Sanioto, and G. Cilento, BBRC 58, 391 (1974). 190. K. E. Joester, G. Jung, U. Weber, and U. Weser, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 25, 25 (1972). 191. K. Goda, T. Kimura, A. L. Thayer, K. Kees, and A. P. Schaap, BBRC 58, 660 (1974). 192. P. D. Markel and D. R. Keams, JACS 94, 7244 (1972). 193. A. P. Schaap, K. Kees, and A. L. Thayer, Abstr., 6th Cent. Reg. Meet., Amer. Chem. SOC.(1974). 194. J. R. Totter, E. C. d e Dugros, and C. Riveito, JBC 235, 1839 (1960). 195. L. Greenlee, I. Fridovich, and P. Handler, Biochemistry 1, 779 (1962). 196. K. D. Legg and D. M. Hercules, JACS 91, 1902 (1969). 197. E. K. Hodgson and I. Fridovich, Photochem. Photobiol. 18, 451 (1973).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

537

between superoxide dismutase and singlet oxygen. Therefore, observations of luminescence quenching by superoxide dismutase should be interpreted with caution.

3. Superoxide Dismutase fromProkaryotes and Mitochondria Bacteria have been found to contain a superoxide dismutase that differs from the animal enzyme in that it contains neither copper nor zinc but rather one atom of manganese per molecule (198-201). It also differs from the copper-zinc enzyme with respect to the molecular weight, amino acid composition, amino acid sequence, and sensitivity to cyanide and chloroform-ethanol treatment. I n addition to the manganese enzyme E. coli contains a closely related protein with iron in place of the manganese (202). It has been found that mitochondria from a variety of animal cells contain a manganese enzyme very similar to the prokaryotic superoxide dismutase (199,200,203-20‘06) . These similarities have lent important material to the discussion concerning the origin of mitochondria (155,206-213 ) . Very recently the marine bacterium, Photobacterium leiognuthi, was found to possess a protein with superoxide dismutase activity which contains one copper and two zinc atoms per molecule ( 2 1 4 ) . It appears to be dissimilar to the eukaryotic copper-zinc enzymes with respect also to the subunit composition, isoelectric point, optical properties, and pH 198. B. B. Keele, Jr., J. M. McCord, and I. Fridovich, JBC 245, 6176 (1970). 199. H. M. Steinman and R. L. Hill, Proc. Nat. Acad. Sci. U. S. 70, 3725 (1973). 200. R. A. Weisiger and I. Fridovich, JBC 248, 3582 (19731. 201. P. G. Vance, B. B. Keele, Jr., and K. V. Rajagopalan, JBC 247,4782 (1972). 202. F. J. Yost, Jr. and I. Fridovich, JBC 248, 4905 (1973). 203. R. A. Weisiger and I. Fridovich, JBC 248, 4793 (1973). 204. G. Beckman, E. Lundgren, and A. Tamvik, H u m . Hered. 23, 338 (1973). 205. S. Marklund, Acta Chem. Scand. 27, 1458 (1973). 206. D. Roodyn and D. Wilkie, “The Biogenesis of Mitochondria.” Methuen, London, 1968. 207. N. K. Boardman, A. W. Linnane, and R. M. Smillie, “Autonomy and Biogenesis of Mitochondria and Chloroplasts.” Amer. Elsevier, New York, 1971. 208. S. S. Cohen, Amer. Sci. 58, 281 (1970). 209. E. Schnepf and R. M. Brown, Jr., in “Results and Problems in Cell Differentiation” (J. Reinert and H. Unprung, e&.), Vol. 2, p. 299. Springer-Verlag, Berlin and New York, 1972. 210. L. Margulis, “The Origin of Eukaryotic Cells.” Yale Univ. Press, New Haven, Connecticut, 1970. 211. E. S. Goldring, L. I. Grossman, D. Krupnick, D. R. Cryer, and J. Marmur, J M B 52, 323 (1970). 212. H. R. Mahler and K . Davidowicz, Proc. Nat. Acad. Sci. U.S. 70, 111 (1973). 213. L. Reindars, C. M. Kleisen, L. A. Grivell, and P. Borst, BBA 272, 396 (1972). 214. K. Puget and A. M. Michelson, BBRC 58, 830 (1974).

538

B.. G. MALMSTROM, L.-E.

ANDREASSON, AND

B. REINHAMMAR

dependence of the activity. It remains to be demonstrated that it is evolutionally related to the eukaryotic copper-zinc enzymes.

B. PURIFICATION AND ASSAYMETHODS 1. Sources Superoxide dismutases containing two atoms each of copper and zinc have been obtained in purified form from a variety of animal, plant, and microorganism material, e.g., blood (15,16,216-218),brain (141,144), liver (16,144,200,219), heart (220),pea (221,222),spinach leaves (223), wheat germ (224),brewers’ yeast (219,226,226), and fungi (227,228). 2. Methods of Purification

a. Chloroform-Ethan02 Procedure. The enzyme can be purified from hemolysates in procedures involving fractionation with chloroform-ethano1 and acetone, usually in connection with precipitation with ammonium sulfate and sometimes heavy metal salts (16,16,816). Because of its unusual solubility properties, the enzyme will be dissolved in the salted out chloroform-ethanol phase. Kimmel et al. (229) modified the original purification method by replacing some of the later precipitation steps by electrophoretic fractionation. The chloroform-ethanol fractionation can be adapted for use in the purification of superoxide dismutase from other sources than blood. b. Ion-Exchange Methods and Gel Filtration. The original methods can be further modified by the use of ion-exchange chromatography with 215. H. Markowitr, G . E. Cartwright, and M. M. Wintrobe, JBC 234, 40 (1959). 216. M. J. Stansell and H. F. Deutsch, JBC 240, 4299 (1965). 217. M. J. Stanaell and H. F. Deutsch, JBC 240,4306 (1965). 218. J. W. Hartz and H. F. Deutsch, JBC 244,4565 (1969). 219. U. Weser, R. Prine, A. Schallies A. Fretzdorff, P. Krauss, W .Voelter, and W. Voetsch, Hoppe-Seyler’s Z. Physiol. Chem. 353, 1821 (1972). 220. B. B. Keele, Jr., J. M. McCord, and I. Fridovich, JBC 246, 2875 (1971). 221. Y. Sawada, T. Ohyama, and I . Yamasaki, BBA 268, 305 (1972). 222. Y. Sawada, T. Ohyama, and I. Yamasaki, in “Oxidases and Related Redox Systems” (T.E. King, H. S. Mason, and M. Morrison, eds.), p. 82. Univ. Park Press, Baltimore, Maryland, 1973. 223. K. Aaada, M. Urano, and M. Takahashi, Eur. J . Biochem. 36, 257 (1973). 224. C. 0. Beauchamp and I. Fridovich, BBA 317, 50 (1973). 225. S. A. Goscin and I. Fridovich, BBA 289,276 (1972). 226. U. Weser, A. Fretzdorff, and R. Prinr, FEBS (Fed. Eur. Biochem. SOC.)Lett. 27, 267 (1972). 227. H. P. Misra and I. Fridovich, JBC 247, 3410 (1972). 228. U. Rapp, W. C. Adams, and R. W. Miller, Can. J . Biochem. 51, 158 (1973). 229. J. R. Kimmel, H. Markowite, and D. M . Brown, BBA 234, 46 (1959).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

539

TABLE VI DISMUTASES FROM VARIOUSSOURCES~ SPECIFICACTIVITIESOF SUPEROXIDE Source

Units of activity/mg protein

Ref.

Human erythrocytes Bovine erythrocytes Chicken liver Wheat germ Green pea Spinach leaves Yeast Neurospora

3000 3300 3295 4300-4700 6400 9320 3330 2650

I6 I6 800 934

dBI 823 886

827

The activities have been determined according to McCord and Fridovich (16), sometimes with minor modifications.

or without a combination with gel filtration (16,216,217,230,231).These methods have been reported to induce less modification of the protein than the original chloroform-ethanol fractionation, a t least of the human enzyme (217) . 3. Methods of Assay

a. Xanthine Oxidase Procedure. Methods for the determination of superoxide dismutase activity (Table VI) exploit the ability of the enzyme to scavenge 0,- radicals which would normally reduce suitable electron acceptors. The original method of McCord and Fridovich (16,232) utilizes a system of xanthine and xanthine oxidase for the generation of 0,- radicals and ferricytochrome c as an electron acceptor. The presence of superoxide dismutase is thus characterized by the inhibition of cytochrome c reduction. Modified versions of this method have been discussed by McCord et al. (232) and Weser et al. (233,234). b. Other Methods. Nitro blue tetrazolium (NBT) can be substituted for ferricytochrome c, and aerobic photoreduction of certain dyes or flavins in the presence of suitable electron donors such as EDTA can be used to generate 0,- radicals (232,235).Superoxide dismutase prevents 230. P. 0. Nyman, BBA 45, 387 (1960). 231. J. Bannister, W. Bannister, and E. Wood, Eur. J. Biochem. 18, 178 (1971). 232. J. M. McCord. C. 0. Beauchamp, S. Goscin, H. P. Misra, and I. Fridovich, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 51. Univ. Park Press, Baltimore, Maryland, 1973. 233. U. Weser, W. Bohnenkamp, R. Cammack, H. J. Hartmann, and G. Voelcker, Hoppe-Seyler‘s Z . Physiol. Chem. 353, 1059 (1972). 234. U. Weser and G. Voelcker, FEBS (Fed. Eur. Biochem. Soc.) Lett. 22, 15 (1972). 235. C. Beauchamp and I. Fridovich, Anal. Biochem. 44,276 (1971).

540

B. G. MALMSTR~M, L.-E. ANDR~ASSON, AND B. REINHAMMAR

the formation of colored insoluble formazan. The NBT photoreduction method can be applied to detect superoxide dismutase activity on polyacrylamide gels (232,236).A simple assay method exploiting the ability of superoxide dismutase to inhibit the autoxidation of epinephrine to adrenochrome a t elevated pH has been devised by Misra and Fridovich (163). A convenient source of superoxide for assay purposes is a solution of the radical anion in an organic solvent, such as dimethylsulfoxide (DMSO). The solution is prepared by electrolytic reduction of dissolved oxygen or by simply dissolving potassium superoxide in the organic solvent (179,236,237). 4. Determination of Purity and Concentration

Starch or polyacrylamide gel electrophoresis of superoxide dismutase from several-sources generally reveals two components: a major band and a faster moving minor component (144,219,231).In some cases [for example, enzyme from chicken liver (200) and erythrocyte enzyme from certain human populations (EM)J the heterogeneity is even greater. The components are generally detected by their ability to prevent formazan formation in the assay method of Beauchamp and Fridovich (236).Consequently, electrophoretic homogeneity is not usually utilized as a criterion of purity. When the electrophoretically heterogeneous enzyme is analyzed in the ultracentrifuge or by isoelectric focusing technique, only one component is usually found (200,219,239).The major and minor components have been suggested to be “size” isomers (.t?s9,240).Because of the unusually low molar absorptivity at 280 nm of superoxide dismutase (Table VII), a slight contamination of most other proteins would immediately reveal itself as an absorbance increase at this wavelength. It is therefore convenient to use the ratio of the absorbances a t 280 nm and at some reference wavelength for the estimation of enzyme purity as described by Sawada et al. (221). The concentration of superoxide dismutase in solution is mostly determined from the absorbances a t 680, 260 ( 1 6 ) ,or 258 nm (341) (see Table

VII) .

236. J. A. Fee and P. G. Hildebrand, FEBS (Fed. Eur. Biochem. SOC.) Lett. 39, 79 (1974). 237. H.J. Forman and I. Fridovich, ABB 158,396 (1973). 238. G. Beckman, L. Beckman, and L.-0. Nileson, Hereditas 75, 138 (1973). 239. W. H. Bannister, D. G . Dalgleish, J. V. Bannister, and E. J. Wood, Znt. J . Biochem. 3, 560 (1972). 240. J. L. Hedrick and A. J . Smith, ABB 126,155 (1968). 241. G. Rotilio, L. Calabrese, F. Boasa, D. Barra, H. Finazzi Agrb, and B. Mondovi, Biochemktry 11, 2182 (1972).

00

8

1

Q

0

2

El

TABLE VII OPTICALPROPERTIES OF SUPEROXIDE DISMUTASE FROM VARIOUSSOURCES~

5

3

EG

Molar absorptivity

Wavelength (nm)

9

m

NeuroMan

Ref.

COW

Ref.

Chicken Ref.

237-3 13

1G,d31,846

Wheat

Ref.

Pea

Ref. Spinach Ref.

Yeast

Ref.

spora

Ref.

#

9

680 675 670 660 655 650 280 265 259 258 a

235-250

290

221

350

223

156

3

226

144J 46,,??18.239

233

200

330-360

227

3

11,700 227

8

231 226

224

490 250-350 16,300-18,500 17,OOO-18,700

2i7,218 146,220.239 146,216-218

250 8,100 9,400 9,840-10,300

295 220

4,400

221

M

231

16,246

Ex

8,750

200

9,740-9.960

224 8,800 22f 9,920 BBS

In a few eases the given molar abaorptivities have been obtained by recalculation of values originally given as Ei3m.

9,800 22G 11,300 226

17,400

227

542

B. G. MALMSTROM, L.-E.

ANDREASSON,AND

B. REINHAMMAR

5. Crystallization Conditions for successful crystal growth have been described by Richardson et al. (242) for the bovine enzyme. Monoclinic crystals grow from a 1.5 mg/ml solution of the enzyme in buffer with 57-5876 Z-methyl-2,4pentanediol, whereas orthorhombic crystals can be obtained from a 300 mg/ml solution of superoxide dismutase a t 4O. Both forms are suitable for X-ray crystallographic work and the unit cell parameters have been reported(242). The monoclinic crystals are blue along one crystal axis and green along another when viewed in unpolarized light. This suggests that the principal axes of all copper atoms in the crystal are not far from parallel (242,243). Rhombic crystals of spinach leaf superoxide dismutase have been grown by Asada et al. (223).

C. MOLECULAR PROPERTIES 1. Native Superoxide Dismutase a. Molecular Weight. The molecular weight of superoxide dismutase has been determined by a number of investigators and is found to be 31,000-33,000regardless of source ( 16,144,600,218-b21 ,223-225,2~?7,229). b. Quaternary Structure. Superoxide dismutase can be split into two subunits of equal size (molecular weight about 16,000)by treatment with denaturing agents, e.g., sodium dodecyl sulfate or guanidinium chloride (200,~23-~25,2~7,244,245). Earlier experiments indicated that reductants, such as mercaptoethanol, are required for the separation of the subunits in the bovine enzyme and this was interpreted in terms of the existence of at least one interchain disulfide bridge (220,241).However, recent investigations by Beauchamps and Fridovich (224) show the absence of such bonds but the presence of intrachain disulfide bonds. The resistance of the enzyme toward dissociation is explained by the stabilizing effect of such bridges on the conformation of the subunits which in their native state are strongly noncovalently associated. Hartz and Deutsch (244) found that removal of the metal, alkylation of two free sulfhydryls, SUCcinylation or reduction and alkylation of the protein facilitates dissociation of the human enzyme. On the basis of these results Hartz and 242. D. C. Richardson, C. J . Bier, and J. S. Richardson, JBC 247, 6368 (1972). 243. R. A. Lieberman and J. A. Fee, JBC 248,7617 (1973). 244. J. W. Hartz and H. F. Deutsch, JBC 247, 7043 (1972). 245. U. Weser, E. Brunnenberg, R. Cammack, C. Djerassi, L. FlohB, G . Thomas, and W. Voelter, BBA 243, 203 (1971).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

543

Deutsch suggested that metal-sulfur bonds may account for the stability of the protein against dissociation. Such possibilities have also been discussed by Fee (246). The two free sulfhydryl groups in human superoxide dismutase were found by Hartz and Deutsch (24.4) in two nonidentical peptides isolated after alkylation, which indicates that the two subunits are not identical (cf. 204). Also, the number of tryptic peptides support this notion. Rotilio et al. (241) found only one carboxyl terminal amino acid and one tryptophan in the bovine enzyme which suggests that this is also composed of two nonidentical subunits. However, according to Steinman and Hill (199) the recovery of peptides in sequence studies is consistent with the presence of identical subunits in the bovine erythrocyte enzyme. c. Amino Acid Composition. The amino acid composition of superoxide dismutase from animal sources has been reported by several investigators (144,6OO,217-621,226-227,229,~Sl,239) (Table VIII) . The content of tryptophan and tyrosine has been a matter of controversy. The finding of tyrosine (two residues) in the human enzyme by Kimmel et al. (229) has not been repeated (144,Zl 7,618,gSQ). Chemical investigations and ultraviolet absorption measurements have indicated zero to three tryptophan residues (144,Sl 7,218,220,229,247) in human superoxide dismutase. Various workers generally agree with the original report by Keele et al. (220) indicating two tyrosines in bovine superoxide dismutase, whereas results on the tryptophan content indicate zero (220,248,249) or one (ZSl,Z4l) residue. A total of six half-cystines are found in the bovine enzyme (219,220,226,241). The native protein does not react with SH reagents (231,241,246) but complete removal of the zinc exposes two SH groups (241). Rotilio et al. (241) found four sulfhydryls in the apoprotein in 6 M guanidinium chloride, Fee et al. (246) reported two under approximately equivalent conditions and claimed that the figure of Rotilio et al. should be reduced to two because of the use of a n incorrect extinction coefficient. The experimentally determined number of half-cystines in the human enzyme varies between 4 and 11 (l44,217,218,229,2S9). One free sulfhydry1 can be detected by titration of the native enzyme with p-chloromercurobenzoate (PCMB) or iodoacetamide (217,618). Under denaturing conditions, however, two SH groups are found ( 2 4 ) . 246. J. A. Fee, R. Natter, and G. S.T. Baker, BBA 295,96 (1973). 247. W. H. Bannnister, C. M. Salisbury, and E. J . Wood, BBA 168, 392 (1968). 248. U. Weser, G. Barth, C. Djerassi, H. J. Hartmann, P. Krausa, G. Voelcker, W. Voelter, and W. Voetsch, BBA 278, 28 (1972). 249. J. A. Fee, BBA 295, 87 (1973).

TABLE VIII TEE AMINOACID COMPOSITION OF SUPEROXIDE DISMUTASIGS FROM VARIOUSSOURCEV Amino acid

kc

Human Bovine erythrocytesberythrocytesc

np

22 16 23 7 36 16 20 26 10 50 20 28 0 16 17 0 8 0

7

10 35 26 20 24 14 50 21 28 0-2 j 17 20 2 10 0 6

MW

33,600

32,600

NHa Arg

%

Ser Glu Pro

GIY

Ala Val Met Ile Leu TYr Phe Half-cystine

a

Chicken liverd

Wheat germ' I I1

Neuro-

Green pea*

Spinach leaf9

Yeasth

spora'

22 16

20 14

10 19

8 15

10 18

13 14

18 11

12 11

8 32 18 14 23 12 22 28 4 14 16 2 8 0 14

8 28 33 15 21 19 55 28 31 2 13 22 0 7 0 6

10 28 30 12 26 19 43 25 34 0 10 31 0 6 0 4

6 45 30 14 19 14 56 21 21 0 20 21 0 9 0 6

7 35 28 10 20 17 42 23 28 2 6 22

7 32 18 20 25 20 40 24 28 2 9

9 36 26 14 20 14 39 20 22 0 13

11

11

0 6 0

2 10 0

4

4

2 6 0 3

30,400

31,000

30,900

31,500

32,200

50

32,700

31,000

The given figures are nearest integral values based on the tabulated molecular weights.

* From Harts and Deutsch From Keele ei al. (880).

(218).

From Weisiger and Fridovich (200). From Beauchamp and Fridovich (284). f From Sawada et al. ($21). 0 From Azada et al. (223). * From Goscin and Fridovich (225). i From Misra and Fridovich (887). j See Beauchamp and Fridovich (884).

W

d

Z

31

4

3

8.

COPPER-CONTAINING

OXIDASES

AND SUPEROXIDE DISMUTASE

545

Methionine has not been found in the human enzyme (144,217,218, 229,239), and Keele et al. (220) reported its absence in superoxide dismutase from bovine heart and blood. Others (219,664,6S1,641) have, however, detected two methionins in enzyme from bovine material. The amino terminal groups are blocked in the human (217) and bovine (631) enzymes. Although superoxide dismutases from sources other than man and cow have been much less studied, available results point a t significant similarities in amino acid composition (Table VIII) . Generally, tryptophan, tyrosine, and methionine are absent or present only in low amounts in enzymes from chicken liver ( Z O O ) , pea (221,222),spinach ( 2 2 3 ) , wheat germ (2241, and Neurospora ( 2 2 7 ) . Two different research groups have characterized superoxide dismutase from yeast (225,226). The two groups, although using essentially the same method for the purification of the yeast enzyme and both claiming homogeneous preparations, arrived a t such deviating results with respect to amino acid composition as well as optical and EPR properties that it is doubtful whether they studied the same protein. The isoelectric point is 4.75 and 4.95, respectively, for the human (218) and bovine (231) erythrocyte enzymes and varies from 5.35 to 6.75 for the different forms of the chicken liver enzyme ($60). d. Spectroscopic Properties. The optical spectrum of the superoxide dismutase is characterized by a broad absorption band in the visible region between 500 and 900 nm with a maximum a t about 680 nm which is responsible for the bluish-green appearance of concentrated solutions of the enzyme. The absorption of this band is enhanced when the protein is cooled to liquid nitrogen temperature (245). This band and another weaker one a t about 340 nm is thought to result from copper. As a result of the amino acid composition the spectral region between 250 and 300 nm is characteristically dominated by absorption by the phenylalanines. Contributions to the CD in the visible region occur a t 350, 440, 610, and 750 nm (245,250,261).The two latter bands are equal in rotatory strength but opposite in sign. The positions of these two CD bands indicate that the broad visible absorption centered a t 680 nm actually is composed of two components. These studies have been extended with magnetic optical rotatory dispersion (MORD) and magnetic circular dichroism (MCD) investigations (945,252). Unfortunately, the copper 250. E. J. Wood, D. G. Dalgleish, and W. H. Bannister, Eur. J. Biockem. 18, 187 (1971). 251. G. Rotilio, A. Finazzi Agr6, L. Calabrese, F. Bossa, P. Guerrieri, and B. Mondovi, Biochemistry 10, 616 (1971). 252. G. Rotilio, L. Calabrese, and J. E. Coleman, JBC 248, 3855 (1973).

546

B. G . MALMSTROM, L.-E.

ANDREASSON,

AND B. REINHAMMAR

ions are only weakly magnetically induced which limits the usefulness of these methods. I n the native superoxide dismutase positive contributions to the CD are observed in the ultraviolet region above 250 nm. At least some of these are induced by metal (239,245,260).Other bands are possibly related to amino acid residues, e.g., cystine, tyrosine, phenylalanine, and tryptophan (245,250). The content of a-helix is probably very low since no evidence has been found for the double minimum in the far ultraviolet C D spectrum characteristic of helix structure (245,250). On the other hand, the position of the negative ellipticity in this region suggests that superoxide dismutase, in addition to “unordered” structure, contains some p-pleated sheet. This is in agreement with a recent conformational analysis by Bannister et az. (253). Since superoxide dismutase contains paramagnetic copper, magnetic resonance methods provide excellent tools for the study of the copper sites. The first report on the EPR properties of superoxide dismutase was published by Malmstrom and Viinngbd in 1960 (254). Since then the results of many studies of the EPR spectrum of superoxide dismutase from various sources and under a variety of conditions have appeared in the literature (144,219,221,243,246,248,249,251,265-257) (Table I X ) . The field around the copper ions is distorted from tetragonal symmetry (243,251,256). Nitrogen superhyperfine structure can be seen a t pH 7.5 (245) but is much more evident a t higher pH values (233). Above pH 12 irreversible changes in the EPR spectrum occur with indications of a near tetragonal symmetry around the copper ion as is usually found with copper proteins at high pH. Addition of cyanide to superoxide dismutase reveals nitrogen superhyperfine structure in the E P R spectrum (251,257,258) with profound alteration of the symmetry of the copper sites. Experiments with I3CN- show that the cyanide ion binds to copper via the carbon atom and that the nitrogen superhyperfine structure resulting from cyanide binding is entirely due to the protein nitrogens (255,259). The cyanide experiments 253. W. H. Bannister, J. V. Bannister, P. Camilleri, and A. Leone Ganado, Int. f. Biochem. 4, 365 (1973). 254. B. G. Malmstrom and T. ViinngKrd, JMB 2, 118 (1960). 255. G. Rotilio, L. Morpurgo, C. Giovagnoli, L. Calabrese, and B. Mondovi, Biochemistry 11, 2187 (1972). 256. J. A. Fee and B. P. Gaber, Fed. Proc., Fed. Amer. SOC. E x p . Biot. 30, 1294 (1971). 257. J. A. Fee, BBA 295, 107 (1973). 258. J. A. Fee and B. P. Gaber, JBC 247, 60 (1972). 259. P. H. Haffner and J. E. Coleman, JBC 248, 6626 (1973).

8.

547

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

TABLE IX ELECTRON PARAMAGNETIC RESONANCE PARAMETERS OF SUPEROXIDE DISMUTASES OF DIFFERENT ORIGINSO Source

8. = 811

Human erythrocytes Bovine erythrocytes Green pea Spinach leaf Yeast Neurospora Fusarium oxysporum

2.260 2,257 2.23 2.243 2.255 2.26 2.27

BY

Bx

2.103

2.025

Bm

2.063 2.04 2.034 2.071 2.08 2.09

A II (G)

Ref.

152 132

$64

124

883 886 888

134 135

894 821

888

In the bovine enzyme the values of all three g tensors have been measured. In the other cases gm represents the g value a t maximum absorption.

may be taken to indicate that the copper ions are surrounded by three strongly bound magnetically equivalent nitrogen nuclei (251,255,257). Recent NMR studies are consistent with the idea that the nitrogen ligands to the copper ions are donated by three histidines (260-262; cf. 263). The cyanide-treated enzyme is more stable a t high p H values than the native enzyme indicating a competition between cyanide ions and denaturation-inducing hydroxide ions a t the same binding position ( 2 5 5 ) . When large quantities of azide are added to superoxide dismutase the spectral properties of the enzyme change drastically indicating the binding of azide to copper. A strong absorption band appears in the visible spectrum around 370 nm which is typical of Cu2+-N3-charge transfer (264) and the 680-nm band shifts t o lower wavelengths with somewhat increased intensity. The EPR spectrum changes to an axial-type spectrum (255,258,265) and the paramagnetic relaxivity is abolished (258,265). The paramagnetic relaxivity results from exchange of water close to the copper ions with bulk water and a decrease in relaxivity indi260. A. M. Stokes, H. A. 0. Hill, W. H. Bannister, and J. V . Bannister, FEBS (Fed. Eur. Biochem. Soc.) Lett. 32,119 (1973). 261. H . J . Forman, H. J. Evans, R. L. Hill, and I. Fridovich, Biochemistry 12, 823 (1973). 262. A. M. Stokes, H. A. 0. Hill, W. H. Bannister, and J . V. Bannister, Biochem. SOC.Trans. 2, 489 (1974). 263. R. C. Bray, S. A. Cockle, E. M. Fielden, P. B. Roberts, G. Rotilio, and L. Calabrese, BJ 139, 43 (1974). 264. W. E. Hatfield and R. Whyman, Transition Metal Chem. 5, 47 (1969). 265. J. A. Fee and B. P. Caber, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.). p. 77. Univ. Park Press, Baltimore, Maryland, 1973.

548

B. G. MALMSTROM, L.-E. ANDRI~ASSON, AND B. REINHAMMAR

cates that the water is displaced from the copper. Fee and Gaber (258,265) and Gaber et al. (266) have shown that water is accessible to two more or less identical copper sites. Rotilio et al. (855) and Morpurgo et al. (267) suggested that azide binds to copper already at [N3-]/[Cu] = 1, whereas Fee and Gaber (258,265) interpreted their results with low concentrations of azide as indicating the binding of the anion to the zinc atoms. They furthermore suggested that the zinc atoms are accessible to SCN- and OCN- ions. Fee and Gaber (258) found only negligible effects of F- (and S2032-)even a t high concentrations on the spectral properties and paramagnetic relaxivity of superoxide dismutase indicating the absence of strong interaction with these anions. Rotilio et al. (255),however, observed large changes in the spectroscopic properties of the enzyme with fluoride, similar to those obtained with azide. No obvious explanation for the discrepancies in the results of the different research groups is presently available, The metal sites in superoxide dismutase have also been studied by X-ray photoelectron spectroscopy (268-271). 2. Apoenzyme a. Preparation. The metal atoms in superoxide dismutase can be removed from the enzyme molecule by various means. Extended dialysis of the bovine protein against EDTA at about pH 3.4-3.8 results in the loss of virtually all metal (18,231), whereas dialysis a t higher pH values against 1,lO-phenanthroline or EDTA is not effective (145). Weser et al. (248,272) passed bovine superoxide dismutase through a column of Sephadex G-25 equilibrated with EDTA a t pH 3.8 with complete loss of the metal as a result as judged from EPR and atomic absorption spectroscopy. The metals can also be removed by dialysis against KCN a t pH 8 a t room temperature (145) and less effectively in the cold (216). Cyanide dialysis, however, has been reported to lead to some damage of the protein (241,248).Rotilio et al. (241) were able selectively to remove copper without significant denaturation by reduction of the copper with ferrocyanide prior to the cyanide dialysis. 266. B. P. Gaber, R. D. Brown, 5. H. Koenig, and J. A. Fee, BBA 271,l (1972). 267. L.Morpurgo, C.Giovagnoli, and G. Rotilio, BBA 322, 204 (1973). 268. G.Jung, M. Ottnad, W. Bohnenkamp, W. Bremser, and U. Weser, BBA 295, 77 (1973). 269. G. Jung, M. Ottnad, W. Bohnenkamp, and U. Weser, FEBS (Fed. Eur. Biochem. Soc.) Lett. 25, 346 (1972). 270. G.Jung and M. Ottnad, 2.Anal. Chem. 263,282 (1973). 271. G. Jung, M.Ottnad, W. Bremser, H. J. Hartmann, and U. Weser, Hoppe-Seyler's Z . Physiol. Chem. 354, 341 (1973). 272. U. Weser and H. J. Hartmann, FEBS (Fed. Eur. Biochem. Soc.) Lett. 17, 78 (1971).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

549

b. Molecular Properties of the Apoenzyme. Removal of the metal leads to significant changes in the spectroscopic properties of superoxide dismutase. Large changes occur in the optical spectrum between 250 and 350 nm and the band a t 680 nm disappears. On the other hand, the region below 250 nm is largely unaffected which indicates that no gross changes in the tertiary structure occurs following the removal of metals (145,6~1,64l,248-25O).The positive ellipticity bands above 250 nm present in the native enzyme are virtually absent in the apoprotein or in the protein containing only zinc (241). The stability of the apoprotein toward extremes in pH and denaturing agents, although still striking, is less than that of the native enzyme (241,248). Obviously, the metals exert a stabilizing effect on the threedimensional structure (231,239,241,246,250). Forman and Fridovich (273) have investigated the effect of different metals on the stability of the enzyme and found that Co2+and Hg2+are able to replace Zn2+in exerting a stabilizing effect toward thermal denaturation. 3. Reconstitution of the Apoenzyme

a. Restoration of the Native Enzyme. Exposure to the apoprotein to divalent copper ions in aqueous solutions leads to restoration of most of the activity of the enzyme (16). Addition of a t least two copper ions per enzyme molecule appears to be necessary for restoration of activity although a t least four can be bound (248). The sites becoming occupied when Cu2+is simply added to the apoprotein, however, differ from the native copper binding sites (246,249,257), which is evident from EPR spectra. The presence of zinc seems to be necessary for the binding of copper to the native sites and it has been suggested that the removal of zinc leads to irreversible changes in the protein such as interchange of S H groups exposed by zinc removal with a disulfide bridge, which prevent the recombination of copper (241), Others have, however, reconstituted the enzyme by dialysis of the apoprotein against a solution of Zn2+ and Cuz+ (257).There is evidence that binding of zinc to the apoprotein preforms the native copper sites and permits the ra,pid incorporation of copper resulting in a catalytically active enzyme indistinguishable from the native one (257). b. Preparation of Modified Enzymes. Because of the physical properties of zinc the binding site for this metal is only to a limited extent accessible to study. Fee (257) reported that when the apoprotein was dialyzed against high concentrations of Cuz+,copper seemed to bind to the zinc site (cf. 273) and a copper E P R signal appeared which indicated 273. H. J. Forman and

I. Fridovich, JBC 248, 2645 (1973).

550

B. G. MALMSTROM,

L.-E.

ANDROPSSON,

AND B. REINHAMMAR

a dipolar coupling between the Cuz+ ions. This was taken as evidence for rather close proximity of the copper and zinc sites. Valuable information has been obtained by substitution of Co2+ for zinc in superoxide dismutase. This can be accomplished by dialysis of the native enzyme against a solution of CoCl, followed by water dialysis for the removal of excess Co2+according to Calabrese et al. (274,276).Fee (276') dialyzed the apoprotein against Co2+and Cu2- in stoichiometric amounts or added Cu2+to the 2 Coz+-protein which had been previously formed by dialysis of the apoprotein against Co2+.These methods result in a product which contains two equivalents each of Cu2+and Coz+ and with retained catalytic activity (275,276). The procedure of Fee, however, may result in some nonspecifically bound copper (276').The replacement of zinc with cobalt results in changes in the optical properties of superoxide dismutase. The absorption of the copper centers is not dramatically altered, but the C D band a t 346 nm resulting from copper is accentuated in the 2 Cu2+ 2 Coz+-enzyme (276). The optical spectrum of cobalt-containing superoxide dismutase is characterized by several more of less resolved bands between 530 and '600 nm which are caused by the Co2+ion. The positions and strength of these bands are almost identical with the corresponding properties of the bands of the cyanide complex with human CoZ+-carbonicanhydrase (262,276'$77) for which it has been suggested that the bands are caused by Co2+in a tetrahedral field (278-280). The CD spectrum of cobalt superoxide dismutase is of opposite sign compared to that of CN--cobalt carbonic anhydrase (262,276).The MCD spectrum, which is less dependent on the external protein potential field (280) than the CD spectrum, should more accurately reflect the symmetry of coordination to the cobalt ions, and MCD spectra of cobalt superoxide dismutase and CN--cobalt carbonic anhydrase are very similar (262). Comparison of EPR spectra of the two proteins will give information as to whether or not the conclusions drawn from the optical spectral similarities are correct. Reduction of the copper ions in the cobalt enzyme results in some change in the MCD of the Co2+indicating the existence of interaction between the copper and cobalt binding sites (252,276).The E P R signal resulting from the paramagnetic ions in oxidized cobalt-containing superoxide dismutase accounts for only 274. A. Rigo, M. Terenei, C. Franconi, B. Mondovi, L. Calabrese, and G . Rotilio, FEBS (Fed. Bur. Biochem. Soc.) Lett. 39, 154 (1974). 275. L. Calabrese, G. Rotilio, and B. Mondovi, BBA 283, 827 (1972). 276. J. A. Fee, JBC 248, 4229 (1973). 277. G. Rotilio, Biochem. SOC.Trans. 1, 50 (1973). 278. S. Lindskog and A. Ehrenberg, J M B 24, 133 (1967). 279. E. Grell and R. C. Bray, BBA 236, 503 (1971). 280. J. E. Coleman and R. V. Coleman, JBC 247,4718 (1972).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

551

a small part of the Co2+and Cult content (275-277,281). On reduction of the protein, so that copper ions become diamagnetic, the cobalt signal increases significantly in amplitude (876,277,281) . This indicates that magnetic interaction exists between the copper and cobalt sites (275,376,281).This possibility is also supported by NMR studies (274) showing that Cuz+in proximity of Coz+is not active in relaxation of water protons (cf. 258,265). Calabrese et al. (275) reported that the weak EPR copper signal of cobalt-containing superoxide dismutase was indistinguishable from that of the native protein, whereas Fee (276) found that the signal differed markedly. The latter author suggested that it resulted from nonspecifically bound Cu2+which could have been introduced during the preparation procedure. The copper signal of Calabrese et al., on the other hand, could well result from unaltered native molecules of the enzyme. The cobalt sites do not seem to be accessible to anions (277,281) with the exception of cyanide at high concentrations (276), which condition leads to removal of cobalt. Recent magnetic relaxation studies by Rigo et al. (274) indicate that the cobalt is not accessible to rapidly exchanging water molecules. If it is true that cobalt is binding to the zinc site in the same manner as zinc does, the above results may call for a reinterpretation of some anion binding data. Reconstitution experiments with metals other than copper and zinc or cobalt have not led to restoration of catalytic activity (16,273). 4. Redox Properties of the Copper Ions in Superoxide Dismutase

The addition of hydrogen peroxide to a solution of superoxide dismutase results in a bleaching of the 680-nm band and a simultaneous disappearance of the copper EPR signal (241,282-284). It has been suggested that the enzyme copper is reduced by H,O, according to the scheme 2 Cu(I1)

+ Hz02 + 2 Cu(1) + 2 Hf + O?

(6)

Sulfide and ferrocyanide can also accomplish a bleaching of the 680-nm band (284,285). The production of ferricyanide shows that a true reduction of the enzyme takes place. From the reaction with ferrocyanide a 281. G. Rotilio, L. Calabrese, B. Mondovi, and W. E. Blumberg, JBC H 249, 3157 (1974). 282. M. A. Simonyan and R. M. Nalbandyan, FEBS ( F e d . Eur. Biochem. Suc.) L e t t . 28, 22 (1972). 283. J. V. Bannister, W. H. Bannister, R. C. Bray, E. M. Fielden, P. B. Roberts. and G. Rotilio, FEBS (Fed. Eur. Biuch.em. Soc.) L e t t . 32, 303 (1973). 284. G. Rotilio, L. Morpurgo, L. Calabrese, and B. Mondovi, BBA 302, 229 (1973). 285. J. A. Fee and P. E. Dicorleto, Biochemistry 12,4893 (1973).

552

B. G. MALMSTR~M, L.-E.

ANDREASSON, AND

B. REINHAMMAR

reduction potential a t p H 7 (Eo’) of the copper ions of +0.42 V has been determined (284; cf. 285). Fee and Dicorleto (285) demonstrated that the reduction of copper is accompanied by the uptake of protons which results in p H dependence of the oxidation-reduction potential of the copper ions. The experiments indicate that the pK, of the protonaccepting group, which may well be a ligand to copper, must be greater than 9. The reaction can be written as ECu(I1) + H+ + eHE+Cu(I) (7) If large excesses of H,O, are added to a solution of superoxide dismutase the enayme is inactivated with concomitant marked changes in optical and EPR spectroscopic properties (224,263,282,286) suggesting changes in the surrounding of the copper ions. Bray et al. (263) determined the amino acid composition after H,O, treatment and found that histidine had been destroyed. Since histidine is not normally oxidized by H202 this reagent seems to act as an active-site-directed modifier of superoxide dismutase. When CN- or N,- is added to the ferrocyanide-reduced enzyme, reoxidation of the copper occurs as judged from optical and EPR spectroscopic measurements (277,284). Simultaneously, ferricyanide is converted to ferrocyanide. A reasonable explanation to these observations would be that the redox potential of the copper ions is affected by the presence of anions as has been observed with other copper-containing proteins (cf. 287,288). Strangely enough, no effect of CN- was found on the protein reduced with hydrogen peroxide (284).

D. THE CATALYTIC MECHANISM 1. The Enzymic Dismutation of 0,-

The first studies of the kinetic properties of superoxide dismutase and the identification of the activity were made in systems where enzymically or chemically generated superoxide was used to reduce cytochrome c or tetranitromethane (16). The presence of superoxide dismutase activity inhibited the reduction, and a rate constant for the reaction between superoxide radical and superoxide dismutase could be calculated. As shown later (237), the rate constant was overestimated by two orders of magnitude but the experiments served to demonstrate the extremely high activity of the enzyme. Forman and Fridovich (237) as well as 286. E. M. Fielden, P. B. Roberts, R. C . Bray, and G . Rotilio, Biochem. SOC.Trans. 1, 52 (1973). 287. B. Reinhammar and T. Vanngird, Eur. J. Biochem. 18, 403 (1971). 288. B. Reinhammar, BBA 275, 245 (1972).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

553

Sawada and Yamasaki (289) have shown that it is possible to obtain useful information on reaction rates and mechanism of superoxide dismutase despite the complexity of the xanthine oxidase-cytochrome c system. By the use of E P R Ballou et al. (290) directly demonstrated that superoxide dismutase accelerated the decomposition of superoxide radicals a t a very high rate. Recently, the introduction of pulse radiolysis into enzymology has provided other means for the direct study of the rapid reaction between superoxide dismutase and superoxide. The main advantages of this technique is a simple reaction system, a high yield of 0,- radicals, and the high time resolution (291,292). Pulse-radiolytic studies have shown that the second-order rate for the reaction of 0,- with the enzyme governs the turnover. This rate constant is about 2 X lo9 M-l sec-I for the bovine (291-293) and human (283) enzymes. This value is close to that expected for a diffusion-controlled reaction. Rotilio et al. (292) and Fielden et al. (294) found that increased viscosity resulting from added glycerol led to a lower reaction rate. The rate constant for the reaction between enzyme copper in the oxidized or reduced form and superoxide radical is about 2 x lo9 M-' sec-1 (294,295), which is consistent with these reactions being rate-limiting under turnover conditions. Analysis of data has so far provided no evidence for a Michaelis complex (283,292,294). The rate constant for the limiting step is independent of pH in the range 5-10 (283,291-293) in contrast to the spontaneous dismutation (296), but decreases below 4.8 (291,293). This implies that the radical anion rather than H02 is the substrate since pK, for HO, is 4.8 (293,296,297). When superoxide is allowed to react with native superoxide dismutase under pre-steady-state or turnover conditions, partial reduction of the copper occurs as judged from the absorbance a t 680 nm or from EPR intensities. If, on the other hand, the reduced enzyme reacts with 02-, copper is partially reoxidized (283,294,295) indicating involvement of the copper in a valence shuttling mechanism of the type

+ 02-e Cu(1) + H+ Cu(1) + e Cu(I1) + Cu(I1)

0 2

2

02-

€1202

(8) (9)

289. Y. Sawada and I. Yamasaki. BBA 327,257 (1973). 290. D. Ballou, G. Palmer, and V . Massey, BBRC 36, 898 (1969). 291. D. Klug, J. Rabani, and I. Fridovich, JBC 247, 4839 (1972). 292. G. Rotilio, R. C. Bray, and'E. M . Fielden, BBA 268, 605 (1972). 293. J . Rabani, D. Klug, and I. Fridovich, Isr. J. Chem. 10, 1095 (1972). 294. E. M . Fielden, P. B. Roberts, R. C. Bray, D. J. Lowe, G. N. Mautner, G . Rotilia, and L. Calabrese, BJ 139, 49 (1974). 295. D. Klug-Roth, I. Fridovich, and J. Rabani, JACS 95, 2786 (1973). 296. J. Rabani and S. 0. Nielsen, J. Phys. Chem. 73, 3736 (1973). 297. D. Behar, G. Czapski, J. Rabani, L. M. Dorfman, and H. A. Schwarz, J. P h s . Chem. 74, 3209 (1970).

554

B. G. MALMSTR~M, L.-E. ANDR~ASSON, AND B. REINHAMMAR

H,O, generated in the process can reduce divalent enzyme copper and irreversibly inactivate the enzyme (263,282,284,285), which can make analysis of kinetic results difficult as discussed by Fielden et al. (294). The molecular activity of the enzyme has been found to be independent of the initial state of the enzyme, oxidized or reduced, as is the bimolecular rate of reaction between enzyme and 0,- (285,886,893,294). The steady-state level of oxidized copper, measured as the strength of the 680-nm absorption band or the copper E P R signal, is strongly dependent on the initial oxidation state of the copper (283,294). This variation in steady-state level of oxidized copper can be interpreted to indicate the participation of only half of the copper atoms in turnover, according to Fielden et al. (894) who presented a model in which the reaction of one of two initially identical copper ions, Cu(I1) or C u ( I ) , renders the other transiently nonreactive toward 0,- according to the following scheme:

-

-kt 0;R-CdII)

k., 0;

N-CdII)

:;-,

R-Cu(II)

ks O<

k - s 0;

R--Cu(I)

Each box represents one enzyme subunit; R and N denote copper, reactive or nonreactive, respectively, toward 02-; k,, k+, k,, and k., control the fast turnover reaction, whereas k , and k-, represent much slower rates. Fielden et al. concluded that the conformational changes controlling the reactivity of the subunits during turnover must take place very quickly with rate constants greater than los sec-I and therefore probably involve only minor movements within the enzyme molecule. Klug-Roth et al. (296) used computer fitting of pulse-radiolytic data to determine the rate constants for the reactions of different forms of the enzyme with superoxide and oxygen. Their results are in many respects similar to those of Fielden et al. (29/t), for example, the equal magnitudes of the rate constants for the reduction of the native enzyme and the reoxidation of the reduced enzyme and the turnover rate constant. Also, the steady-state level of oxidized copper of the native enzyme is similar to that reported by Fielden et al. Furthermore, Klug-Roth et al. (995) found that the reduction of the second copper ion must be considerably slower than the reduction of the first (cf. 294). Their model can be summarized as follows:

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

555

Eo refers to the native enzyme with both copper ions in their divalent state while E- and E2- are molecules with one or both copper ions, respectively, in their univalent form. Reactions (11) and (12) are very rapid with rate constants of the order of lo9 M-l sec-I. These steps are proposed to account for the activity of the enzyme in the absence of H,O, since reaction (13) is very much slower than (11) or (12). Hydrogen peroxide, through reaction (15), is believed to modify the enzyme and in such cases reactions ( l l ) , (12), and (14) control the activity. I n the model of KlugRoth et al. (295) the copper ions are not initially equivalent but are given different molar absorptivities a t 650 nm, whereas in the more elaborate model of Fielden e t al. (294) the copper ions are initially indistinguishable. The latter situation seems to be favored by EPR investigations of the enzyme in the resting state or in kinetic experiments (255,257,294). The-results of Fee and Dicorleto (285) indicate that the reactions between superoxide dismutase and superoxide might be written as

+ +

+ H+ HE+Cu(I) + + a+-+ECu(I1) + H202

(16) (17) However, catalysis is not pH-dependent. In addition, the authors demonstrated that involvement of a hydronium ion in any rate-limiting catalytic step is excluded on the basis of the limited diffusion of free hydronium ions and concluded that the mechanism by which the oxidizing 0,- accepts two protons is not understood. Hodgson and Fridovich (298) have presented evidence indicating that at high pH O 2 can be reduced by H,O, in the presence of superoxide dismutase, i.e., a reversal of the dismutation reaction, according to the folIowing scheme: ECu(I1) HE'Cu(1)

0 2 02-

+

+

0 2

ECu(II) H0z- + ECu(I)-OzH ECu(I)-02H + ECu(1) Hf ECu(1) 0 2 -+ ECu(I1) 0 2 -

+

+

+

+

02-

The superoxide radicals are effectively scavenged by tetranitromethane (TNM) to yield nitroform (237,299).The rate of production of nitroform is dependent on the concentrations of 0, and enzyme but independent of TMN concentration. This reaction is also saturable with respect to the concentration of H,O,. The reaction between reduced superoxide dismutase and molecular oxygen has been shown to be extremely slow (15,663,677,285,695).This can be expected in view of the redox potentials of the couples 02/0,-and ECu(II)/ECu(I). The latter is +0.42 V a t pH 7 according to Fee and 298. E. K. Hodgson and I. Fridovich, BBRC 54,270 (1973). 299. J. Rabani, W. A. Mulac, and M. S. Matheson, J. Phys. Chem. 69, 53 (1965).

556

B. G : MALMSTROM, L.-E. ANDR~ASSON, AND B. REINHAMMAFI

Dicorleto (285).The work by George (300) is usually cited when discussing the E,’ value of the O,/O,- couple. George gives the value -0.45 V (0, fugacity 1 atm) a t pH 7 and a pK, for HO, of 2.2. With a pK, of 4.8 (296,297) E,’ decreases to -0.59 V. Recently, data have become available which permit recalculation of this important value. From the equilibrium constant for the reaction 02-

+ duroquinone

O2

+ durosemiquinone

(21)

( S o l ) , the semiquinone equilibrium data (302) and E,‘ for duroquinone/durohydroquinone, Wood (140) calculated an E,’ value for O,/O,-

of -0.33 V with a partial pressure of 0, of 1 atom as standard state (303,304) or -0.16 V with a standard state of 1 M Oz. With the former value AGOfor reaction (20) will be about 72 kJ mole -l. This corresponds to the minimum free energy of activation of the reaction and represents a significant barrier which may explain the slowness of the reoxidation (285). The E,’ value for the copper ions in the enzyme is intermediate relative to the E,’ values of the O,/O,- and 0,-/H,O, redox couples which is advantageous in a catalytic mechanism involving the dismutation of superoxide. Fielden et al. (294) observed that the aerobic reoxidation of reduced superoxide dismutase could proceed a t a considerable rate in the presence of catalase and suggested that catalase acts as a peroxidase with reduced superoxide dismutase as electron donor. This observation is in conflict with results by Fee and Dicorleto (285) and Klug-Roth e t al. (295) who observed only very slow reoxidation in the presence of catalase. Fee and Dicorleto (285) found that a stimulating effect by catalase on reoxidation was obtained with prior addition of H,O,. They, however, noted that the hydrogen peroxide should be decomposed within seconds after addition of catalase, whereas the stimulated reoxidation of superoxide dismutase proceeded for several minutes. 2. Inhibition of the Enzymic Activity

Cyanide has been found to be a potent inhibitor of superoxide dismutase (16,223,224,292)which accentuates the role of copper in the catalytic mechanism. Azide, on the other hand, has only a marginal effect on the activity (292),which seems surprising in view of the affinity of enzyme copper for this anion (255,258,267). 300. P. George, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 3. Wiley, New York, 1965. 301. K.B.Pate1 and R. L. Willson, JCS, Faraday Trans. 1, 816 (1973). 302. C. A. Bishop and L. K. J. Tong, JACS 87,501 (1965). 303. V. M. Berdnikov and 0. S. Zhuravleva, Zh. Fiz. Khim. 46, 2658 (1972). 304. J. Chevalet, F. Rouelle, L. Gierst, and P. Lambert, J . Electroanul. Chem. 39, 201 (1972).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

557

3. Catalytically Active Model Complexes

Joester et al. (190) found that Cu(I1) complexes of lysine, histidine, and tyrosine or of some di- and tripeptides displayed superoxide dismutase activity amounting to some 5% of the activity of native superoxide dismutase. Aqueous Cu2+, Cu (11)-EDTA, or copper bound to bovine serum albumin were totally inactive (cf. 232). As noted earlier a significant activity is shown by complexes between apo-superoxide dismutase and divalent copper even though copper is not bound in the native site. Rabani et al. (305) found that copper ions in acidic aqueous solution are capable of catalyzing the dismutation of superoxide a t a rate higher than that of the enzyme-catalyzed reaction a t physiological pH. It is pointed out (294) that this is a unique example of a model reaction which is more efficient than the enzymic reaction itself. Cleveland and Davis (306) have observed that galactose oxidase has the ability to catalyze the dismutation of 0,- radicals. The activity of the enzyme, however, seems to be rather low or somewhat less than the activity of model copper complexes. Concerning some very important recent studies on superoxide dismutase, see addendum ( 3 0 6 ~ ) . IV. The Blue Copper-Containing Oxidases

A. INTRODUCTION 1. History

As seen from Table I the blue oxidases include ascorbate oxidase, ceruloplasmin, and laccase. Of these laccase was discovered first. I n 1883, 305. J. Rabani, D. Klug-Roth, and J. Lilie, J . Phys. Chem. 77, 1169 (1973). 306. L. Cleveland and L. Davis, B B A 342, 517 (1974). 306%.After the completion of this review, a few important contributions concerning the structure of superoxide dismutase have appeared. Thomas et al. (306%) have described the X-ray structure at 5.5 A resolution, and Steinman and coworkers (SD6c,S06d) have presented the complete primary structure of the bovine superoxide dismutase subunit. Apart from giving a clear picture of the general shape of the dimeric molecule, these studies have given information about secondary structure, the position of the disulfide bridges, the nature and location of the copper ligands, and the possible position of the zinc-binding sites. I n addition, ambiguities concerning the amino acid content have been removed. 306b. K. A. Thomas, B. H. Rubin, C. J. Bier, J. S. Richardson, and D. C. Richardson, JBC 249, 5677 (1974). 306c. H. M. Steinman, V. R . Naik, J. L. Abernethy, and R. L. Hill, JBC 249, 7326 ( 1974). 306d. J. L. Abernethy, H. M. Steinman, and R. L. Hill, JBC 249, 7339 (1974).

558

B. G. MALMSTROM,

L.-E. ANDRJ~ASSON, AND B. REINHAMMAR

Yoshida (307) demonstrated that the darkening and hardening of the latex of the Oriental lacquer tree involves a thermolabile substance, which, about a decade later, was shown to be an enzyme and given the name “laccase” by Bertrand (4; cf. Section I and 5,6). The discovery of ascorbate oxidase was made by Szent-Gyorgyi (308), who found this enzyme in cabbage leaves in 1931. In neithey case were the enzymes recogniied to be copper proteins by their discoverers. Somewhat ironically, the occurrence of a specific ascorbate oxidase was, in fact, for a long time doubted by many investigators because ionic Cu2+was known to catalyze ascorbate oxidation (see 309).Incontrovertible evidence for the existence of a specific copper protein with ascorbate oxidase activity was first provided by Dawson and his collaborators (see 310) as late as around 1950. By that time the involvement of copper as a prosthetic group in oxidases was, of course, established through the work of Keilin (see Section I). In 1948, Holmberg and Laurell (311) reported the isolation of a copper protein from pig and human serum. It was distinguished from other copper proteins found earlier in mammalian blood by, among other things, its very intense blue color. This led the discoverers to suggest the name “coeruloplasmin” (31.2) for the new protein. They also showed that it has oxidase activity with a rather broad specificity similar to that of laccase. Good substrates are, for example, quinol (hydroquinone) and p-phenylenediamine. The specific activity was, however, found to be very low compared to that of laccase. 2. Biological Distribution and Functions

Laccase is widely distributed in plants and microorganisms. Soon after his discovery of the enzyme in the lacquer tree, Bertrand (313) demonstrated its presence in the fruit bodies of many fungi. The function of the enzyme in trees is apparently a protective one similar to that of the blood clotting system of animals. When a lacquer tree becomes damaged, the latex, containing various phenolic compounds and laccase, seeps out. I n contact with air laccase induces a catalytic oxidation of the phenols 307. H. Yoshida, JCS 43,472 (1884). 308. A. Szent-Gyorgyi, JBC 90, 385 (1931). 309. T. P. Singer and E. B. Kearney, in “The Proteins” (H. Neurath and K. Bailey, eds.), 1st ed., Vol. 2, Part A, p. 124. Academic Press, New York, 1954. 310. C. R. Dawson and W. B. Tarpley, “The Enzymes,” 1st ed., Vol. 2, Part 1, p. 454, 1951. 311. C. G. Holmberg and C. B. Laurell, Acta Chem. Scand. 2, 550 (1948). 312. The spelling ceruloplasmin is usually employed nowadays, particularly by

American authors. 313. G. Bertrand, C. R. Acad. Sci. 123,463 (18961.

8.

COPPER-CONTAINING

OXIDASES AND SUPEROXIDE DISMUTASE

559

to free radical products (cf. Section IV,E,2,a). These then polymerize spontaneously to form a protective natural plastic. The biological function of laccase in fungi is less obvious. A possible clue comes from the common occurrence of the enzyme in lignin-decomposing Basidiomycetes. It has been found (see, for example, 314) that the mycelia of these secrete laccase as an inducible exoenzyme. Its function may thus be a digestive one, making nourishment from the rotting wood available to the fungus. Laccase-catalyzed reactions by which lignin could be decomposed have been demonstrated (315,316). Synthetic functions, for example, in the formation of structural pigments (cf. the tree enzyme) or even of lignin (see S I B ) , have also been suggested. Despite its enzymic activities ceruloplasmin, which is of general occurrence in mammalian blood, has been considered a storage or transport protein for copper (see 9). This view is, in particular, supported by observed low levels of ceruloplasmin in Wilson’s disease, a rare hereditary disorder involving abnormal copper metabolism. Broman (317’) suggested that ceruloplasmin is a specific precursor for the incorporation of copper in cytochrome oxidase, and clinical observations in support of this idea have been reported (318). Recently, it has been argued (319) that it is a “ferroxidase,” since Fez+ is a fairly good substrate (see Section IV,E,2). The fact that the same reaction is catalyzed by several components in blood, including proteins as well as simple compounds, such as citrate, speaks against this proposed function. In addition, patients with Wilson’s disease may lack ceruloplasmin but have normal hemoglobin levels. An ascorbate-oxidase function of ceruloplasmin has also been considered, but ascorbate is, in fact, a very poor substrate (320). Ascorbate oxidase is found in many plants (see 310,321) but its function is unclear. An early suggestion, which was sometimes made for laccase as well, is that the enzyme serves as a terminal oxidase (see SO9), but this appears unlikely for several reasons. First, ascorbate oxidase, like laccase, catalyzes the direct transfer of electrons from the reducing substrate to dioxygen. Second, electron transfer chains with cytochrome oxidase as the terminal enzyme are also found in plant tissues (322). 314. G. Fkhraeus, Physiol. Plant. 5,284 (1952). 315. M. A. Pickard and D. W. S. Westlake, Can. J. Biochem. 48, 1351 (1970). 316. T. K. Kirk, J. M. Harkin, and E. B. Cowling, BBA 165, 145 (1968). 317. L.Broman, Actu Soc. M e d . Upsal. 69, Suppl. 7 (1964). 318. M. H. K. Shokeir and D. C. Schreffler, Proc. N a t . h a d . Sci. U . S. 62, 867 (1969). 319. S.Osaki, D.A. Johnson, and E. Frieden, JBC 241, 2746 (1966). 320. G.Curzon and S. N. Young, BBA 268,41 (1972). 321. G.R.Stark and C. R. Dawson, “The Enzymes,” 2nd ed., Vol. 8,p. 297, 1963. 322. J. Bonner and J. E. Varner, “Plant Biochemistry.” Academic Press, New York. 1965.

560

B. G. MALMSTROM,

L.-E. ANDRJ~ASSON, AND B. REINHAMMAR

B. PURIFICATION AND SOMEMOLECULAR PROPERTIES The purification methods for the blue oxidases available a decade ago have been reviewed ( 9 ) , and here some recent developments only will be mentioned. Ascorbate oxidase has been purified from species of squash, particularly, green zucchini (323-366). The enzyme has also been prepared from cucumber, Cucurnis sativus (326).The procedures all include extraction with acetone, ammonium sulfate fractionation, and chromatography on cellulose ion exchange columns. The purified proteins show a high degree of homogeneity in ultracentrifuge and electrophoresis experiments. The main sources of laccase have been the Japanese lacquer tree, Rhus vernicifera (327,328), and the fungus, Polyporus versicolor (329).Fungal laccase is an inducible exoenzyme. When a suitable synthetic medium is used, the protein fraction of the medium consists almost exclusively of laccase, so that a small number of purification steps is sufficient (369). The procedures used for the tree enzyme are similar to those for ascorbate oxidase but, because of the high solubility of the enzyme, increased yields are obtained when ammonium sulfate precipitation is avoided (328).The enzymes isolated from both sources show a high degree of purity, even if fungal laccase B yields a large number of components in isoelectric focusing electrophoresis (330). While pure ceruloplasmin was first prepared from pig plasma (311), most work has been with the protein from human blood. In the late 1940’s Gurd and others, working in E. J. Cohn’s laboratory, had prepared a greenish blue plasma protein fraction, and it was soon recognized that the color came from the ceruloplasmin of Holmberg and Laurel1 [see Peisach et al. (9, p. 513) 1. Cohn fraction IV-1 (331) is a common starting material for the final purification of human ceruloplasmin (see 336). Chromatographic procedures starting from outdated blood bank plasma 323. L. Avigliano, P. Gerosa, G. Rotilio, A. Finasai Agrb, L. Calabrese, and B. Mondovi, Ital. J. Biochem. 21, 248 (1972). 324. M. H. Lee and C. R. Dawson, JBC 248, 6596 (1973). 325. A. Marchesini, BBA (in press). 326. T. Nakamura, N. Makino, and Y . Ogura, J. Biochem. ( T o k y o ) 64, 189 (1968). 327. S. Osaki and 0. Walaas, ABB 123, 638 (1968). 328. B. Reinhammar, BBA 205, 35 (1970). 329. G. Fihraeus and B. Reinhammar, Acta Chem. Scund. 21, 2367 (1967). 330. M. Jonsson, E. Pettersson, and B. Reinhammar, Acta Chem. Scund. 22, 2135 (1968). 331. E. J. Cohn, L. E. Strong, W. L. Hughes, Jr., D. L. Mulford, J. N. Ashworth, M. Melin, and H. L. Taylor, JACS 68, 459 (1946). 332. H. Bjarling, Voz Sung. 8, 641 (1963).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

561

TABLE X MOLECULAR WEIGHTSA N D CHEMICAL COMPOSITION OF THE BLUE OXIDASES

Enzyme Ascorbate oxidase Laccase Tree Fungal Ceruloplasmin

Molecular weight

Carbohydrate CU content content (wt-%) (atoms/molecule)

Ref.

132,000-140,000

10

8-10

3,%$,326

110,000-140,000 64,000 135,000-160,000

45 10 8

4-6 4 6-8

328,339 389 333,340

(333,334) are also often used. According to Ryd6n (335) it is important to use fresh serum if artificial subunit formation by proteolysis is t o be avoided. Several procedures for the crystallization of ceruloplasmin have been described (333,336,33'7), and preliminary X-ray crystallographic data have been published (338). Investigations of the blue oxidases have generally been centered around the prosthetic copper ions and their role in the catalytic function (Sections IV,C-E) and only limited information is available on other molecular properties. Data on molecular weight, carbohydrate content, and estimated number of copper atoms per molecule are summarized in Table X (324,32S,S28,S29,S,3S9,340). All the enzymes are seen to be glycoproteins, but the carbohydrate moiety has not, in general, been characterized. An exception is ceruloplasmin, from which glycopeptides have been isolated and characterized (341,342).It was shown that the main molecular forms differ in carbohydrate composition. Such differences have also been suggested as the basis of laccase heterogeneity (330). Except for fungal laccase, the reported molecular weights vary considerably (Table X ) . As a consequence the values for the number of copper atoms per molecule are uncertain even when accurate copper analyses 333. 334. 335. 336.

H. F. Deutsch, ABB 89, 225 (1960). L. Broman and K. Kjellin, BBA 82, 101 (1964). L. RydCn, FEBS (Fed. Eur. Biochem. Soc.) Lett. 18,321 (1971). A. G. Morell, C. J. A . van den Hamer, and I. H. Scheinberg, JBC 244, 3494

(1969). 337. J. A . P. Trip and J . van Dam, Clin. Chim. Acta 36, 561 (1972). 338. B. S. Magdoff-Fairchild, F. M. Lovell, and B. W. Low, JBC 244, 3497 (1969). 339. T. Omura, J. Biockem. (Tokyo) 50,264 (1961). 340. L. RydCn, Eur. J. Biochem. 26, 380 (1972). 341. L. RydBn, Int. J. Protein Res. 3, 191 (1971). 342. L. RydCn, Acta Univ. Upsal. N o . 222 (1972).

562

B. G. MALMSTR~M, L.-E. ANDR~~ASSON,AND B. REINHAMMAR

are available. On the other hand, the range of these values is larger than can be accounted for by the uncertainties in the molecular weight. Ceruloplasmin has been found to have a loosely bound, “chelexable” copper (343), and EPR studies have shown that the content of so-called Type 2 Cu2+can vary between one and two ions per molecule (see 34.4).The variation in the copper content of ascorbate oxidase (345) may have a similar explanation (cf. 346). If the protein preparations are homogeneous, considerations based on the contents of the various forms of copper (see Section IV,C) suggest that the number of copper atoms per molecule is four for the laccases (328,3479, seven for ceruloplasmin (344), and eight for ascorbate oxidase (346). The amino acid composition has been determined for ascorbate oxidase (324,348), for fungal (329) and tree laccase (328), and for human ceruloplasmin (342,949).Since the content of some amino acids is of importance in considerations of subunit structure and of metal coordination, a summary of the analytical results are given in Table XI. The quaternary structure of the blue oxidases is of interest in relation to the occurrence of three distinct forms of copper (Section IV,C) . Ascorbate oxidase can be dissociated into two subunits by a variety of treatments (350). The subunits appear identical by several criteria, but the presence of three Type 1 Cu2+and only one Type 2 Cu2+is not consistent with a dimer structure. The formation of subunits at alkaline pH has been reported for ceruloplasmin (351) and fungal laccase (366),but it has been pointed out (350) that disulfide bonds are unstable under the conditions used. Evidence that ceruloplasmin consists of a single polypeptide chain has been reported (340).As in the case of ascorbate oxidase, the presence of a single Type 2 Cu2+in the laccases (347,353) and in ceruloplasmin (344) is inconsistent with a symmetrical dimer structure. I n addition, laccase appears to contain a single sulfhydryl group (328,354),in which case any possible subunits cannot be identical. D. J. McKee and E. Frieden, Biochemistry 10,3880 (1971). J. Deinum and T. VanngBrd, BBA 310, 321 (1973). M. H. Lee and C. R. Dawson, JBC 248,6603 (1973). J. Deinum, B. Reinhammar, and A. Marchesini, FEBS (Fed. Eur. Biochem. Soc.) Lett. 42, 241 (1974). 347. B. G. Malmstrom, B. Reinhammar, and T. Vanngbd, BBA 156,87 (1968). 348. G. R. Stark and C. R. Dawson, JBC 237, 712 (1962). 349. C. B. Kasper and H. F. Deutsch, JBC 238,2325 (1963). 350. K. G. Strothkamp and C. R. Dawson, Biochemistry 13, 434 (1974). 351. W. N. Poillon and A. G. Bearn, BBA 127,407 (1966). 352. J. J. Butaow, BBA 168, 490 (1968). 353. B. G. Malmstrom, B. Reinhammar, and T. VanngBrd, BBA 205, 48 (1970). 354. C. Briving and J. Deinum, FEBS (Fed. Eur. Biochem. Soc.) Lett. 51, 43 (1975).

343. 344. 345. 346.

8.

563

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

TABLE XI AMINOACID COMPOSITION OF BLUE OXIDASES Number of residues/moleculec Amino acid Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Half-cystine Methionine lsoleucine Leucine Tyrosine Phenylalanine Amide ammonia Lysine Histidine Arginine Tryptophan 4

Ascorbate oxidaseb

Fungal laccasec

Tree laccased

Ceruloplasmin#

132 70 73

79 41 35 27 42 43 57 35 6 4 30 35 15 31 55 6 17 14 4

60 50 37 37 38 36 39 43 7 11 31 34 24 30 79 26 17 13 6

121 73 62 117 48 76 50 64 15 18 54 72 65 48 68 44 42 22

100

92 94 76 79 18 17 72 91 50 53 (144) 55 39 37 50

Nearest integral value. From Stark and Dawson (348). From Fhhraeus and Reinhammar (329.) From Reinhammar (328). From R y d h (342).

C. THEFORMS OF COPPER: MAGNETIC AND SPECTROSCOPIC PROPERTIES 1. Definitions and Distribution of Copper Forms

The first indication that blue oxidases contain more than one form of copper came from quantitative EPR measurements with ceruloplasmin and fungal laccase (355).These showed that in both proteins only about half of the copper contributes to the EPR spectrum. The EPR-nondetectable part was later shown not to be paramagnetic by magnetic susceptibility measurements (356-358). 355. L. Broman, B. G. Malmstrom, R. Aasa, and T. Viinngbrd, JMB 5, 301 (1962). 356. A. Ehrenberg, B. G. Malmstrom, L. Broman, and R. Mosbach, J M B 5, 450 (1962). 357. P. Aisen, S. H. Koenig, and H. R. Lilienthal, J M B 28,225 (1967). 358. T. H. Mom and T. Viinngbrd, BBA 371, 39 (1974).

564

B. G. MALMSTR~M,L.-E. ANDRI~ASSON, AND B. REINHAMMAR

It was observed early (365) in EPR studies of the blue oxidases that there is a superposition of spectra from Cu2+in two different environments. Quantitative work on fungal laccase (347) revealed that in one protein molecule there is one ion in each environment, these being called Type 1 and Type 2 Cus+, respectively. Type 1 Cu2+is characterized by an unusually small hyperfine splitting constant, and it is also responsible for the anomalously strong blue color of the enzymes (Section IV,C,2). The spectral properties of Type 2 Cu2+,on the other hand, are more normal, and this ion was at first believed to represent Cu2+in denatured molecules (359). It was, however, later shown to be indispensable for catalytic activity (360,361).It also appears to be the binding site for certain inhibitors (Section IV,C,3). Since the most common nonparamagnetic form of copper is Cu+, the EPR-nondetectable copper ions, here called Type 3, were for a long time thought to be in this valence state (362).The situation changed, however, when i t was found that a molecule of laccase under anaerobic conditions can accept as many electrons as there are copper ions (363).The electrons not involved in the reduction of Type 1 and Type 2 Cu2+were found to reduce a cooperative two-electron acceptor associated with an absorption band a t 330 nm (364). The presence of such a band in the blue oxidases had earlier been stressed by Nakamura and Ogura (365).Various models for Type 3 copper have been proposed (Section IV,C,4). Early studies by the Japanese workers indicated that tree laccase does not conform to the pattern just described since all copper was thought to be paramagnetic (366,367). More recently, Makino and Ogura (368) have, however, confirmed that the blue color and the 330-nm band are associated with different copper ions. In addition, a careful E P R investigation of tree laccase has clearly demonstrated the presence of three types of copper (353).Since ascorbate oxidase has now also been found to have three types of copper (345,346),the occurrence of these three forms of the metal has been established as a general property of the blue oxidases. 359. B.G.Malmstrom, R. Aasa, and T. Vinngkd, BBA 110, 431 (1964). 360. R. Malkin, B. G. Malmstrom, and T. Vanngbrd, FEBS (Fed. Eur. Biochem. Soc.) L e t t . 1, 50 (1968). 361. R. Malkin, B. G. Malmstrom, and T. Vanngbrd, Eur. J . Biochem. 7 , 253 (1969). 362. B. G. Malmstrom, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 207. Wiley, New York, 1965. 363.J. A. Fee, R. Malkin, B. G. Malmstrom, and T. Vanngbrd, JBC 244, 4200 ( 1969). 364. R. Malkin, B. G. Malmstriim, and T. Vanngbrd, Eur. J. Biochem. 10, 324 (1969). 365. T.Nakamura and Y. Ogura, J . Biochem. (Tokyo) 59,449 (1966). 366. T.Nakamura, BBA 30, 640 (1958). 367. T. Nakamura, A. Ikai, and Y. Ogura, J. Biochem. (Tokyo) 57, 808 (1965). 368. N. Makino and Y. Ogura, J . Biochem. (Tokyo) 69,91 (1971).

8.

COPPER-CONTAINING

OXIDASES AND SUPEROXIDE DISMUTASE

DISTRIBUTION OF

THE

565

TABLE XI1 FORMS OF COPPERI N BLUE OXIDASES Number of ions/moleeule

Enzyme Ascorbate oxidase Laccase Tree Fungal Ceruloplasmin

Type 1 Cu2+ Type 2 Cu2+ Type 3 copper

Ref.

3

1

4

346

1 1 2

1 1 1

2 2 4

363 347 344

The enzymes differ, however, in the number of ions of each type present, as summarized in Table XII. Before a more detailed discussion of the properties of the various forms of copper is presented, it should be mentioned that one common approach in the study of metalloenzymes, namely, the preparation of apoenzyme followed by reactivation with various metal ions, appears difficult to apply to most blue oxidases. It is true that Tissihres (369) as early as 1948 reported a partial reactivation of cyanide-treated tree laccase on the addition of Cu2+,but he presented only activity measurements and no copper analyses. Later, Omura (370) found that, he could get partial reactivation with Cut but not with Cu”, and this was confirmed by Ando (371).The best reactivations were achieved when not all copper had been removed (370,371). A study of the spectral properties of the “apoenzymes” (370,371) reveals that good reconstitution was, in fact, obtained only when a considerable amount of Type 1 Cu2+had been retained. This is consistent with the finding that with fungal laccase only Type 2 Cu2+ can be reversibly removed (361). With ceruloplasmin, on the other hand, it appears possible to obtain good reconstitution if the apoprotein is made by treatment with sodium diethyldithiocarbamate in the presence of ascorbate (372,373). The same method fails, however, when applied to fungal laccase (374). A reversible dissociation of copper has been reported for ascorbate oxidase (375), but, as with tree laccase, good reactivation is achieved only on partial removal of the metal. 369. 370. 371. 372.

A. TissiBres, Nature (London) 162, 340 (1948). T. Omura, J. Biochem. (Tokyo) 50, 389 (1961). K. Ando, J. Biochem. ( T o k y o ) 68, 501 (1970). A. G. Morell, P. Aisen, W. E. Blumberg, and I. H. Scheinberg, JBC 239, 1042

(1964). 373. P. Aisen and A. G. Morell, JBC 240, 1974 (1965). 374. P. Aisen, unpublished results. 375. Z. Penton and C . R. Dawson, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 222. Wiley, New York, 1965.

566

B. G. MALMSTROM, L.-E. A N D R ~ S S O N , AND B. REINHAMMAR

The properties of the three types of copper in all the blue oxidases show great similarities. Consequently, an extensive discussion of individual enzymes would appear unnecessary, particularly since detailed compilations of data for all except ascorbate oxidase are available in two reviews (10,Il). 2. Type 1 Copper a. Optical, ORD, and CD Spectra. Type 1 CuZcis responsible for the beautiful, strong color of the blue oxidases. I n all the enzymes the dominating absorption band in the visible region is slightly above 600 nm. The molar extinction coefficient a t the absorption maximum is almost two orders of magnitude larger than that commonly found for Cuz+complexes (see 10). It is most easily estimated for the laccases, which have only one Type 1 Cu2+.For the tree enzyme it is 5700 M-' cm-l (365) and for the fungal enzyme 4900 M-' cm-' (361). If i t is assumed that the two and three Type 1 Cu2+ions found in ceruloplasmin and ascorbate oxidase, respectively, are identical, then the extinction coefficients per Type 1 Cu2+are 4400 (376) and 3300, M-' cm-l (346). There is also an unresolved band with lower extinction coefficient between 700 and 800 nm in all the proteins. Possible absorption bands a t shorter wavelengths are obscured because of an overlap with the broad 330-nm band associated with Type 3 copper. There are, however, a number of blue electron transport proteins, such as azurin and stellacyanin, which have a single Type 1 Cuz+,and in these a band a t 450 nm is clearly resolved. Circular dichroism spectra (377,378) reveal the presence of optical transitions a t this wavelength in all the blue oxidases as well, and this band has also been observed by ORD measurements (379). In view of the unusual optical properties, the electronic structure of Type 1 Cu2+has been discussed extensively (377). The high extinction coefficients make it unlikely that d-d transitions are involved. Indeed, since the number of bands seen in the CD spectra often are greater than four, charge transfer and possibly also ligand transitions must be considered (see 377). On the basis of resonance Raman spectroscopy (380), it has been suggested that the charge transfer involves sulfur. It is noteworthy that all blue oxidases have one cysteine (Section IV,B), and that 376. W. E. Blumberg, J. Eisinger, P. Aisen, A. G. Morell, and I. H. Scheinberg, JBC 238, 1675 (1963). 377. K.-E. Falk and B. Reinhammar, BBA 285,84 (1972). 378. S.-P. W. Tang, J. E. Coleman, and Y. P. Myer, JBC 243, 4286 (1968). 379. F. Bossa, G. Rotilio, P. Fasella, and B. G. Malmstrom, Eur. J . Biochem. 10, 395 (1969).

380. V. Miskowski, $.-P. W. Tang, T. G. Spiro, E. Shapiro, and T. H. Moss, Biochemistry 14, 1244 (1975).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

567

this is an invariant residue in the sequences of several azurins (581). With the related protein, stellacyanin, chemical evidence for the involvement of a sulfhydryl group in copper coordination has also been obtained (382). The spectral properties have been interpreted in terms of fivecoordination (380,383).Unfortunately, the crystal structure has not been determined for any blue protein, even if preliminary X-ray data have been published for ceruloplasmin (338) and azurin (384). b. EPR Spectra. Malmstrom and Vannghrd (254) were the first to observe that fungal laccase and ceruloplasmin have unusually low g values and hyperfine splitting constants in their E P R spectra. The same was later found for tree laccase (353) and ascorbate oxidase (545,346).An illustrative spectrum is shown in Fig. 1. Since the ascorbate oxidase used has three Type 1 Cu2+and only one Type 2 Cu2+,the former dominates the spectrum. The good resolution indicates that all three ions have the same spectrum, which may justify the assumption made earlier that they have the same extinction coefficient (Section IV,C,2,a). Good simulations can be obtained if the symmetry is assumed to be less than axial (Fig. 1). The same is true for tree laccase (355), while the 9 GHz spectra of fungal laccase and ceruloplasmin (254) indicate a more axial symmetry. If the spectra are recorded a t 35 GHz, however, a deviation from axial symmetry is observed with these enzymes as well (347’).With ceruloplasmin simulations have usually been made on the assumption that both Type 1 Cu2+have the same parameters (3&,385), but a better result may be obtained by using a slight difference in hyperfine coupling (386). A detailed discussion of the E P R properties of the blue oxidases is available (11). 3. T y p e 2 C o p p e r

The EPR parameters associated with Type 2 Cu2+in all blue oxidases are similar to those generally found for simple Cu2+complexes. A typical example is seen in the simulated spectrum of Fig. 1. Because the Type 2 spectrum is broader than that of Type 1 Cu2+,the low-field line is entirely associated with Type 2 Cu2+.This line is, therefore, generally used in quantitative EPR studies of this ion. So far i t has not been possible 381. R. P. Ambler, in “Chemotaxonomy and Serotaxonomy” (J. G. Hawkes, ed.), p. 57. Academic Press, New York, 1968. 382. L. Morpurgo, A. Finazzi Agrb, G. Rotilio, and B. Mondovi, BBA 271, 292 (1972). 383. H. B. Gray, Advan. Chem. Ser. 100, 365 (1971). 384. G. Strahs, Science 165, 3888 (1969). 385. L.-E. Andreasson and T. VanngLd, BBA 200, 247 (1970). 386. P.-0. Gunnarsson, U. NylCn, and G. Pettersson, Eur. J. Biochem. 37, 47 (1973).

568

B. G. MALMSTROM, L.-E. A N D R ~ S S O N , AND B. REINHAMMAR

hcurbita pep0 msdulloso kscorbate oxidase

I

1

b Simulated

I

J

2 500

3000

3500

Mognetic field ( g a u s s )

FIQ.1. Experimental (a) and simulated (b) EPR spectrum of ascorbate oxidase from green zucchini at about 9 GHz and 77°K.The simulated spectrum (b) is the sum of two components, Type 1 (---) and Type 2 Cua+ (- -1, with relative intensities of 3 and 1, respectively. Lorensian line shape was assumed. The parameters used in the simulations and the experimental conditions are given hy Deinum et al. (346).

-

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

569

to associate any part of the visible and ultraviolet spectrum with Type 2 Cu2+;for example, on removal of this ion the absorbance around 600 nm remains unchanged (361) . The most characteristic property of Type 2 Cu2+is its ability to interact strongly with a number of anionic inhibitors, which include not only classic metal poisons such as N,- and CN- but also the halides. The inhibition by these agents had been studied considerably before their binding to a specific Cu2+ was known, particularly with ceruloplasmin (see 387) , and in some cases characteristic spectral changes were also observed (388). In 1970, Andr6asson and Vannglird (385) showed with EPR that the spectral changes and inhibition occurring at low concentrations of azide are entirely correlated with binding to Type 2 Cu2+;a t higher concentrations more complex changes, involving other copper ions as well, occur (377,389). The fact that the inhibitor F- binds to Type 2 Cu2+in the blue oxidases, as seen from changes in the low-field EPR line (S90), is particularly interesting, since this ion is generally not considered a good ligand for Cu2+.Fungal laccase is most exceptional in this respect since the stability constant for the Type 2 Cu2+-F-complex is greater than 10 pM-I (390), compared to less than 10 M-I for the Cu2+-aquoion. Because of this remarkable affinity the enzyme as commonly prepared has more than 15% of its molecules contaminated with F- (36S), and it has been necessary to develop a special procedure for its removal (391).Tree laccase also shows a considerable affinity for F-, while the binding to Type 2 Cu2+ in ceruloplasmin is much weaker (390). The interaction between Type 2 Cuz+ and F- is most clearly seen in EPR spectra, since the low-field line is split into a doublet because of a coupling of the unpaired electron to the nucleus ISF,which has a nuclear spin with I = 4 (S60). A spectral change around 320 nm is, however, also associated with this interaction in the laccases and ceruloplasmin (390). While this change is definitely related to the binding of the inhibitor to Type 2 C$+, it does not necessarily reflect a change in the absorption properties of this ion. The reason for this is that in the Type 2 Cu2+-F- complex, the properties of Type 3 copper are also affected 387. G. Curzon and B. E. Speyer, BJ 105, 243 (1967). 388. C. B. Kasper, JBC 243, 3218 (1968). 389. W. Byers, G. Curzon, K. Garbett, B. E. Speyer, S. N. Young, and R. J. P. Williams, BBA 310, 38 (1973). 390. R. Briindkn, B. G. Malmstrom, and T. Vanngird, Eur. J. Biochem. 36, 195 (1973). 391. R. Brand&, B. G . Malmstrom, and T. Vanngird, Eur. J. Biochem. 18, 238 (1971).

570

B. G. MALMSTROM,

L.-E.

ANDREASSON, AND

B. REINHAMMAR

(288,364,392). Concomitant changes in the absorption around 600 nm on the binding of inhibitors (390) should be interpreted with caution since Type 1 Cu2+may undergo reduction by endogenous reducing agents when the enzymes are inhibited. This has been clearly demonstrated in the case of fungal laccase (393). When fungal laccase is treated with CN-, drastic but reversible changes in the E P R spectrum occur (347,360). The intensity is reduced to onehalf and extensive superhyperfine structure resulting from N develops. Since these changes are not found with samples from which Type 2 Cu2+ has been removed (360,36l), the remaining spectrum probably results from this ion. Experiments with isotopes (360) show that the N hyperfine structure is not caused by CN- but by protein ligands. These results show that Type 2 Cu2+is coordinated to three to four N, at least in the CNcomplex. With ceruloplasmin N y l h and Pettersson (394) have shown that on chemical modification of histidine residues, the EPR spectrum of Type 2 Cu2+,but not that of Type 1, changes. It is not, however, possible to draw any conclusion about Cu2+ligands from their experiments. The effects of inhibitor binding to Type 2 Cu2+on the oxidation-reduction and catalytic properties of the blue oxidases will be considered later (Sections IV,D,2 and IV,E). Type 2 Cuz+can form a complex with H202, and this compound has characteristic spectral properties (390; cf. Section IV,E,3).

4. T y p e 3 Copper

As already mentioned (Section IV,C,l), the EPR-nondetectable copper ions (Type 3) of the blue oxidases were long thought to be in the Cu+ state. Anaerobic redox titrations of fungal laccase (363,364) showed, however, the presence of a cooperative two-electron acceptor which can be monitored a t 330 nm. For three reasons this was believed to involve the Type 3 copper ions (10):the stoichiometry, the high oxidation-reduction potential (Section IV,D), and the 330-nm absorption, not found in simple copper proteins. A reducible center not involving Type 1 or Type 2 Cuz+ but having an absorption band a t 330 nm has now also been demonstrated in tree laccase (287,368) and ceruloplasmin (395). Ascorbate oxidase also has 392. L.-E. AndrCasson, B. G. Malmstrom, C. Stromberg, and T. Vinnglird, Eur. J . Bbchem. 34, 434 (1973). 393. J. A. Fee, B. G. Malmstrom, and T. Vannglird, BBA 197, 136 (1970). 394. U. Nylkn and G. Pettersson, Eur. J. Biochem. 27, 578 (1972). 395. R. J. Carrico, B. G. Malmstrom, and T.Vanng%rd,Eur. J . Biochem. 20, 518 (1971).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

571

Type 3 copper and a 330-nm band (345,346). Makino and Ogura (568) implied a discrepancy in the results for fungal and tree laccase, since in anaerobic titrations the absorbancies at 610 and 330 nm decrease together in the fungal (364) but not in the tree (368) enzyme. This is explained, however, by differences in the relative oxidation-reduction potentials of the two enzymes (287,288; Section IV,D). With ceruloplasmin many earlier results suggested that each molecule can accept half an electron per copper only (376,396,397). This discrepancy might be explained by a disappearance of the 330-nm band without reduction under certain circumstances (344). I n view of the lack of paramagnetism (356-358), the only possibilities to be considered for the state of Type 3 copper is Cu+, Cu3+,or an antiferromagnetically coupled Cuz+-Cu2+pair. Cu3+would appear unlikely (see 10 and 358). Williams and co-workers (389) have suggested the following structure:

[I]:I:

Against this proposal is failure to detect an increase in the number of sulfhydryl groups on complete reduction (398). For this reason, among others, the Cuz+-Cu2+model is preferred, but it is not possible a t this stage to make a firm choice among possible models. It is interesting in this connection that on treatment of reduced ceruloplasmin with NO, an EPR spectrum ascribed to Cu-Cu dimers (399) is obtained (4OO).

D. OXIDATION-REDUCTION PROPERTIES It is natural that the oxidation-reduction properties of the different reducible centers in the blue oxidases have attracted a good deal of attention. In general, two types of experiments have been used. One type is anaerobic reductive titrations with a reducing agent having a lower potential than any of the electron acceptors in the enzyme. Suc,h experiments can yield information about differences in midpoint potentials of the different acceptors but do not provide absolute potential values. The second type of experiments, namely, direct potentiometric titrations with 396. C . B. Laurell, in “The Plasma Proteins” (F. W. Putnam, ed.), Vol. 1, p. 349. Academic Press, New York, 1960. 397. B. F. van Gelder and A. Veldsema, BBA 130,267 (1966). 398. J. Deinum, unpublished results; see also ref. 36.4. 399. A. J. M. Schoot Uiterkamp, FEBS (Fed. Eur. Biochem. Soc.) Lett. u),93 (1972). 400. F. X. R . van Leeuwen, R. Wever, and B. F. van Gelder, BBA 315,200 (1973).

572

B. G. MALMSTROM, L.-E. ANDREASSON, AND B. REINHAMMAR

TABLE XI11 OXIDATION-REDUCTION POTENTIALS OF LACCASE

A N D CERULOPLASMIN

Oxidation-reduction potential (mV) Enzyme

PH

Type 1 copper

Type 3 copper

Ref.

Tree laccase +10 mM NaF Fungal laccase +1 mil4 NaF Ceruloplasmin

7.5 7.5 5.5 5.5 5.5

394 390 785 780 490,580

434 390 782 570

288

-

288

344

electrodes or with redox buffers, is intended to give such absolute values. Early studies (397,401,402) were made before the presence of three separate reducible centers had been established. They generally employed the absorbance around 600 nm to monitor the redox state and thus, a t best, provide information about the Type 1 copper only. I n many cases the results could be misinterpreted, however ; for example, in the absence of other information, the reductive titrations of fungal laccase (363) could be taken as evidence that this enzyme contains a t least three Type 1 coppers. Consequently, the present discussion will be limited to the few studies in which both the 610- and 330-nm bands have been employed. These deal with the laccases (287,288,364,368) and ceruloplasmin (344,396,403,404),while the redox properties of ascorbate oxidase have not been investigated. With all three oxidases studied it has been established that each molecule can accept as many electrons as it has copper ions (288,364,396), but with ceruloplasmin misleading results can be obtained if only the optical bands of the protein are measured (344,403). Tree laccase has been found to interact with ferrocyanide (688), so that experiments using this reducing agent (401) do not give correct results. What appears to be the best values available for the oxidation-reduction potentials of Type 1 copper and the two-electron acceptor (Type 3 copper) are summarized in Table XIII. The potential for Type 2 copper has not been determined directly since EPR measurements must be employed and these are disturbed by the mediators used. From the reductive titrations it can, however, be seen to have a potential close to Type 1 copper in the two laccases and ceruloplasmin (see 287). For ceruloplasmin, which 401. T. Nakamura, BBA 30, 44 (1958). 402. J. A. Fee and B. G . Malmstrom, BBA 153,299 (1968). 403. A. Veldsema, Ph.D. Thesis, Drukkerij Elinkwijk, Utrecht, 1973. 404. A. Veldsema and B. F. van Gelder, BBA 293,322 (1973).

8. COPPER-CONTAINING OXIDASES

AND SUPEROXIDE DISMUTASE

573

has two Type 1 Cu2+,two potentials are given for this acceptor. These are calculated on the assumptions that both ions have the same extinction coefficient and that there is no interaction between them. This is not a unique interpretation, however, and an equally good fit with the experimental results is obtained on the assumption of identical acceptors having a negative cooperativity with a free-energy change corresponding to 60 mV (344; cf. 405). As already mentioned, the 330-nm band is not simply related to the redox state of Type 3 copper in ceruloplasmin; thus, the potential of this acceptor cannot be obtained by the method used. From redox titration data, Veldsema and van Gelder (404) concluded that ceruloplasmin contains one Type 1 and three Type 2 copper ions. Later EPR studies from the same laboratory (406) showed, however, the same contents as reported by other investigators (344,385; see Table XIII) . The almost linear decrease in 610 nm absorption up to full reduction in the anaerobic titrations of fungal laccase (363), which was puzzling in view of the presence of only one Type 1 copper, is explained by the fact that Type 1 and Type 3 copper have nearly the same potential in this enzyme (Table XIII). I n the presence of F- they titrate separately (36‘41, since this ion lowers the potential for Type 3 copper. I n tree laccase F- has the same effect, but in this the different potentials in the native enzyme become the same in the inhibited complex. Thus, in this case the titration becomes linear in the presence of F- (287). It should be recalled that F- is bound to Type 2 Cu2+ (Section IV,C,3), so there is apparently an interaction between this center and the two-electron acceptor. The potential of Type 1 copper, on the other hand, is not affected by F- (Table XIII), Type 1 Cu2+in fungal laccase has been found to be reduced without the addition of a reducing agent if the p H is raised above 6 (359,393). This process is associated with the dissociation of one proton from a group with an apprent pK of 7.45. I n view of the extremely high potential of Type 1 copper in this enzyme, the possibility that H,O is the reducing agent is thermodynamically feasible and has been considered (407). Later studies showed, however, that O2 is consumed rather than produced in the process (393).Apparently an endogenous reducing agent, possibly denatured enzyme, is responsible for the reduction. Since it is relatively 405. B. G. Malmstrom, Quart. Rev. Biophys. 6, 384 (1973). This paper gives an extensive discussion of interactions in redox proteins with several electron-accepting sites. 406. R. Wever, F. X. R. van Leeuwen, and B. F. van Gelder, BBA 302, 236 (1973). 407. J. A. Fee, B. G. Malmstrom, and T. VanngBrd, in “Biochemie des Sauerstoffa” (B. Hem and Hj. Staudinger, eds.), p. 29. Springer-Verlag, Berlin and New York, 1968.

574

B. G. MALMSTRbM, L.-E. ANDRkASSON, AND B. REINHAMMAR

slow, it is not seen a t low pH where reoxidation is rapid. A likely explanation is that OH-, like F-, becomes bound to Type 2 Cuz+and that this prevents reoxidation by inhibiting electron transfer to the two-electron acceptor (see 392). The finding that the two-electron acceptor remains oxidized in the pH-reduced enzyme (36‘4,393) is consistent with this view. I n such a case the pK of 7.45 would represent the hydrolysis of Type 2 cu2+.

E. CATALYTIC PROPERTIES 1. Specificity, Inhibition, and Steady-State Kinetics

The blue oxidases have a rather low degree of specificity. Before 1960 ascorbate oxidase was believed to be quite a specific enzyme (SO), but later studies revealed that many phenolic substances are quite good substrates (408). Laccase has long been known to catalyze the oxidation of various diphenols and diamines (49).Monophenols, on the other hand, were not considered substrates, and this was generally one of the criteria used to distinguish laccase from tyrosinase. With the fungal enzyme FBhraeus (410) showed, however, that the failure to detect monophenol oxidation resulted from a rapid reaction inactivation, which could be prevented by the addition of detergents. Later i t was shown that monophenols are substrates for three laccase as well ( d l 1) . The products of laccase-catalyzed oxidation of monophenols are quite distinct from those in the case of the tyrosinase reaction. With laccase the enzymic reaction leads to the formation of free radicals (see Section IV,E,2a) which then undergo further nonenzymic reactions, while tyrosinase appears to catalyze a two-equivalent oxidation without free radical intermediates (412). Ceruloplasmin was shown by its discoverers t o have a specificity similar to that of laccase (even if the turnover numbers are about two orders of magnitude lower with all substrates, see 419) and the enzyme has, in fact, been referred to as blood laccase (309). Ascorbate as a possible ceruloplasmin substrate has been a subject of considerable debate. Curzon 408. C. R. Dawson, ipa “The Biochemistry of Copper” (J. Peisach, P. Aisen, and W. E. Blumberg, eds.), p. 305. Academic Prees, New York, 1966. 409. W. G. Levine, in “The Biochemistry of Copper” (J. Peisach, P. Aispn, and W. E. Blumberg, eds.), p. 371. Academic Press, New York, 1966. 410. G. Fihrseus and H. Ljunggren, BBA 48,22 (1961). 411. G. Benfield, S. M. Bocks, K. Bromley, and B. R. Brown, Phytochemistry 3, 79 (1964). 412. H . S. Mason, E. Spencer, and I . Yamazaki, BBRC 4, 236 (1961). 413. J. Peisach and W. G. Levine, JBC 240,2284 (1965).

8.

COPPER-CONTAINING

OXIDASES AND SUPEROXIDE DISMUTASE

575

and O’Reilly (414) had shown that Fez+is a substrate for ceruloplasmin, and this suggested that iron can mediate the oxidation of any compound which can reduce Fe8+.Morell et al. (415) reported that in the presence of EDTA ascorbate is not oxidized. More recently, Curzon and Young (416) have, however, reinvestigated the question and shown that ascorbate appears to be a true, albeit poor, ceruloplasmin substrate. In recent years, Frieden and co-workers (319,417) have emphasized Fez+as a substrate (see Section IV,E,2,b). It has already been mentioned that many anions inhibit the blue oxidases by binding to Type 2 Cu2+ (see Section IV,C,3). In many cases the situation is more complex, however, and other inhibition mechanisms appear involved; for example, the Type 2 CuZ+-azidecomplex in ceruloplasmin has a stability constant of 15 mM-l (385),but a much stronger binding (500 mM-l) occurs to the reduced enzyme (418). Consequently, the strength of inhibition in this case, as with CN-, is dependent on the redox state of the enzyme. Other inhibitors (for example, the halides and certain carboxylic acids) act on both the oxidized and the reduced ceruloplasmin (419). The steady-state kinetics of the blue oxidases, particularly ceruloplasmin, has been studied a good deal, mostly with dimethyl-p-phenylenediamine as substrate. Apart from defining maximum turnover rates and inhibition constants, such measurements have, however, not provided much information about reaction mechanisms, which has mainly been derived from transient-state and spectroscopic studies (Section II,E,2). Because of this only a few representative recent steady-state investigations will be quoted (@O-QSS). Despite the undoubtedly complex mechanism, at constant pH and oxygen concentration Michaelis-Menten kinetics is followed, deviations being caused by the free radical product (Section IV,E,2,a). The maximum velocities with the best substrates are highest for fungal laccase (of the order of 100 sec-l) and lowest for ceruloplasmin ( <1 sec-l) . 2. The Reducing Substrates a. Free Radical Formation. Oxidation of the reducing substrate appears, with all the blue oxidases, to involve a one-electron transfer, result414. 415. 416. 417. 418. 419, 420. 421. 422.

G. Curzon and S. O’Reilly, BBRC 2, 284 (1960). A. G. Morell, P. Aisen, and I. H. Scheinberg, JBC 237, 3455 (1962). G. Curzon and S. N. Young, BBA 268, 41 (1972). S. Osaki, D. A. Johnson, and E. Frieden, JBC 246,3018 (1971). P.-0. Gunnarsson, U. Nylkn, and G. Pettersson, Eur. J. Biochem. 27, 572 (1972). P.-0. Gunnarsson and G. Pettersson, Eur. J . Biochem. 27, 564 (1972). G. Pettersson and I . Pettersson, Acta Chem. Scand. 23, 3235 (1969). G. Pettersson, Acta Chem. Scand. 24, 1809 (1970). S. N. Young and G. Curzon, BJ 129,273 (1972).

576

B. G. MALMSTROM, L.-E.

ANDREASSON,AND

B. REINHAMMAR

ing in the formation of a free radical which then dismutates spontaneously. Free radical intermediates can be observed optically but are most clearly demonstrated by EPR spectroscopy. With this technique, Nakamura (493) in 1960 found a radical intermediate in the reaction catalyzed by tree laccase, and similar observations were later made with ascorbate oxidase (424), fungal laccase ( 4 2 5 ) ,and ceruloplasmin (425). Because of the nonenzymic dismutation reaction, free radicals may form in a side reaction and thus not represent intermediates, as shown with tyrosinase (412). In all the cases quoted, however, kinetic data showed that the radical is a primary product of the reaction between enzyme and substrate. If the initial electron acceptor is a single Cu2+, radical formation would appear mandatory since this ion must be reduced to Cu+in a one-electron step. With ceruloplasmin the radical formed from dimethyl-p-phenylenediamine (Wurster's red) has been found (420) to be not only a product but also a substrate in a second one-electron step. This finding apparently accounts for the deviation from Michaelis-Menten kinetics found in some cases (420,429). b. The Ferroxidase Activity. It has already been mentioned repeatedly (Sections IV,A,2 and IV,E,l) that Fez+is a good substrate for ceruloplasmin. It should be noted, however, that the turnover rate (426') is not much higher than with organic substrates. The large difference is instead found in the rate of reduction of Type 1 Cu2+,which appears to be the primary electron acceptor in ceruloplasmin (426,427,42628) as in all blue oxidases (10). This has a second-order rate constant of about los M-I sec-I with Fez+ compared to about lo3 M-l sec-l with the best organic substrates (427).The reason that this large reduction rate does not lead to a much higher catalytic velocity is that other steps are rate-limiting during turnover. Thus, the rate of reoxidation of Type 1 Cut is low, as seen from the steady-state level which corresponds to about 95% reduction of Type 1 Cu2+ (428). The rate-limiting step has been reported to have a first-order rate constant of about 1 sec-l (427).Osaki and Walaas (427) suggested that it involves a change in protein conformation, but another possibility is a slow intramolecular electron transfer step (428). c. Anaerobic Reduction Experiments. Attempts to determine the kinetic properties of steps involving the reduction of electron acceptors in 423. T. Nakamura, BBRC 2, I11 (1960). 424. I. Yamaaaki and L. H. Piette, BBA 50, 62 (1961). 425. L. Broman, B. G. Malmstrom, R. Aasa, and T. Vanngbrd, BBA 75, 365 (1963). 426. S. Osaki, JBC 241,5053 (1966). 427. S.'Osaki and 0. Walaas, JBC 242,2653 (1967) 428. R. J. Carrico, B. G. Malmstrom, and T. Vanngbrd, Eur. J . Biochem. 22, 127 (1971).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

577

oxidases are generally based on anaerobic experiments. Many studies of the anaerobic reduction of the blue oxidases have been performed, but before the recognition of the two-electron acceptor and the 330-nm band (Section IV,C,4) the results were difficult to interpret. For this reason the present section will be limited to recent investigations (392,403,428-493).

The presence, in a blue oxidase, of electron acceptors other than Type 1 and 2 Cu2+was first recognized for fungal laccase in anaerobic reduction experiments by Malmstrom et al. (429). These authors also showed that the initial electron acceptor is Type 1 Cuz*, which with good substrates is reduced in a reaction characterized by a second-order constant of about lo6 M-l sec-I, as confirmed in later, more complete studies (392,430). The two-electron acceptor is reduced much more slowly in an intramolecular step with a rate constant of 1 sec-*. An important question is if the reductive steps observed anaerobically are fast enough to form part of the catalytic mechanism. This is clearly the case for the reduction of Type 1 Cu2+,while the two-electron acceptor appears to react much too slowly in relation to the turnover, which is faster than 100 sec-’ (392). Despite this finding, Type 3 copper appears to participate in electron transfer reactions in the catalytic mechanism, however. This can be concluded, for example, from the fact th a t the steady state of the 330-nm chromophore corresponds to partial reduction (428). Since the chromophore can be both reduced and oxidized by the respective substrates, this is only explicable in terms of an electron transfer via i t during turnover. To overcome this apparent dilemma i t is necessary to postulate that the presence of oxygen alters the rate constants for intramolecular electron transfer steps. The same type of results have been obtained for cytochrome c oxidase (434). The disheartening conclusion is that anerobic experiments do not yield information of relevance to the catalytic reaction, except possibly for the rate of electron transfer from the reducing substrate to the initial acceptor in the enzyme. I n the presence of the inhibitor F- the internal electron transfer to 429. B. G. Malmstrom, A. Finazzi Agrb, and E. Antonini, Eur. .I. Biochem. 9, 383 (1969). 430. L.-E. Andrkasson, R. Brandkn, B. G. Malmstrom, C. Stromberg, and T. Vanngird, in “Oxidases and Related Redox Systems” (T. E. King, H. S. Mason, and M. Morrison, eds.), p. 87. Univ. Park Press, Baltimore, Maryland, 1973. 431. J. W. Dawson, H. B. Gray, R. A. Holwerda, and E. W. Westhead, Proe. Nut. Aead. Sei. U S . 69, 30 (1972). 432. I. Pecht and M. Faraggi, Nature (London), New Biol. 233, 116 (1971). 433. M. Faraggi and I. Pecht, JBC 248, 3146 (1973). 434. L.-E. AndrBasson, B. G. Malmstrom, C. Stromberg, and T. Vanngird, FEBS (Fed. Eur. Biochem. Soc.) Lett. 28,297 (1972).

578

B. G. MALMSTR~M,L.-E. A N D R ~ A ~ S O NAND , B. REINHAMMAR

the two-electron acceptor is even slower with a rate constant of 0.008 sec-l for fungal laccase (39%).The rate of reduction of Type 1 Cuz+,on the other hand, is not affected. Apparently, F- must inhibit the enzyme by preventing the reoxidation of Type 1 Cut and, thereby, also the oxidation of more substrate. The reduction of tree laccase by Cr2+has been studied (4.91). It appears to reduce the 330-nm chromophore faster than Type 1 CuZ+.The hydrated electron does not reduce Type 1 Cu2+in fungal laccase (43%)and ceruloplasmin (433) directly but via protein residues.

3. The Reduction of Oxygen a. Experimental Results. When blue oxidases in their fully reduced state are mixed with dioxygen, a rapid reoxidation takes place (468,4299). I n fact, the rate is larger than that estimated from the steady-stake levels in turnover experiments ( 4 2 8 ) .Apparently the rate of reaction of one center is dependent in the oxidation state of other centers, and again the transient-kinetics experiments do not yield results relevant to the catalytic reaction. An explanation could be that in the experiments just described all centers are reduced, while some may remain oxidized during catalysis in order to bind an intermediate such as 0,- or H202,in the reduction of dioxygen. In molecules having the Type 1 center reduced and the other ones oxidized, the blue color does not return rapidly on mixing with dioxygen (428,429).The rate of color return, i.e., return of Type 1 Cu2+,is limited by an intermolecular electron transfer between two partially reduced molecules (429).A likely explanation for this is that the 330-nm center cannot be reduced unless two reducing equivalents are available since it functions as a cooperative two-electron acceptor (Section IV,D) . A natural hypothesis is that the purpose of the two-electron acceptor is to facilitate two-electron steps in the reduction of dioxygen. If such steps occur, H,O, should be formed as an intermediate. Any possible intermediate must, however, remain ,firmly bound to the enzyme since H,O is the final reduction product. Recently, kinetic evidence for an intermediate in the reaction with dioxygen has been obtained with ceruloplasmin (436) and fungal laccase (436).It is interesting to note that these intermediates have spectral properties similar to those of an artificial H,02 complex of fungal laccase (391). 435. T. Manabe, N. Manabe, K. Hiromi, and H. Hatano, FEBS (Fed. Eur. Biochem. Soc.) Lett. 23, 268 (1972). 436. L.-E. AndrBasson, R. Brand&, B. G. Malmstrom, and T. VanngBrd, FEBS (Fed. Eur. Biochem. Soc.) Lett. 32, 187 (1973).

8.

COPPER-CONTAINING OXIDASES AND SUPEROXIDE DISMUTASE

579

b. General Considerations. The most difficult mechanistic problem with the blue oxidases, as with cytochrome c oxidase, would appear to be the coupling of the one-electron step in the oxidation of the reducing substrate to the four-equivalent reduction of dioxygen. It might be argued that this could easily occur via a radical chain mechanism involving oneelectron reactions only. Such a mechanism would seem unlikely for several reasons, however (see 1 0 ) . First, it must involve several quite distinct intermediates, which all must be bound strongly. The very reactive OH radical would, in particular, be an undesirable species. Second, a one-electron transfer t o dioxygen from the reduced form of a high-potential center is energetically unfavorable. From the oxidation-reduction potentials for the superoxide-dioxygen couple (140) and fungal laccase (688),the minimum activation energy for a one-electron mechanism can be estimated to be 106 kJ mole-I. In view of the considerations just given, it would seem desirable to formulate alternative mechanisms. Some authors have suggested that the presence of a t least four reducible centers in the blue oxidases, and in cytochrome c oxidase, might allow a concerted mechanism in which four electrons are donated to dioxygen in a single step. Many considerations, including the asymmetry of the oxidases (437), make this an unlikely alternative. The presence of a cooperative two-electron acceptor, and the kinetic results described in the previous section, make two double-electron transfers with H,O, as an intermediate much more attractive. It would, in fact, seem worthwhile to apply the lessons learned from the extensive mechanistic information available for the blue oxidases also to the oxidase, cytochrome c oxidase (cf. 405). 437. B. G. Malmstrijm, in “Symmetry and Function of Biological Systems a t the Macromolecular Level” (A. Engstrom and B. Strandberg, eds.), p. 153. Wiley (Interscience), New York, 1969.

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Author Index Numbers in parentheses are references numbers and indicate that an author’s work is referred to, although his name is not cited in the text.

A

Aleman, V., 325, 389, 391(286), 395(287),

Aasa, R., 7, 563,564, 573(359), 576 Ahbott, B. J., 533(152), 534 Abbott, M. T., 152, 154(145), 158, 170,

Aleo, J. J., 162 Alfano, J., 266, 269 Allison, M. J., 61 Allison, W. S., 77 Alvares, A. P., 270 Amadori, E., 284 Amaral, D., 531 Ambike, S. H., 280 Ambler, R. P., 567 Ameraal, R. M., 58(17), 59, 62(17), 99

533( 146), 534

171, 172, 178, 174, 175(145, 317, 319, 325, 328, 329), 176(316, 317, 319, 322, 3281, 177, 178, 179, 184, 185(145), 186, 187(145, 320, 325), 188(145) Abe, T., 484 Aheles, R. H., 199, 204(24-26), 428, 431, 451(31), 485(31), 486, 488(31), 489 (186), 490(31), 498 (17) Amick, R. M., 89 Abernethy, J. L., 557 Anan, F., 146 Abramovitz, A. S., 473 Andersen, R. D.. 60, 66, 70, 71(28, 65), Achee, F. M., 515(55), 516, 519 83,911(105), 92(126), 93(126), 447 Ackerman, E., 361, 368 Anderson, B. M., 428 Ackers, G. K., 445, 447(58) Anderson, R., 263 Adachi, K., 146 Adachi, O., 513, 514(43), 518(41, 43), 519, Anderson, R. E., 37 Ando, N., 267 520(79), 521(41, 79) Ando, W., 536, 565 Adachi, T., 142 Andrhasson, L.-E., 567, 569, 570, 573(385), Adams, E., 164, 189 574(392), 575(385), 577, 578 Adams, G. E., 358 Andres, R., 315, 389, 391(285), 407(285) Adams, M. J., 77 Andrews, L., 94, 99(131) Adams, W. C., 538,547(228) Addink, R., 112, 241, 242, 243(157), 251 Andrews, P., 306, 313, 314 Anger, G., 37 (157) Antoine, A. D., 403, 404(302) Adman, E. T., 32,33,37, 71,367 Aisen, P., 563, 565, 566, 571(357, 376), 575 Antonini, E., 447, 499, 500(202), 521, 577, 578 (429) Akeson, A., 150 Anttinen, H., 166 Akino, M., 239, 240(143), 290, 291(179) Apgar, P. A., 66, 70, 71(65), 83, 91(105), Al-Adnani, M. S., 162 447 Albrecht, S. L., 53 Appaji Rao, N., 225,247(88) Alden, R. A., 32, 33, 35(90-92) 581

AUTHOR INDEX

582 Appleby, C. A., 280 Appleby, M., 427, 428(26), 442(26), 464, 465(26), 483(26) Argento-Ceru, M. P.,535 Arima, K.,211, 212(63), 214, 511 Armstrong, M. D.,180, lSl(348) Arneson, R.M.,358, 536 Arnon, D. I., 19, 58, 59, 60, 61, 62(24), 64(24, 25) Aro, H.,155,293 Aronson, R. B., 158, 165, 166, 167(281), 186(219) Arst, H.N.,406, 413(307b) Asatoor, A. M.,466 Ashford, J. S.,516, 517(63), 518(63), 524 (631,525, 526 Ashwora, J. N.,560 Atkins, P.W.,329 Aubert, L.,280 Aubort, J.-D., 182 Aures, D.,514, 519(48), 523(48) Aust, S.D.,275, 533(151), 534 Autor, A. P.,272 Autuorio, F.,535 Averill, B. A.,8, 11(27b), 35, 37(27b), 43 (1061,44, 46 Avigad, G.,528, 530(122) -4viglian0, L.,536, 560 Avis, P. G.,303, 305, 306, 307, 308(49), 309(49), 310, 311(21), 312, 313, 359 (21), 3971491, 398(49) Avron, M., 26 Axelrod, B., 150 Axelrod, J., 186,254,269(7) Aaada, K.,538, 539(223), 5410231, 542, 544, 545(223), 547(223), 556(223) Azoulay, E.,280

B Babin, D. R., 302,366(16) Babu, B. H., 470 Bacher, A., 311, 328(60), 341(60) Bachmayer, H., 8, 12(26) Bachofen, R.,61 Bachrach, U.,516 Bacila, M.,527,531(119) Raillie, D.L.,314, 407(67c) Baker, B. S.,407 Baker, G.8. T., 543, 546(246), 549(246) Baldes, K.,287

Baldwin, R. L., 110, 117(172) Baldwin, T.O.,227 Bale, J. R.,47 Ball, E.G.,303, 305(19) Ballou, D.P.,31, 56, 105, 106(155, 156, 158, 159), 192, 213, 214, 215(73), 216 (691,221(73), 246(2, 3), 304, 306, 313 (451, 314(46), 326(46), 327(46), 328 (45),331(46), 332(128), 336(46, 337(46), 338(46), 339( 1281, 340(45), 341(461, 344(128), 353(45), 355(45), 356(45, 46), 362(45), 364(45), 365 (45), 368(46), 369(46), 371(46), 372 (31, 451, 374(45, 46, 1281, 375(46, I%), 376(46, 128), 377(128), 378(46), 379(46), 380(46), 381(46), 382(46), 383(45, 46), 384(45, 46, 201), 385(45, 46), 386(46), 387(46), 388(45, 46), 399(46, 201), 405(46), 424, 504, 505, 553 Balny, C., 228, 248(103), 251 (103) Bankel, L.,174, 177, 178(332) Bannister, J. V.,539, 540(231), 541(231, 239), 543(231, 239), 545(231, 239), 546, 547,548(231), 549(231, 239), 551, 553(283), 554(283) Bannister, W. H.,539,540(231), 541(231, 2391, 543(231), 545(231), 546, 547, 548(231), 549(231, 239, 2501, 551, 553 (2831,554(283) Barb, W. C., 358 Barber, M. J., 321, 322, 336(90b), 369 (gob), 374, 378(265), 391(9Ob), 393 (gob), 394, 395(90b, 91a), 396(90b), 397(90b), 399(90b, 91a, 288) Barboni, E., 149 Bardsley, W. G.,516, 517, 518(63), 523, 524,525(101), 526 Barker, H. A., 345, 346(168), 348(168), 349, 350(168), 360, 361(168), 369 (168), 390(168), 391(168), 397(168), 398(168) Barman, B. G.,82, 83, 84(100, 107, 1081, 85, 86(108), 87(107), 99(100), 100 (loo), 104(100) Barnes, M. J., 152 Baron, J., 19,268,273 Barra, D.,148, 540, 542(241), 5430411, 545(241), 548(241), 549(241), 551 (241)

583

AUTHOR INDEX

Barrett, J., 259, 263(26), 264(26) Barry, G., 325 Barth, G., 311, 328(60), 341(60), 543, 546 (248), 548(248), 549(248)

Bartholomaus, R. C., 263 Bartlett, P. O., 284 Barton, L. L., 60 Bartsch, R. G., 32,33(90), 35 Bates, C. J., 153, 162 Battelli, M. G., 321, 322 Bauer, D. J., 359 Baumann, E., 179 Baxendale,-J. H., 358 Baxter, R. M., 280 Bayer, E., 311, 328(60), 341(60) Bays, J. P., 96, 97(134) Beacham, L. M., 306, 307(44), 308(44), 310(44), 311(44), 312(44), 315(44), 317(44), 324(44), 325(44), 326(44), 327 (44), 352 (441, 368(44), 374(44), 382(44) Bearden, A. J., 10(66), 20, 26, 29, 30, 341 Bearn, A. G., 562 Beattie, D. S., 230 Eeauchamp, C. O., 357, 535, 538, 539, 540 (232), 541(224), 542(224), 544, 545 (224), 552(224), 556(224), 557(232) Beckman, G., 537, 540, 543(204)

Beckman, L., 540 Behar, D., 356, 357(206), 553, 556(297) Behrman, E. J., 146 Beinert, H., 10(66), 22, 23(50), 26, 30, 32, 40, 56, 57, 59, 62(21), 64,104(21), 109, 110, 111(176), 115(1), 116(1, 176), 117(1), 118, 195, 261, 262(37), 263, 266, 269(67), 280(37), 303, 308(22), 325, 327, 328, 329, 337, 338(146), 341, 342(118), 343, 356, 369, 370, 372(263), 373(263), 374(263), 376(263), 377 (263), 383(263), 385(263), 388(146), 389, 391(2831, 394(283), 395(162, 283), 396(162, 283), 424, 449, 479, 517, 518(66), 519(66), 520(66), 521(66), 533(147), 534 Belyaeva, V. K., 333, 334(133), 335(133), 339(136) Benedict, J. J., 368 Benemann, J. R., 59, 62, 64(24, 25) Benfield, G., 574 Bennett, L. E., 14, 15, 46, 152

Bennett, R., 445 Benoiton, N. L., 240 Bentley, R., 466 Berdnikov, V. M., 556 Berekovskii, V. M., 112 Bereman, R. D., 530 Berends, W., 112, 241, 242, 243, 245, 251 (157)

Berg, R. A,, 153, 155, 156, 159, 160(195), 162, 163(195), 292, 293

Bergel, F., 302, 303, 305, 306(21), 307, 308 (49), 309(49), 310(21, 49), 311(21), 312(40, 49), 313(40), 326(39), 359 (21), 397(49), 398(49) Bergeret, B., 517 Bergmann, F., 345, 346, 348,349, 350, 351 (166) Berlin, C. M., 128, 172(32) Bernhardt, F. H., 286, 287 Bernstein, L., 162 Bertrand, G., 508, 558 Bessey, 0. A., 362,363(240) Betcher-Lange, S., 414 Beyer, E., 317 Bhaduri, A. P., 108, 246,424 Bhatnagar, R. S., 158(229), 159, 186 Bieber, S, 302 Bier, C. J., 542,557 Biggo, D. R., 125, 149(18) Bishop, C. A., 556 Bjorkhem, I., 126 Bjorling, H., 560 Bjur, R. A., 110, 115(166) Blair, J. A., 242, 244, 251(162) Blankenhorn, G., 108, 246, 424, 473 Blaschko, H., 466, 512, 513(36), 515, 517, 518, 519, 522(74), 523(74) Blout, E. R., 158(226), 159, 160(226) Blumberg, W. E., 2, 22, 139, 261, 262(37), 280(37), 414, 529, 530(129), 551, 565, 566, 571(376) Blumenfeld, 0. O., 152, 165(139) Blumenkrantz, N., 167 Boardman, N. K., 537 Bobrik, M. A., 47 Bock, R. M., 393 Bocks, S. M., 574 Boden, N., 518, 519(73), 520(73) B@hmer, T., 167 Bogacki, I. G., 227

584 Bohnenkamp, W., 539, 546(233), 548 Bokman, A. H.,145 Boldingh, J., 536 Bonner, J., 559 Booth, V. H.,345, 346(167), 349067) Bornstein, P.,152, 159, 160, 166 Borst, P.,537 Bossa, F., 540, 542(241), 543(241), 545, 546(251), 547(251), 548(241), 549 (241), 551(241), 566 Bothe, H.,59, 61(19, 201, 62, 63, 64, 65 (19), 90(51), 99(51), 102(51), 104(19, 20, 51) Boucek, M., 164 Bouchilloux, S., 296, 297(225) Bowman, M.,23 Boyd, G.S.,266,269(66) Boyer, P.D.,125, 150(19) Bradley, P.L.,359 Bradshaw, W.H., 345, 346(168), 348(168), 349, 350(168), 360, 361(168), 369 (1681, 390(168), 391(168), 397(168), 398(168) Brady, F. O., 129, 312, 319, 320(78), 353 (78), 354(78) Briindkn, R.,569, 570(390), 577, 578 Brandt, K.G.,465,483(121) Braneoli, U.,317, 319(77c), 323, 354(77c), 390(77c), 391(77c), 393(96b), 398 (77~1,445, 446(59), 499, 500(202) Bray, R. C., 50, 54(123), 301, 302, 303, 304, 305, 306, 307, 308, 309(26, 36, 49), 310, 311, 312(4, 36, 40, 491, 313(36, 40, 43), 314(4, 43), 315, 316, 317, 319, 320(70), 321, 322, 323,324, 325(5, 98), 327,328,329,330(32, 73, 98), 331, 332, 333, 334, 335, 336, 337, 338, 339, 340 (1531,341, 342(25, 26, 1591, 343(153), 344 (4, 5), 352(4, 41), 354(4), 355, 357, 358(4), 359(21, 361, 360(4), 361, 362(4), 363(4, 1251, 366(4), 368(5, 691, 369, 370, 372, 373, 374, 375, 376, 377, 378(70, 153, 2651, 381(125), 382 (98, 126), 383, 384(197), 385(29, 263), 386, 387, 388, 389, 391(90b, 2861, 393 (36,90b, 2861,394,395(90b,91a, 286), 396(90b, 2861, 397(47, gob), 398(49), 399(90b, 91a, 2881, 419(272), 510, 533 (147),534,547,550,551,552, 553, 554, 555, 556

AUTHOR INDEX

Bremer, J., 167 Bremser, W.,548 Brennemann, A. R.,238,239(139) Bridgers, W.F.,232,295 Bright, H. J., 154, 157(189), 158(189), 159(189), 427, 428, 431, 432, 434(46), 435(39), 440, 442(26, 39, 51), 443, 445 (39), 449, 450(39), 451(39), 452(39, 46), 453(39), 454(87), 456, 461(53), 463(51), 464, 465(26, 271, 476, 477 (51), 479(46), 480(30, 39, 531, 481 (165), 482(46, 51, 165), 483, 485, 486 (39), 488(39, 185), 490(39), 491(39, 185), 492, 494(165, 187), 495(39, 165, 187), 496(39, 165), 497(165), 501 (165), 502(165, 182) Brignac, P. J., 531 Briley, M. S.,322 Brill, A. S.,361, 368 Brill, W.J., 50,52(122) Brinteinger, H.,4, 22, 301, 327(2), 368(2), 372(2) Briving, C.,562 Brockman, H.L., 110, 116(175), 117(175) Brodie, B. B., 186,254, 269(7) Broide, J. D.,532 Broman, L.,559, 561, 563, 564(355), 571 (356), 576 Bromley, K.,574 Broyuist, H.P.,167 Brown, B. R.,574 Brown, D.J., 345, 346(164, 166), 348(164, 1661,349(164), 351(166) Brown, D. M., 538, 542(229), 543(229), 545(229) Brown, G.B., 352,355(180, 182), 361(180, 182) Brown, J. R.,59, 60(27), 66(27), 99(27) Brown, L.E., 500,501 Brown, N. C.,180 Brown, R.D.,548 Brown, R. M.,Jr., 537 Brownie, A. C.,266,269(67) Briistlein, M.,90, 108,246, 248, 319, 328, 424,480, 497 Bruice, T.C.,496, 497, 499, 500, 501, 502, 503(193), 505(198) Brumby, P. E., 305, 307(35), 308(35), 310 (351, 311(35), 312(35), 313(35), 314 (35), 315, 326, 327(35), 328(35), 353

585

AUTHOR INDEX

(35), 361(35), 362(35), 368(35), 369 (351, 372(35), 374(35), 381(35), 388, 432, 433(43), 440(43), 445, 446 Brundage, G. B., 239 Brunnenberg, E , 541(2451, 542, 545(245), 546(245) Brunori, M., 447, 499, 500(202) Bruschi, M., 61, 64(39), 104(39) Bublitz, C., 185 Buchanan, B. B., 61 Buehner, M., 77, 447 Buffoni, F., 515, 517, 518, 519, 520, 521 (81), 522(74), 523(74), 525 Bulen, W. A., 59, 60(22), 63(22), 99(22), 104(22) Bulgrin, V. G., 440 Bulla, L. A., 535 Bunbury, E., 325 Bunnenberg, E., 311, 328(60), 341(60) Bunton, C. A., 184 Burger, A., 158, 160(223) Burnett, J. D., 296,297(231) Burnett, R. M., 66, 70, 71(65, 691, 72, 73(66), 74(69), 75(69), 76(69), 77 (691, 78, 79(69), 80(69), 81(69), 82 (691, 94, 98(66), 101(69), 102(69), 114(66), 447 Burris, R. H., 32, 34(85), 44, 47(108), 50, 52, 55 Butler, W. T., 165, 166 Butzow, J. J., 562 Byers, W., 569, 571(389)

C Cain, G. D., 164 Cain, R. B., 143 Calabrese, L., 125, 149(16), 357, 535, 536 (178), 540, 542(241), 543(241), 545, 546, 547, 548(241, 2551, 549(241), 550, 551, 552(263, 2841, 553, 554(263, 284, 294), 555(255, 263, 294), 556(255, 2941, 557(294), 560 Calabrese, R. L., 275, 276(128) Calvin, M., 211, 212 Cambier, H. Y., 403 Cameron, J. C., 401, 402(295a) Camilleri, P., 546 Cammack, R., 8, 9(24), 10(24), 11(98), 12(24), 15, 17, 24, 29, 30, 33, 35, 37,

341, 342(159), 539, 541(245), 542, 545 (2451, 546(233, 245) Canale-Parola, E., 32, 34 Candido, E. P., 314, 407(67c) Cannella, C., 148 Cantor, C. R., 17 Capalna, S., 270 Capeilkre-Blandin, C., 386, 419(272) Carbon, J. A., 524, 525( 100) Cardinale, G. J., 152, 153, 154(174), 155 (1441, 158(144), 159, 160, 162, 163 (2531, 184, 185, 186, 188 Cardini, G., 280 Carnahan, J. E., 58, 61(2) Carper, W. R., 526 Carrico, R. J., 533, 538(144), 540(144), 541(144, 145), 542(144), 543(144), 545(144), 546(144), 548(145), 549 (1451, 570, 572(395), 576, 577(428), 578(428) Carter, C. W., Jr., 32,33(90-92), 35(90-92) Cartwright, G. E., 533, 538, 541(215), 548 (215) Cattau, E. L., 389, 397(279a), 398(279a) Caughey, W. S., 534 Cavallini, D., 125, 148, 149(17) Chakrabarty, A. M., 259 Chalmers, R. A., 402 Chambers, R. W., 536 Champion, P. M., 143 Chan, H. W. S., 150, 536 Chance, B., 427, 437, 442, 452, 453 Chandrasekhar, K., 77 Chang, J. J., 227 Chang, L. O., 163 Chapman, H. R., 305, 306(36), 307(36), 308(36), 309(36), 310(36), 311(36), 312(36), 313(36), 317(36), 319(36), 326(39), 359(36), 393(36) Charlton, S., 40, 41(102), 42(102), 43(102) Charlton, S. C., 518, 519(73), 520(73) Chassy, B. M., 446,447 Chaykin, S. J., 351, 355, 367(190), 368 (190), 389(179), 390(179), 391(179), 393(179), 397(179) Chen, J. S., 50 Chertock, H., 239, 291 Chervenka, C. H., 515(55), 516, 519 Chevalet, J., 556

586 Chiancone, E., 447, 517,518(66), 519(66), 520(66), 521(66) Childs, R. E., 523 Chisholm, A. J., 306, 310(41), 352(41) Chovnick, A., 314,407(67c),410, 411 Chrispeels, M. J., 154, 163, 164 Christensen, J. J., 362 Christian, W., 461 Christman, D. R., 166 Chu, A. E.-Y., 351, 389(179), 390(179), 391(179), 393(179), 397(179) Chu, J. W., 267,268 Chuang, H. Y. K., 467, 468, 469, 470, 471 (139), 498(139) Chvapil, M., 157, 164 Cilento, G., 536 Cinti, D. L., 270, 277 Civen, M., 181 Clark, C. T., 186 Clark, W. M., 100, 356 Cleere, W. F., 316, 317, 319(75b), 323 (75d), 345, 346(169), 348(169), 349, 350, 351(169), 353(75b), 360, 361 (1691, 367(75d), 390(169), 391(169), 394(75b, 169), 396(75b), 397(169), 398 (169) Cleland, R., 163 Cleland, W. W., 369, 429, 525 Clerin, D., 501 Cleveland, L., 557 Closs, G. L., 478 Closs, L. E., 478 Cockle, S. A., 547, 552(263), 554(263), 555(263) Coddington, A., 405 Coffey, D. S., 455 Coffman, R. E., 24 Cahen, B. S., 275,276(125), 279 Cohen, H. J., 309, 414, 415, 416, 417(333, 334, 3371, 418(333, 336a, 3371, 419 (336a) Cohen, S. S., 537 Cohn, E. J., 560 Cohn, M., 82,96(99), 97(99) Coleman, J. E., 545, 546, 550, 566 Coleman, R. V., 550 Collins, G. G. S., 469 Collins, J. F., 407, 408, 409(311, 3161, 410 Comai, K., 187 Comstock, J. P., 163

AUTHOR INDEX

Conney, A. H., 270,273 Cook, D. E., 229 Cook, K. A., 50, 51(119), 53(119) Coon, M. J., 6, 8, 13, 14, 271, 272, 273, 280

Cooper, D. Y., 254, 255(8), 265, 266, 267, 268(59, 68, 691, 270, 274, 275, 276 ClZS), 282(69), 283(68) Cooper, G. W., 153 Cooper, J. A. D., 527, 531(119) Cooper, J. M., 427 Cooper, J. R., 232, 254, 269(7> Corcoran, W. H., 103 Cordes, E. H., 373 Cordes, M. M., 213 Cormier, M. J., 227 Corran, H. S., 303, 305(18), 306(18), 352 (18) Correia, M. A., 279 Costa, M. T., 125, 148, 149, 516, 517, 518 (66), 519(66), 520(66), 521(66), 522, 523(89), 525, 526(106) Costilov, R. N., 535 Cotton, F. A., 77, 334, 339 Coughlan, M. P., 316, 317, 319(75b), 321, 322, 323, 324(96), 325, 326(96), 336 (gob), 345, 346(169), 348(169), 349, 350, 351(169), 353(75b), 360, 361 (1691, 367(75d), 369(90b), 390(169), 391(90b, 1691, 393(90b, 961, 394, 395 (gob, 91a), 396(75b, 9Ob, 961, 397 (gob, 169), 398(169), 399(90b, 918, 288) Courtright, J. B., 407, 408(315), 409(315) Cove, D. J., 402, 405, 406, 413(307b) Cowling, E. B., 559 Cox, J. M., 79 Crabbe, M. J. C., 523, 525 Craine, J. E., 233, 295 Crandall, D. I., 146, 182 Crane, F. L., 110, 111(176), 116(176) Crawford, E. J., 362, 363(240) Crawhall, C. J., 183 Crespi, H. L., 58(17, 181, 59, 62/17), 96, 97, 99(17, 181, 109 Cretney, W. C., 50, 52(121), 53(121) Cronin, J. R., 110, 115(177), 117(177) Croasley, H. L., 153(177), 154 Cryer, D. R., 537

587

AUTHOR INDEX

Csoregh, I., 76, 82(89), 102(89), 424, 425

(21) Cunningham, L. W.,165 Curti, B.,203, 430, 431, 432, 433(43), 435

(381, 437(38), 440(37, 38, 431, 442 (38), 445, 446, 447(62), 448(37), 455, 457, 458, 476(38), 480(38), 499, 500 (202), 504(38) Curtin, W.E., 94 Curaon, G.,534, 559, 569, 571(389), 575, 576(422) Cusanovich, M. A., 58, 59(14), 60(14), 62(14), 90(14), 95(14), 104(14) Cushman, D. W.,263 Cutroneo, K. R.,153(180), 154, 162 Czapski, G.,356, 357(206), 553, 556(297) D Dagley, S., 143 Dahm, H.,182 Dalgleish, D. G.,540, 541(239), 543(239),

545, 546(239, 2501, 549(239, 250) H.,65, 322, 336, 389, 391(286), 393(286), 394, 395(91a, 2861, 396 (286), 399(91a, 288) Daly, J. W.,182, 219, 220, 221(81), 234, 238,240(140),284,290 Dalziel, K., 429 Daniels, G. H.,295 Danielson, H., 126 D’Anna, J. A., Jr., 66, 82, 83032, 101), 84(62), 85(62), 87, 93(62), 94, 95 Danowske, T.S.,456 Darling, G.D., 66, 70(65, 66), 71(65, 69), 72(66), 73661, 74(69), 75(69), 76 (69), 77(69), 78(66), 79(69), 80(69), 81(69), 82(69), 94(66), 98(66), 101 (69), 102(69), 114(69), 447 Darlington, A. J., 302, 412(9) Darnell, J., 153, 159(149) Davidowicz, K., 537 Davis, L., 557 Dawson, C. R., 296, 558, 559, 560, 561 (324), 562, 563, 564(345), 565, 567 (3451,571(345), 574 Dawson, J. W., 577,578(431) Debrunner, P. G.,10(97b), 35, 263, 264, 282(54) de Dugros, E. C.,358,536 Dalton,

Dehm, P.,153 Dehner, E. W.,229 Deinum, J., 562, 564(346), 565(344, 346),

566(346), 567(344, 3461, 568(346), 571, 572(344), 573(344) Dekker, E.E.,189 Dekker, I., 113 DeKlerk, H.,32 De Kok, A., 456 de Lamirande, G.,302 De Lapp, N. W., 397 Delcambe, L., 204 Della Corte, L., 320, 321(87), 322, 366, 389(95), 402(95), 518, 519, 520, 521 (811,525 DeI Sole, P.,535,536(178) DeLuca, D. C., 454,470 DeLuca, M.,188 Delwiche, C. V., 187 Demain, A. L.,63, 65(50) De Marco, C., 125, 148 Dempseyj M. E.$ Dent* E.p 400 DePamphilisy 46 De Renzo, E. ‘., 301r 303(7)v 410 Dervartanian, D. V.,64,263 DeSap R. J., 4589 460 D’Eustachio, A. J., 58, 59(6), Q9(6) Deutsch, H.F., 533, 538, 539(216, 217), 540( 1441, 541 (144, 145, 216-218), 542, 543, 544, 545(144, 217, 2181, 546(144), 548(145), 549(145), 561, 562 de Vries, A., 359 de Vries, J., 404 Dewant J. G.$ 305(18)* 306(18), 352 (18) Diamond, R ~ 74, , 76(85), 77(85), 78(85), SO(%) Dickerson, R. E., 69 Dickinson, W.J., 407, 408 Dickson, D. P. E., 8, 9(27a), 10(27a), 11(98), 35,37 Dicorleto, P. E., 551, 552, 554(285), 555, 556 Diegelman, R. F., 16% 164 Diehl, H., 270, 277, 278(130), 279, 284 (130) Dikstein, S., 350 Dimitrijevic, L., 534 D’Iorio, A., 240 v*j

4227

303j

588 Dixon, M.,242, 303, 448 Djerassi, C., 311, 328(60), 341(60), 541 (245), 542, 543, 545(245), 546(245, 248),548(248),549(248) Dorfman, L. M., 356, 357(206), 358, 553, 556(297) Dorfman, R.I., 255 Douzou, P.,385 Dragila, T. A,, 170, 171(319), 175(319), 176(319) Draper, R. D., 98, 99(141), 102, 499 Drew, R.T.,414 Dubourdieu, M.,58, 59(10, ll), 60, 61, 62(10, ll), 64(39), 66, 67(67), 69(67), 70(29), 82(11), 99(11), 104(39) Dubrov, Y. N.,333, 334(133), 335(133), 339(136) DuBus, R.,259, 263(29) Duck, P.,411 Dudley, K.H., 244, 251,480 Duke, E. J., 312(65, 65a, 66), 314, 407, 408(316), 409(316), 410 Duke, F. R,., 440,499 Dukes, G.R.,47 Dunham, W. R., 10(66), 20, 21, 26, 341 Dunn, D. K.,227 Duppel, W.,272, 280 Dupr6, S.,148,149 Dus, K.,18,32, 35, 259, 260(28), 262, 263 (28), 264, 268(55), 283(39) Duthie, R., 512 Dylewski, I., 183

AUTHOR INDEX

Edwards, J. O.,184 Edwards, P.,306, 313(43), 314(43) Edwards, S. W.,146, 179, 181(344) Ehrenberg, A., 37, 78, 90, 95(117, 1211, 96, 100(121), 248, 249, 251, 304, 307 (26), 309(26), 311(26), 327, 341(26), 342, 361(26), 369(26, 271, 370(26), 372(26), 374(26), 387(26), 424, 480, 550,563, 571 (356) Ehrig, H., 286, 287(161) Ehrlich, E., 164 Ehrlichova, M., 157 Eisenberg, F.,232, 295 Eisenger, J., 566, 571 (376) Eisenthal, R.,322 Elion, G. B., 302, 303(10), 306, 307(44), 308(44), 310(44), 311(44), 312, 315 (44, 611, 317(44), 324(44), 325(44), 326(44), 327, 337(110), 345(10), 346 (lo), 348(10), 349(10), 350(10), 352 (10, 441, 354(10, 611, 359(10), 363, 364(61), 367(61), 368(44), 374(44, 611, 382(44), 387(61), 397(10), 398 (lo), 402(10), 408(10), 413(10) Ellenbogen, L.,239 Ellin, A., 270 Elsden, S.R.,64, 110 Embden, G.,287 Engel, L.L.,254 Engel, P.C.,106, 108(160), 110, 192, 225 (4),246(4), 534 Engelman, K., 302 Engelman, R.,401 Entsch, B.,63, 82(46), 91, 99(46, 125), E 100(125), 104(125), 213, 214, 215, 216 (691,221(73), 505 Eady, R. R., 50, 51, 52, 53(119), 55 Eaton, W. A., 5, 6, 8, 12, 24, 91, 92, 93 Erbes, D. L.,262, 283(39) Erdin, N.,286, 287(162) (126) Ericsson, J. L.E.,274 Ebata, M.,515(57), 516 Erickson, R. H.,309, 404, 405(305, 306) Edelstein, S.J., 445,446(60) Edmondson, D., 56, 306, 307, 308(44), Eriksson, G.,327 310(44), 311, 312, 315(44), 313(44), Eriksson, L.E. G., 96,424 323, 324(44, 62), 325(44), 326, 327 Erlenmeyer, H.,250(181), 251 (441,331, 332(128), 336, 337(62), 339 Ermakov, A. N.,333, 334(133), 355(133) (128),344(128), 352(44), 368, 374(44, Ernster, L.,274 Erwin, V. G.,467, 468, 469(129), 470 1281, 375(128), 376, 377, 382(44) Edmondson, D. E., 58, 59, 60(14), 62(14), Estabrook, R.W.,19, 254, 255, 265, 267, 268, 270, 271, 274, 275, 276(119, 125, 65, 82, 83(61), 84(61, 1081, 85, 86(61, 127), 279 102, 108), 87(23, 611, 90, 95(14, 231, 99(61), 100(23), 104(14), 107(102) Ettinger, M. J., 530, 532

589

AUTHOR INDEX

Ettlinger, L., 297 Evans, H. J., 547 Evans, M. C. S., 11(98), 35, 37(98) Evans, M. C. W., 8, 9(24), 10(24), 12 (24),29,30, 53 Everse, J., 77 Ewall, R. X., 46 Ewetz, L., 125, 149

F Fahien, K. A., 240 Fiihraeus, G., 559, 560, 561(329), 562 (329), 563, 574 Fairbain, D., 164 Falk, I<.-E.,566 Falkenbei-g, B., 59, 61(20), 63, 104(20) Faraggi, M., 384, 399(271), 577, 578(432, 433) Farber, T., 153(178), 154 Faria Oliviera, 0. M. M., 536 Fasella, P., 566 Federici, G., 125, 148, 149 Fee, J. A., 8, 21, 22, 23, 24(23), 26(45), 263, 535, 540, 542, 543, 546, 547, 548, 549(246, 249), 550, 551, 552, 554(285), 555, 556, 564, 569(363), 570, 572, 573, 574(3931, 579(407) Feigelson, P., 128, 129, 130, 359 Feldberg, R., 66, 105(63), 106(63), 108 (631, 116(63), 192, 246(5), 248(5), 399, 422 Felix, A., 533(152), 534 Felix, A. M., 159, 160(231) Fellman, J. H., 180, 181(349, 352), 183 (349) Felsted, R. C., 351, 389(179), 390(179), 391(179), 393, 397(179) Ferri, G., 445, 446(59) Fetzer, V., 239, 291 Fielden, E. M., 322, 357, 394(91a), 395 (91a), 399(91a), 547, 551, 552, 553, 554, 555, 556, 557(294) Finazzi Agrh, A., 516, 517, 518(66), 519 (661, 520(66), 521, 522, 523(89), 525, 526(106), 535, 536, 540, 542(241), 543 (2411, 545, 546(251), 547(251), 548 (241), 549(241), 551(241), 560, 567, 577, 578(429) Fink, K., 169, 172(315), 177(315)

Fink, R. M., 169, 170, 171(318), 172(315), 174(318), 177(315) Finley, R. G., 173 Finnerty, V., 406, 407(307d), 409(307d), 411 Fiori, A., 148, 149 Fischer, E. H., 522, 523(91) Fisher, D. B., 193, 234(13), 236(128), 237, 238, 240(13), 241, 242(13), 251 (1361,288,289 Fisher, J., 498 Fisher, J. R., 302, 358(226), 359, 360(232), 361, 369(226), 397 Fitch, W. M., 67, 68(73), 69 Flamm, W. G., 146 Flashner, M. S., 188, 193, 194(10), 200, 201(33), 203(32), 210 Flohk, L., 535, 541(245), 542, 545(245), 546(245) Flood, T. C., 289 Fory, W., -93 Foguelman, A. I., 412, 413(328) Folch, J., 533, 538(141) Fonda, M. L., 428 Fong, K.-L., 358 Foote, C. S., 536 Forchielli, E., 255 Ford, G. C, 447 Ford, G. L., 77 Forman, H. J., 129, 130, 540, 547, 549, 551(273), 552,555(237) Forti, G., 31, 108 Foust, G. P., 19,31, 62,65, 79(42), 98(58), 99(58), 100(58), 102(58), 106(58), 107(58), 108(42), 192, 246(5), 248(5), 356, 399, 422, 472, 473(159) Fouts, J. R., 524, 525(100) Fowlks, W. L., 120, 254, 296(2) Fox, A. S., 407 Fox, J. L., 59, 60(27), 66, 67(27), 69(67), 99(27) Francke, A., 150 Franconi, C., 550, 551(274) Frank, M., 359 Frankel, R. B., 8, 11(27b), 25, 35(27b), 37(27b), 46, 341 Frazier, W. A., 403, 404(302) Freedland, R. A., 234 Freer, S. T., 32, 33, 35(90-92) Fretzdorff, A., 538, 540(219), 541(226),

590

AUTHOR INDEX

542(219), 543(219, 226), 545(219, 226), 546(219) Fridovich, I., 186, 256, 326, 345, 353, 354, 355, 356, 357, 359(215), 362, 366, 367 (250), 368, 385, 386(236), 389, 392 (2821, 397(171, 282), 414, 415, 417 (333, 3371, 418(333, 336a, 337), 419 (336a); 510, 533, 534, 535, 536, 537, 538, 539, 540, 541(16, 200, 220, 224, 225, 227, 2951, 542(16, 200, 220, 224, 225,227), 543(200, 220, 225, 227), 544, 545(200, 220, 224, 225, 2271, 547, 548 (161, 549, 551(16, 273), 552, 553, 554 (293,2951, 555, 556(16, 224, 295), 557 (232) Fried, L.W., 302, 366 Fried, R.,302,359, 366 Frieden, E.,559,562,575 Friedman, P. A., 241, 291, 292(188) Friedman, R.,302 Friedman, S.,232,294,295 Frisell, W.R.,110, 115, 117(177) Fritchie, C.J., Jr., 76 Fritz, I. B.,167 Fritz, J., 96, 263 Frommer, U.,275,276(123), 277(123), 284 Fruton, J. S.,508,558(6) Fuchs, S., 26 Fujimoto, D.,154, 156, 168, 159(224), 164 (1851, 292 Fujioka, M., 147 Fujisawa, H.,130, 134, 135, 136, 137, 138 (49, 50, 771, 139, 140, 141, 151(72) Fujita, T.,271 Fujita, T.S.,180, lSl(349, 352), 183(349) Fujit8a, Y.,160 Fujiwara, M.,135, 137, 139(78), 151(78) Fukami, H.,516, 522 Fukumoto, J., 511 Fuller, G. C.,153,154 Furthmayr, H.,166 Furuya, E.,146 Futai, N.,258 G Gaal, A.,221,223(82) Gaber, B. P.,546,547, 548, 551(258, 265), 556 (258) Gainulin, I. F., 333, 336(138)

Gajda, L., 58(17), 59, 62(17), 99(17) Gallo, M.,280 Gallop, P.M.,152, 165(139) Gander, J. E.,145 Ganther, H.,424, 425(19), 430, 432, 433 (43), 440(37, 43), 441(19), 446, 448 (37), 452(19), 484(19) Garbett, K., 328, 341(122), 569, 571(389) Gardner, K.G., 334 Garfinkel, D., 254, 270 Garif’yanov, N. S., 333, 336(138) Garrett, N.J., 469 Gartner, B.,471, 498(154) Cast, R.,87 Gaudette, L.,254,269(7) Gauthier, J. J., 147 Gaylor, J. L.,187, 269, 272, 275, 276(128), 282 Geary, P. J., 143 Gee, P.,515(57), 516 Gehring, U.,60 Gelbart, W. M., 411 George, P.,256,358, 556 Gerbeck, C. M., 531 Gerosa, P.,560 Gersonde, K.,45 Ghisla, S., 91, 93(124), 102, 112, 113(181), 114(181), 115(124, 1811, 116(181), 199, 424, 446, 480, 493(7), 494, 496 (1891, 502(189) Gibb, J. W., 290 Gibian, M.J., 500 Gibson, D. T.,124(11), 125, 144 Gibson, J. F., 22, 327, 341, 355, 384(197), 534 Gibson, Q. H., 105, 192,203,206, 207(54), 208(54), 228, 246(1), 427, 430, 432, 434(25), 435(25, 361, 436(25, 36), 437, 438,440, 441(25,361, 442, 443(36, 511, 448, 449(36), 451(36), 452, 454(36), 458, 460, 462, 463, 464(51), 465, 476(25, 36)’ 477(25, 51), 479, 480 (36), 481(36), 482(51), 503, 504(218) Gierst, L., 556 Gigon, P.L.,277 Gilbert, D.A., 302, 363 Gilchrist, F. M. C. 64, llO(54) Gill, D.,6 Gillard, J. M.,77, 83(90), 107, 108 Gillard, R. D.,328, 341(122)

591

AUTHOR INDEX

Gillette, J. R.,274,277,279 Giordano, R.S.,530 Giovagnoli, C., 535, 536(178), 546, 547 (2551, 548, 555(255), 556(255, 267) Glahn, P.E.,456 Glassman, E.,407, 408, 409, 410, 411 Glick, D.,291 Goda, K., 536 Goldberg, B.,153 Goldring, E. S.,537 Goldstein, A., 164 Goldstein, M.,240,295 Golovleva, L.,135 Gomes, B.,467,468, 470(131) Gonnerma, W.A.,527 Goodman, B.A., 329 Goodman, D.S., 126 Goodman, P.A., 345, 346(170), 349(170) Goodwin, S., 182 Gordon, A. H.,303, 305(18), 306(18), 352 (18) Gorkin, V. Z., 467 Goryachenkova, E.V., 516, 518(60), 519 (60), 520(60), 521(60), 522(60) Goscin, S. A.,534,535,538, 539, 540(232), 541(225), 542(225), 543(225), 544, 545(225), 547(225) 557(232) Goswami, M. N. D., 179(347), 180, 181 Gottlieb, A.,160 Gowie, E.A.,88 Gozzetti, G., 322, 389(95), 402(95) Grahame-Smith, D. G., 241, 291 Gram, T.E.,277 Grant, M.E.,152,292 Gravey, T.Q.,111, 295 Gray, H. B., 8, 11(27b), 24, 35(27b), 37 (27b),567,577,578(431) Graziani, M. T., 148 Green, A. A., 227 Green, A. R.,534 Green, D. E.,303, 305(18), 306(18), 308 (22),352(18), 393,456 Green, H., 153 Green, N. M.,156 Greenawalt, J. W., 230,467 Greengard, O.,129 Greenlee, L.,317, 351(77), 355, 362(77), 536 Gregerman, R. I., 179, 180(346), 181(346), 183(346)

Gregory, D., 345, 346(170), 349 Gregory, E.M.,256,534 Grell, E., 550 Griffin, B. W., 130, 260, 261(32), 263, 264 (53),283(53) Griswold, W.R., 173, 174(329), 175(329), 178, 179 Grivell, A. R., 172, 173(326) Grivell, L.A., 537 Groseclose, E.E.,224 Grossman, L.I., 537 Grosso, E.D.,148 Grunthan, F. J., 8, 11(27b), 35(27b), 37 (2%) Guerrieri, P., 545, 546(251), 547(251) Guilbault, G. G., 531 Gunnarsson, P.-O., 567, 575 Gunsalus, I. C.,10(97b), 18,19,22, 23(49, 50), 35, 255, 259, 260, 261, 262, 263, 264, 268(31, 55), 280(37), 282(54), 283(39) Gurin, S., 179 Guroff, G., 220, 221(81), 234, 235, 238, 240(10) Gurtoo, H. L.,353 Gutfreund, H., 352, 353(181), 369, 372, 373, 432 Gutman, A. B., 302, 363(12), 401(12) Gutman, M.,301 Guaman, N. A., 153(180), 154, 162

H Haas, W., 108, 244, 246, 424, 501 Hackert, M.L.,77 Haddow, A.,302 Haffner, P.H., 546 Hafner, E. W.,456, 461(95), 480(95) Hager, S. E.,179, 180(346), 181(346), 183 (346) Hall, C. L., 110, lll(l68), 115(168) Hall, D. O., 8, Q(24, 27a), lO(24, 27a), 11(98), 12(24), 15, 17, 22, 24, 35, 37 (981,341, 342(159) Hall, E.S., 233 Hall, F. B.,412, 413(328) Hall, P.F.,269 Halme, J., 153, 154, 155, 158, 292 Hamberg, M.,126 Hamilton, G. A., 184, 185, 186, 194, 225,

592 248, 284, 368, 382, 500, 501, 528, 529, 530, 531, 532 Hamilton, W.D.,50, 52(122) Hammes, G.G.,477 Hampton, J., 463 Handler, P., 301, 305, 306, 307(34), 310 (34), 312, 313, 314, 315, 317, 319, 320 (78), 323, 324(96), 325, 326, 343, 350, 351(77, 175), 353(78), 354(78), 355, 360(175), 361(175), 362, 366, 367, 368, 369(102), 386(236), 389, 390(102, 281), 391(102, 283, 287), 392(282), 393(96, 102, 175, 258, 281), 394(102, 175, 283), 395, 396(96, 102, 162, 175, 2831, 397(102, 175, 2821, 398(102, 1751, 399(102), 533(146), 534, 536 Haniu, M., 18,66,67(68,701,267 Hannuksela, M.,154 Hansen, R. E.,22, 23(50), 30, 56, 263, 517, 518(66), 519(66), 520(66), 521 (66) Harada, M., 446 Harbury, H. A., 89 Harding, B.W., 255,265,269 Hardy, R. W.F., 58, 59(7, 8), 60(8), 61, 62(8), 63, 64(7), 70(8), 82(8), 87, 90 (81, 99(7, 8), 100(8) Hargrave, K. H., 358 Harkin, J. M.,559 Harris, S.,359 Harsch, M.,153, 159(148, 149) Hart, L.I., 305,306, 307, 308, 309, 310(36, 421, 311, 312, 313, 317(36), 319(36), 323, 352(41), 357, 359(36, 210), 393 (36) Hartley, M. J., 413 Hartmann, G.,358 Hartmann, H. J., 535, 539, 543, 546(233, 2481, 548, 549(248) Hartmann, U., 424, 480(7), 493(7), 494 Hartree, E. F.,357, 464, 466, 507 Harta, J. W.,538, 541(218), 542, 543, 544, 545(218) Harvey, E. N.,227 Hasada, M.,527 Hashimoto, T.,137, 141(79) Hashimoto, Y.,271, 272(101) Hashimoto, Z.,120 Hastings, J. W.,105, 192, 227, 228, 246(1), 248(103), 251(103), 503, 504(218)

AUTHOR INDEX

Hasumura, Y ., 187 Hatano, H., 578 Hatchikian, E. C.,58, 59(9), 61, 62(9), 64(39), lOa(39) Hatfield, W.E.,547 Haussmann, E.,156,166 Hawkins, E.G.E., 184 Hayaishi, O.,120, 121, 124(8, lo), 125, 126, 127, 128, 129, 130, 131, 132, 133, 134, 135, 136, 137, 138(49, 50, 77), 139, 140, 141, 142, 143, 151(72, 781, 186, 193, 195, 199, 200, 201, 202, 206, 207(49), 226, 230, 241, 253, 256, 291 Hayashi, Y., 388 Hayashikawa, R., 317 Hayden, T.J., 312(66), 314 Hayes, M.B.,456 Hazen, E.,Jr., 77 Hedegaard, J., 255,259 Hedrick, J. L.,540 Heesen, T.C.,184 Heidema, J. K.,272,273 Helbert, J., 24 Hellerman, L., 446, 454, 455, 467, 468, 469,470, 471, 498(139) Hemmerich, P., 64,78, 90, 95, 99(51), 100 (121),102, 104(51), 108, 112, 244, 246, 248, 249, 250, 251, 319, 320(79), 327, 328, 342(118), 384, 399(271), 424, 425 (101, 471, 480, 482, 493(7), 494, 496 (1891,498(6, 1541, 501, 502(189, 213), 504(11), 505 Henderson, L. M., 145 Hendricks, R.H., 163 Henn, S. W.,445, 447(58) Hercules, D. M.,536 Herriott, J. R.,5,74,76(83), 78(83) Hersh, L. B.,498 Herskovitz, T.,8, 11(27b), 35, 37(27b), 43(106), 44, 46 Hesp, B., 211 Hevesi, L.,502 Hewitt, E.J., 309 Hewitt, J., 156 Heytler, P.G.,410 Hickey, R.J., 63, 65(49) Hiedemann-Van Wyk, D.,26 Higashi, N.,211, 212(63), 214 Higgins, I. J., 224 Higgins, R.,536

593

AUTHOR INDEX

Higuchi, K., 130, 131 Hildebrand, P. G.,540 Hildebrandt, A, G., 260, 261(33), 268, 271, 275 Hilkens, J. G. H., 213 Hill, C. M.,516, 517, 518(63), 524, 525(63, 67, 101) Hill, H. A. O., 282, 547 Hill, J. M.,514, 518(49), 519(49), 523, 525(96) Hill, R. L., 537, 543(199), 547, 557 Hinkson, J. W.,59, 60(2), 63(22), 65, 82 (601,83(60), 88(60), 99(22), 104(22) Hiramoto, M., 158(227), 159 Hirata, F.,131, 132, 142,202 Hiromi, K.,130, 138(49, 501, 139(50), 202, 578 Hirsch, J. G., 515 Hitchings, G. H.,302, 303(10), 345(10), 346(10), 348(10), 349(10), 350(10), 352(lo), 354(lo), 359( lo), 397(lo), 398(10), 401, 402(10), 408(10), 413 (10) Ho, P. P. K., 123,127(7) Hobza, P.,157 Hodgson, E.K.,536, 555 Hodgson, W.C.,333 Hofrichter, J., 91, 92(126), 93(126) Hogenkamp, H . P.C., 177 Hollins, R.A.,536 Hollunger, G.,468 Holm, R. H., 8,11(27b), 25, 35, 37(27b), 43,44,46,47,341 Holman, R. T., 150 Holmberg, C. G.,558, 560(311) Holme, E., 169, 174, 175, 176(336), 183 Holmes, M. C.,518, 519(73), 520(73) Holmquist, B.,8 Holwerda, R. A., 577, 578(431) Hong, J. S.,42, 45 Horecker, B. L., 529, 530(129), 531 Hori, K.,137, 141(79), 201 Horie, S.,265, 266,267 Horrocks, W.D., 43 Hosein, B.,26,27,28(67), 29(67), 45(67) Hoskins, D. D.,110, 115(166, 167) Hosoda, S.,291 Hosokawa, K.,211, 212 Houslay, M.D., 469 Howell, L.G.,31, 106, 108(160), 192,211,

212, 213, 225(4), 246(4, 5), 248(5), 399, 422, 472, 473(159), 534 Hsia, D. Y. Y., 179,181 (344) Hsu, C. A., 171, 172(324), 173(324, 325), 175(3251, 179(3251, 187(325) Hsu, M.,470 HSU, S.-T., 187 Huang, C. Y., 234, 236(125, 126), 251 (126), 289 Huang, H. S., 126 Huang, J.J., 19,23, 267,268 Huang, T.C., 55 Hbcko-Haas, J. E.,522, 523(93) Hughes, W.L.,Jr., 560 Hume, R.,266, 269(66) Hurych, J., 157 Hutchings, B. C.,410 Hutton, J. J., 151, 153, 154(133), 157, 158, 159(133) 160(223) 161 Hyde J. S., 96,344 Hysert, D.W., 227,249

I Ibers, J. A., 25, 35, 46(65, 97), 47, 341 Ichihara, A., 200 Ichikawa, Y.,272 Ichiyama, A.,241,291 Igaue, I.,467,470(131) Ignesti, G.,525 Iizuka, H., 258 Iizuka, T., 21 Ikai, A., 564 Ikeda, M.,240 Imai, Y.,271, 274 Inamasu, M.,469, 470(144), 522, 525, 526 (103) Infante, A. A.,162 Ingraham, L. L.,98, 99(141), 102, 473, 499 Inoue, H., 134 Inouye, K., 153 hie, K.,61,64(40), 70(40), 82(40>,88(40) Isaacson, E.L.,273 Isaka, S.,269 Isemura, T.,461 Ishii, H.,187 Ishimoto, M.,61, 64(40), 70(40), 82(40), 88 (40) Ishimura, Y., 129, 130, 133, 263, 264(53), 283(53)

594

AUTHOR INDEX

Isomoto, A., 476 Itada, N., 124(8, lo), 125, 140, 200 Itaya, K., 511 Ito, K., 445 Iwatsubo, M., 386, 419(272), 445 Iwayama, Y.,146 Iyanagi, T.,273 Izatt, R.M.,362

J Jacks, C. A., 14, 15(31) Jackson, J. F.,172, 173(326) Jackson, P. J., 8, 9(24), 10(24), 12(24) Jacobs, H.G.,165 Jacobson, M.,273 Jacoby, G. A., 180, 181(356) Jaaskelainen, M.,153, 154(162) Jakobsson, S. V., 270 James, D. W. F., 305, 312(40), 313(40) James, T. L., 82, 96(99), 97(99) Jefcoate, C . R. E., 266, 269, 275, 276 (1281,282 Jencks, W.P.,478 Jensen, L. H.,4, 5, 32, 33, 37(93), 60, 66, 70, 71, 72(78), 73(86), 74, 75, 76 (83),78(64, 83), 79(64, 781, 98(64), 114(64), 367 Jequier, E., 241,291 Jerina, D.M., 219,235,284 Jermyn, M.A., 466 Jezewska, M. M.,350, 401 Jimenez, M.H.,159,160(231) Jimenez, S.,153, 159(148, 149) Joester, K. E.,536,557 Joh, T.H., 240, 295 Johns, D.G., 353,354, 363, 389, 402(279) Johnson, A. R.,153(177), 154 Johnson, C: E.,8, 9, lO(24, 27a), ll(98, 98a), 12(24), 35, 37(98), 304, 341, 342 (159), 375(28) Johnson, D. A., 559, 575 Johnson, D.B.,316 Johnson, F. H.,188, 227 Johnson, J. L.,309, 414,415(52, 55a, 3341, 417(334) Johnson, P. W., 32, 34 Jolley, R. L.,296, 297(227) Jollow, D.,229

Joly, J.-G., 187 Jones, G., 309 Jonsson, M.,560,561(330) Jordan, S. R.,66, 71(69), 74(69), 75(69), 76(69), 77(69), 79(69), 80(69), 81 (691, 82(69), 101(69), 102(69) Jowitt, R. N., 323, 324(96c) Joyce, P.,312(65,65a),314 Jukes, T.H.,17 Juneau, M.,531 Jung, G.,536, 548, 557(190) Junk, K. W.,271, 272(100), 273(100) Jurtshuk, P.,280 Juva, K., 153, 157, 159, 161(164)

K Kadner, R. J., 170, 171(318), 174(318) Kagamiyama, H., 15, 140 Kahn, A. U.,536 Kainosho, M.,447 Kajita, Y.,201 Kaleita, E.,410 Kallio, R.E.,124(11), 125 Kamen, M.D.,32 Kamimoto, M.,136, 142, 143(74), 151(93) Kamin, H.,206, 207, 208, 213(52), 225 (531, 247, 273 Kanai, M., 126 Kanda, M.,316,319, 320(78), 353(75, 781, 354, 391(75), 394(75), 396(75) Kanetsuna, F., 134, 135(63), 140 Kang, H.H., 145 Kannagara, C.C.,26 Kao, K.-Y. T.,153 Kapeller-Adler, R.,512, 513, 516, 521 Kapit, R.,240 Kaplan, A., 156 Kaplan,N. O., 77 Kappas, A., 270 Kappe, T., 180, 181(348) Kappelman, A. H.,241, 291, 292(188) Karam, J. D.,411 Karkuen, P.,153 Karlsson, R., 76, 82(89), 102(89), 424, 425(21) Kaschnitz, R.M., 272,277 Kasha, M.,536 Kasper, C.B.,662,569 Kasperek, G.,501

AUTHOR INDEX

Katagiri, M., 120, 133(3), 134, 140, 146, 195, 206, 207, 208, 210, 255, 259(18), 260(18),262,283(39) Katz, E., 154, 165 Kate, J. J., 58(18), 59, 96, 97(18, 134), 99(18), 109 Kaufman, S., 158, 193, 232, 233, 234, 235, 236, 237, 238, 239, 240, 241, 242(13), 251, 288, 289, 290, 291, 292(188), 294, 295 Kawasaki, H., 516, 517(61), 518(61), 519 (61) Kawashima, N.,445 Kawashiri, Y.,158(227), 159 Kay, A.,324 Ke, B.,30, 31 Keams, D.R.,536 Kearney, E. B., 456, 470, 558, 559(309), 574(309) Kearns, D. R.,536 Keele, B.B.,358 Keele, B. B.,Jr., 535, 537, 538, 541(220), 542(2201,543, 544, 545 Kees, K., 536 Keilin, D., 357, 464, 466, 507, 508, 510, 533, 538(15), 555(15), 558(5) Keiser, H.R.,153 Kelly-Falcoe, F.,527(121), 528, 529, 530 (121, 129), 531(121) Kendall, D. S., 66,70(66), 71(69), 72(66), 73(66), 74(69), 75(69), 76(69), 77 (69), 78(66), 79(69), 80(69), 81(69), 82(69), 94(66), 98(66), 101(69), 102 (69),114(66) Kennaway, E. L., 325 Kennel, S.J., 32 Kenney, W.,470 Kent, A. B.,522,523(91) Kerber, R. C.,502 Kertesz, D.,296, 297(230) Kessler, D. L.,301, 414, 415(8, 3341, 417 (8,334) Kester, N.,235 Ketchum, P.A,, 403,404 Kevan, L.,23,24 Khanpaa, K., 153 Kielley, R. K., 388 Kierkegaard, P.,76, 82(89), 102(89), 424, 425(21) Kikuchi, Y.,158, 159(224)

595 Kimmel, J. R., 538, 542(229), 543, 545 (229) Kimura, T.,8, 19, 23, 24, 267, 268, 536 Kinemuchi, H.,470, 471 (1531, 498(153) King, L. A., 499 Kinoshita, T.,265 Kirk, T.K., 559 Kirkwood, R., 237, 251(136), 288, 289 (171) Kishida, Y.,153, 158(228), 159, 162 Kita, H.,136, 142, 143(74), 151(93) Kitao, T.,206 Kituchi, Y.,158(227), 169 Kivirikko, K. I., 153, 154, 155, 156, 157, 158, 159, 160, 161, 165, 166, 292, 293, 294 Kjellin, K., 561 Klassen, W. H., 533 Kleisen, C. M.,537 Kleppe, K., 448 Klinenberg, J. R.,302,401 Klingenberg, M.,254, 270 Kloepfer, H.C.,467,470(131) Klug-Roth, D.,541(295), 553, 554(293, 295), 555(295),556 (2951,557 Knappe, J., 19,58(16), 59, 62(16), 64(16), 90(16), 91(16), 99(16), lOO(16) Knappe, W.-R., 108, 246, 319, 424, 480, 494 Knight, E., Jr., 58, 59(6-8), 60(8), 61, 62 (8),63,64(7),70(8), 82(8), 87, 90(8), 99(6-81, lOO(8) Knight, S. G., 456 Knoell, H.E.,19 Knowles, P. F., 304, 315, 324, 327, 328, 331(99), 334, 337(99), 34108, 1221, 355, 368(69), 375(28, 99), 376, 377 (69), 384(197), 517, 518, 519, 520(73, 811,521(81), 525(71), 526(71), 534 Knox, W. E., 128, 129, 146, 179, 180, 181, 182, 183(346), 428 Kobayashi, K., 61,64(40), 70(40), 82(40), 88(40) Kobayashi, S.,124(8), 125 Kobayashi, Y.,153 Kobrle, V.,164 Koch, J. R.,124(11), 125 Kodicek, E.,152 Koenig, S. H.,548, 563, 571(357)

AUTHOR INDEX

Kohama, Y., 227 Koike, K., 128, 129(36) Kojima, Y., 124(10), 125, 134, 135, 136, 137, 138(77), 139(77), 140, 151(72), 200 Komai, H., 305, 307(35), 308(35), 310 (351, 311(35), 312, 313(35, 63), 314 (35), 315(61), 316, 318, 319, 320, 327, 328(35, 631, 337(110), 344(63), 352 (631, 353(35, 631, 354(61, 63), 355 (631, 361(35), 362(35), 363(61), 364 (61), 387(61, 631, 368(35), 369(35), 372(35), 374(35, 611, 381(35), 387 (61), 393(63) Komiyama, T.,140 Korkes, S., 499 Kornblum, N.,502 Kosman, D.J., 530,532 Kosower, E. M.,424, 471, 481, 497 Koster, J. F.,448,451,454 Kotake, Y.,127 Kotaki, A., 446, 447 Kotani, S.,133, 135, 136, 140, 141, 161(72) Kotowycz, G.,212 Kovensky, A,, 401 Krauss, P., 311, 328(60), 341(60), 538, 540(219), 542(219), 543, 545(219), 546(219, 2481, 548(248), 549(248) Kraut, J., 32, 33, 35(90-92) Krebs, E.G.,522,523(91) Krenitsky, T.A., 302, 303, 345, 346, 348, 349, 350(10), 352(10), 354(10), 359, 389, 397(10, 279a), 398(10,279a), 402 (101, 408,412,413(10) Krimm, S., 27 Krishnamurty, H.G.,125 Krodel, E.,486,489(186) Krueger, R.C.,146 Krupnick, D., 537 Krusic, P. J., 46 Kubo, H.,445 Kubowitz, F.,254 Kuczenski, R.T.,239,290,291 Kumagai, H.,516, 517(61), 518(61), 519 (61),522 Kumar, R. P.,124(9), 125, 534 Kunitake, T.,501 Kuno, S.,124(8), 125, 130 Kuntsman, R.,273 Kuroda, M.,476

Kurtz, J., 158, 160(223) Kust, R.N.,499 Kuttan, I., 152 Kuttan, R.,164 Kwiatkowski, L. D.,532 Kwietny, H.,345, 346(164), 348(164), 349(164) Kyogoku, Y.,447 1

Labeyrie, F., 386,419(272) LaDu, B. N.,179, 180, 181,238, 254, 269 (7) Laishley, E. J., 32 Lamar, G.N.,43 Lambert, P.,556 Lamberti, A.,404, 405(304) Lambeth, D.O.,29,103 Lamport, D.T.A., 163 Land, E. G.,328 Land, E.J., 90 Landis, W ., 284 Lang, G.,53, 55(131) Lang, H.M.,61 Langner, R.O.,153, 154 Langness, U., 153, 163 LaNoue, K. F.,89 Lash, S. W.,153 Laskowska-Klita, T.,180, 181(350, 353) Latimer, W. M.,499 Laurell, C. B.,558, 560(311), 571 Lavelle, F.,534 Lazanas, J. C.,524,525(100) Lazarides, E.L., 162 Laazarini, R. A,, 107, 108(161a) Lebar, R.,204 Lebeault, J. M.,280 Lebherz, H.G., 315, 389(68a) Lee, B.,77 Lee, K.-Y., 309, 403, 404(302), 405(304306) Lee, M.H.,560, 561(324), 562, 564(345), 567(345), 571(345) Lee, R.. F.,174, 186(334) Lee, T.-C., 352, 355(182), 361(182) LeGall, J., 32, 58, 59(9-111, 60, 61, 62(9111, 64(39), 66, 67(67), 69(67), 70 (291,82(11), 99(ll), 104(39) Legg, K. D., 536

AUTHOR INDEX

Lehrmann, Ch., 270 Leibman, K. C., 268,275 Leigh, J. S., 342 Leigh, S., 270 Leijonmarck, M., 76, 82(89), 102(89), 424, 425(21) Le May-Knox, M., 180 Lemberg, R., 259, 263(26), 264(26) Leo, A., 463 Leone Ganado, A., 546 Leong, J. L., 270 Le Peuch, C., 228, 248(103), 251(103) LeQuesne, M. E., 66, 70, 71(65), 72(66), 73(66), 78(66), 83, 91(105), 94(66), 98(66), 114(66), 447 Lerch, K., 297 Lerner, A. -B., 296 Leterrier, F., 58, 59(10), 62(10) Le-Thi-Lan, 204 Levenberg, B., 158, 288, 294 Levene, C. I., 153 Levene, L., 345, 346(166), 348(166), 351 (166) Lever, A. B. P., 7 Levi, A. J., 466 Levin, B., 401,402(295a) Levin, E. Y., 158,294 Levin, G., 345, 346(164), 348(164), 349 (164) Levin, S. S., 266, 268(68), 270, 283(68) Levin, W., 270, 273 Levine, C. I., 162 Levine, W. G., 574 Levitt, M., 238, 239(138), 240(140), 290 Levy, C. C., 217 Lewis, D., 64, 110 Lewis, N., 412 Lhoste, J.-M., 91, 93(124), 102(124), 115 ( 124), 248, 424,446 (20) Liberman, U. I., 359 Lichtenberger, F., 279 Liebelt, A. G., 153(180), 154 Lieber, C. S., 187 Lieberman, R. A., 542, 546(243) Lieni, H. H., 262 Lilie, J., 557 Lilienthal, H. R., 563, 571(357) Liljas, A., 73, 77 Lin, E. C. C., 181 Lin, S., 173, 174(329), 175(329), 179(329)

597 Lindblad, B., 169, 170, 180, 182, 183, 184 (313), 293 Lindskog, S., 550 Lindstedt, G., 167, 168, 169, 174, 175, 176 (3361, 177(332), 178(332), 182, 183, 184, 185, 186(306), 293 Lindstedt, S., 167, 168, 169, 174, 175, I76 (3361, 177(332), 178(332), 180, 182, 183(311, 351), 184(313), 185, 186 (3061, 293 Lindstrom, A., 517, 518, 519(72), 520, 521 (72, 761, 522, 523(72), 524(92), 525, 526 (110) Lindy, S., 153, 154 Linnane, A. W., 537 Linneweh, W , 167 Lipscomb, J. D., 260, 261(31), 264, 268 (31, 55), 282(54) Lipton, M., 290 Liu, C. K.,-171, 172, 173, 174(329), 175, 176(328), 177(328), 178, 179(325, 3291, 189(325) Liu, T. Z., 186 Ljones, T., 50, 52(122), 53, 55 Ljunggren, H., 574 Llinas, M., 4 Loach, P. A , 89 Lobley, R. W., 524, 525(101) Locke, R. K., 186 Lockridge, O., 188, 195, 196(23), 197, 199, 201(23), 203(23), 204(24, 261, 431, 498 Lode, E. T., 6,8, 13, 45, 46 Lombardini, J. B., 125, 149, 150(19) Lorenzi, D. J., 158(226), 159, 160(226) Lorenzoni, E., 321 Lovell, F. M., 561,567(338) Lovenberg, W., 4, 5, 6, 7(17), 8, 9(17), 12, 13, 14, 15(31), 24(23), 33, 241, 291 Low, B. W., 561, 567(338) Lowe, D. J., 50, 54(123), 304, 321, 322, 330(32), 331, 332(127a), 336, 338(32), 339, 340(127a, 1531, 341(153), 343, 344, 357, 369(90b), 372(32), 374, 375 (1531, 377(32), 378(154, 265), 388 (154), 389, 391(90b, 286), 393(90b, 286), 394, 395(90b, 91a, 2861, 396(90b, 286), 397(90b), 399(90b, 91a, 2881, 547(294), 553, 554 (2941, 555(294), 556(294), 557(294)

598 Lowe, H. J., 100, 356 Lowry, 0.H., 362, 363(240) Lowther, P.A., 156 Lu, A. Y. H.,271, 272(100), 273 Ludwig, M. L., 60, 66, 70, 71(28, 65, 69), 72(66), 73(66), 74(69), 75(69), 76 (69), 77(69), 78(66), 79(69), 80(69), 81(69), 82, 83, 91, 92(126), 93(126), 94(66), 96(99), 97(99), 98(66), 101 (691, 102(69), 114(66), 447 Lukens, L. N., 153, 159, 161, 162 Lindgren, E.,537,543(204) Luthy, J., 440 Lynden-Bell, R. M., 339, 340(153), 341 (1531, 343(153), 374(153), 375(153), 378(153), 388(153) Lynn, S., 103

AUTHOR INDEX

McGee, J. O’D., 154, 159, 160(231), 162, 163 McGeer, P. L., 466 Machinist, J. M.,229 McKee, D. J., 562 McKee, E. M.,229 Mackenzie, C. G., 110, 115, 117(177) MacKensie, R.E., 93 Mackler, B., 393 MacKnight, M. L., 77, 83(90), 94, 99 (131) McMahill, P., 296, 297(225) Madrid, V. O.,178, 179 Maeno, H.,128, 129, 206 Magdoff-Fairchild, B. S., 561, 567(338) Mager, H.I. X., 241, 242, 243, 245, 251 (157) Mahler, H.R., 373, 393, 511, 537 M Mailer, C.,24 Main, L., 501 Ma, D., 239, 291 Maitra, U., 189 McCapra, F.,227,249 Maki, Y.,200, 201, 226 McCarron, M., 411 Makinen, M. W.,91, 92(126), 93(126) McCarthy, J., 19 Makino, N.,560, 561(326), 564, 570(368), McCarthy, K.,4 571, 572(368) McCay, P.B., 358 McCord, J. M., 354, 357(185), 366, 510, Makita, T., 200 533, 534, 535, 537, 538, 539, 540(16, Malkin, R., 29, 30, 45, 295, 509, 564, 565 (361), 566(10, 3611, 569(360, 361, 2321, 541(16, 2201, 542(16, 2201, 543 3631, 570(10, 360, 361, 363, 3641, 571 (2201, 544(220), 545(220), 548(16), (10, 3641, 572(363, 3641, 573(363, 549(16), 551(16), 552(16), 556(16), 3641, 574(364), 576(10), 579(10) 557(232) Malrnstrom, B. G., 295,304,310, 311, 327 McCormack, J. J.,234,349 (24,25), 329, 342(25), 509, 521, 546, McCormick, D. R., 88, 93,445, 446, 447 547(254), 562, 563, 564, 565(347, 353, McCroskey, R. P., 170, 171(316, 317, 361), 566,567(254,347, 353), 569, 570, 319-3211, 172, 173, 174(329), 175(317, 571(10, 356, 3641, 572, 573, 574(364, 319, 320, 328, 3291, 176(316, 317, 319, 392, 3931, 576, 577, 578, 579 320, 3281, 177(316, 317, 328), 178(317, Manabe, T., 578 3281, 179(329), 187(320) 456 McDonald, C. C., 6, 7(17), 9(17), 12(17), Mrtnchon, P.H., Mandansky, C . H., 403 33, 96 MacDonald, D. W.,405, 406, 413(307b) Mandell, A.J., 239,290 Mann, P.J. G., 514,518(49), 619(49), 523 McElroy, W.D.,227 Mann, T.,508,510, 533,538(15), 555(15) MacFarlane, H., 516,521 McGartoll, M. A,, 305, 306(36), 307, 308 Mannering, G. J., 271,279 (361, 309(36), 310, 311, 312(36), 313 Manning, J. M., 292 (361, 315(48), 316(48, 58), 317(36), Manning, T., 239,291 319(36, 48), 323, 324, 325(98), 329 March, W.F.,470 (981, 330(98), 337(58), 359(36), 374 Marchesini, A., 560, 562, 564(346), 565 (58), 382(98), 388(58), 393(36) (3461, 566(346), 567(346), 568(346), McGavack, T. H.,153 571 (346)

AUTHOR INDEX

Marglin, A., 158, 160(223) Margolis, R.L.,153 Margulis, L.,537 Markel, P.D., 536 Marklund, S.,325, 537 Markowitz, H.,533, 538, 541(215), 542 (2291, 543(229), 545(229), 548(215) Marmur, J., 537 Marov, I. N.,333, 334, 335, 339 Marshall, V.,264, 282(54) Martin, D.F.,526 Martin, R.L.,53 Marsluff, W.F.,333 Masayama, T.,127 Mason, H.S.,120, 146, 182, 254,257, 269, 271, 272, 273, 282, 296, 297(225), 355, 368(191), 519, 520(83), 521(83), 574, 576(412) Mason, J. I., 19 Mason, R.,32,34(86) Massey, V.,31, 56, 58,59(12), 60, 61(12), 62, 63(12), 64(12), 65, 66, 67(70), 71 (281, 79(42), 89, 90, 91, 93(124), 95 (114, 1211, 96(120), 98(58), 99(58), lOO(58, 1211, 102, 103, 104(149), 105, 106, 107(58), 108(42, 63, 160),' 112, 113(179), 114(179), 115(124, 179), 116(63, 179), 188, 192, 193, 194(10), 195, 196(23), 197(23), 199, 200, 201 (23, 321, 203, 204(24, 261, 210, 211, 212, 213, 214, 215(73), 216(69), 217, 218(77), 220, 221(73), 225(4), 246 ('2-5), 247,248,249,300, 301,305,306, 307(35, 44), 308, 310, 311, 312, 313, 314(35, 46), 315, 316, 317, 318(63), 319, 320, 324(44, 621, 325(441, 326, 327, 328, 329(117), 331, 332(128), 336, 337, 338, 339(117, 128), 340(45), 341, 344(1, 63, 1281,345, 352(44, 63), 353, 354, 355, 356, 361, 362,(35, 45), 363, 364, 365(45), 367(61, 631, 368, 369, 371(46), 372, 374(35, 4446, 61, 117, 128), 375(46), 376(46, 128), 377(128), 378(46), 379(46), 380(46), 381(35, 46), 382(44, 461, 383(45, 46), 384(45, 46, 201), 385(45, 461, 386(46), 387 (46,61, 117), 388, 389, 390(77c), 391 (77~9,293(63, Ssb), 398(77c), 399, 405(46), 422,424,425(13, 18, 191, 427, 430, 431, 432, 433(43), 434(25), 435

599 (25,36, 381, 436(36), 437(25, 38), 438 (25),440, 441(19, 25, 361, 442(25, 36, 38), 443(36), 444, 445, 446, 447, 448, 449, 451(36), 452, 454(36), 455, 457, 458, 461, 462(25), 463(25), 465(25), 472, 473, 476(25, 36, 381, 477(13, 251, 479, 480(36, 38), 481, 482, 484(19), 486, 489(171, 186), 498, 499, 500(202), 501, 502(213), 504, 505, 534, 553 Masters, B. S. S., 268, 273 Matheson, M. S.,357, 555 Mathews, R. A.,40, 41(102), 42(102), 43 (102),45(103a) Mathews, R. G.,31 Matsubara, H., 15,17 Matsueda, G.,66 Matsui, H.,516, 517(61), 518(61), 519(61) Matthews, R. G.,106, 108(160), 192, 225 (4), 246(4, 5), 248(5), 399, 422, 472, 473, 534 Mauger, K. K., 153(177), 154 Mautner, G. N.,357, 547(294), 553, 554 (2941, 555(294), 556(294), 557(294) Max, E.E.,234,236(125), 289 Maxwell, J. C.,534 May, F.,309 May, S. W., 533(152), 534 Mayer, V.W., 186 Mayerle, J. J., 25, 43(106), 44, 46(65, 106), 341 Mayhew, S. G., 19,31,58,59(12,13), 60,61 (12),62,63, 64,65,66,67(68, 70), 70, 71(28, 651, 72(66), 73(66), 78(66), 79 (42), 82(13, 591,83,84(13, 53, 591, 85 (59),86(53), 87(59),88,89,90(12,13), 91, 92(13), 93(59, 124), 94(66), 95 (114), 96, 98(13, 58, 66), 99(13, 581, 100(13,58),101,102,103,104,105,106, 107(58),108(42,53,160),110,111,112, 113, 114(66, 179-1811, 115(124, 170, 178-181), 116(170, 178, 179, 1811,117 (170, 1781, ll8(170), 192, 225(4), 246 (4,5), 248(5), 356, 399, 422, 424, 446 (20),447,455, 480, 534 Meany, J. E., 345, 346(170), 349(170) Medina, H., 527, 531(119) Mehler, A. H.,121,128,145,296 Meighen, E.A., 227 Meister, A., 156, 292, 456, 457, 499 Melandri, B. A., 31

600 Melin, M., 560 Meriwether, L. S., 306, 310(41), 316, 324, 330(73), 331(99), 333, 334, 335, 337 (99), 352(41), 375(99) Metz, E., 401 Meyer, T. E.,32 Michaliszyn, G.A,, 227 Michelson, A. M., 534,537 Midg&t, R. J., 170, 171(316), 174, 176 (3161, 177(316), 186(334) Midvedt, T.,167, 168, 169(305) Mihara, K.,146,206,208 Mildvnn, A. 5, 98 Miller, E., 153(178), 154 Miller, F. K.,163 Miller, R.,167 Miller, R.L.,162,165,293 Miller, R. W.,326, 366, 367(251), 358 (1071,538, 547(228) Milligen, L. P.,110, 117(172) Millington, R. H.,179 Milne, M. D.,466 Mimoun, H.,186 Miskowski, V.,566,567(380) Misra, H.P.,356, 358, 533(148, 149), 534, 537(153), 538, 539, 540(153, 232), 541 (2271, 542(227), 543(227), 544, 545 (2271,557(232) Mitani, F., 265, 266, 267 Mitani, S., 445 Mitchell, C.H.,229 Mitchell, H.K.,170, 173, 175(323), 178, 407, 409(312), 411(312) Mitchell, J. R., 279 Mitchell, P.C.H., 323, 324, 336, 337, 338 Mitchell, R.A., 145 Mitoma, C.,232,254,269(7),291 Miyake, Y.,134, 135, 136, 143(74), 272, 484 Miyamoto, T., 127 Mochnacka, I., 180, 181(353) Moldeus, P.,277 Monacelli, F.,125 Monaco, M. E.,129 Mondovi, B.,148, 516, 517, 518(66), 519 (661, 520(66), 521, 522, 523(89), 525, 526(106), 535,536(178),540,542(241), 543(241), 545, 546, 547(251, 2551, 548 (241, 255), 549(241), 550, 551, 555 (255),556(255), 560,567

AUTHOR INDEX

Montgomery, R., 531 Moon, H.M., 411 Moore, T. A., 92(127), 93, 94, 350, 351 (178), 368(178) Moras, D., 73, 447 Morell, A. G.,561, 565, 566, 571(376), 575 Morell, D.B.,307, 318(47), 374(47), 387 (47),‘397,398(291) Morgan, P. H.,165 Mori, M., 124(10), 125 Morpurgo, L.,546,547(255), 548, 551, 552 (2841, 554(284), 555(255), 556(255, 267), 567 Mortensen, L. E., 39, 47, 52, 53, 54(134), 55 Mortenson, L. E., 58, 61(2), 65 Moss, T. H.,19, 32(37), 563, 566, 567 (3801, 571 (358) Muller, F.,66, 78, 87, 89, 90,95(114, 117, 1211, 96, 100(121), 105(63), 106(63), 108(63), 116(63), 192, 213, 245, 246 (51, 248, 249, 251, 327, 328, 399, 422, 424, 425(18), 455, 480, 493(7), 494, 496(189), 502(189), 504(11), 505 Muller-Eberhard, U., 262 Miinck, E., 263,264,282(54) Munck, M., 143 Mukai, K., 19, 23, 24 Mulac, W.A., 357, 555 Mulford, D.L.,560 Mulhern, J. F.,350 Mulliken, R.S.,482 Mullinger, R.,37 Multani, J. S., 50, 52(121), 53(121) Mulveny, T.,166 Munck, E.,10(97b), 35 Murakami, K.,271 Murakami, T.,258 Murphy, L.,153, 159(149) Murphy, P.J., 187,280 Murray, C.L.,45,46(114) Murray, K. N., 355, 367(190), 368(190) Murthy, V. V., 297 Mussini, E.,153 Myer, Y. P.,566 Myles, A., 352, 355(182), 361(182)

N Nagami, K., 134, 135, 136(65, 71) Nagate, T.,522

601

AUTHOR INDEX

Nagatsu, T., 238, 239, 290, 527 Nagelschneider, G.,424 Nagler, L. G.,314 Naik, V. R.,557 Naito, A., 258 Nakagawa, H.,134 Nakai, Y.,195 Nakamo, M.,456 Nakamura, M.,207, 208(58), 210, 322, 354,356, 367(186) Nakamura, S., 186, 211, 212(63), 214, 241, 291, 355, 356, 432, 434, 435(48), 436 (451, 440(45), 441(45), 448(45), 449 (45), 451(45), 452(45), 458(45), 463 (481,465,476(48),481(45) Nakamura, T.,130, 195,207, 208(58), 210, 432, 434, 436(45), 440(45), 441(45), 448, 449(45), 451(46), 452, 458(45), 462, 463, 465, 471, 481(45), 482(82), 560, 561(326), 564, 572, 576 Nakano, G., 527 Nakano, H.,268 Nakazawa, A., 134, 135(63), 136, 142 Nakazawa, T.,134, 135, 136, 140, 141(85), 151(72), 201 Nakacowa, K., 195 Nakos, G.,65 Nalbandyan, R. M.,551, 552(282), 554 (282) Namtvedt, M. J., 22, 23(49), 263 Naoi, M.,446,447 Nara, S.,468 Narasimhulu, S.,270,274 Nason, A , 309, 403,404 Nastainczyk, W.,286, 287(159) Natter, R., 543, 546(246), 549(246) Neims, A. H.,446,454,455 Nelson, C. A,, 305, 306(34), 307(34), 310 (34),312,313,314,315 Nelson, D.H., 265 Nelson, E.B.,273 Nelson, J. M.,296 Nelson, N., 62, 79(43), 108(43) Nelson, R.M.,296,297(227) Netter, K.J., 268 Neuberger, A., 466 Neujahr, H.Y.,221,223(82) Neumann, J., 62, 79(43), 108(43) Newman, D. J., 32 Newman, W.F.,156

Niel, S. M.,302, 303(10), 345(10), 346 (lo), 348(10), 349(10), 350(10), 352 (lo), 354(10), 359(10), 397(10), 398 (lo), 402(10), 408(10), 413(10) Nielsen, S. O., 356, 553, 656(296) Nieman, Z., 345, 346(166), 348(166), 351 (166) Nilason, L.-O., 540 Nishibayashi, H.,275, 276( 124) Nishikimi, M., 444, 448, 449,450, 452, 477 Nishino, T.,389, 390(284), 391(284) Nishizuka, Y.,128, 241, 291 Nordwig, A., 153, 154(168), 159, 164 Norman, R.0.C., 186 Norrestam, R.,76, 82(89), 102(89), 424, 425(21) Norris, J. R., 58(18), 59, 96, 97(18, 134), 99(18), 109 Norris, R.K.,502 North, J. C.,269 Norton, S. J., 235 Nose, K.,511 Notton, A.,309 Novello, F.,322, 389(95), 402(95) Nocaki, M.,129, 130, 133, 134, 135, 136, 137, 138(49, 50, 77), 139(50, 77, 78), 140, 141, 142, 143, 151(72, 78), 193, 201, 202, 226, 230 Nygaard, A. P., 83 NylBn, U., 514, 515, 519(50), 520, 521(50), 523(501,525, 567, 570, 575 Nyman, P.O.,539 Nyns, E.J., 123, 127(7)

0 Oae, S., 206 Ochiai, H.,140 Oda, H.,476 O’Dell, B.L., 527 Ogasawara, N., 145 Ogata, K., 513, 514(43), 516, 517(61), 518,(43, 61), 519(61) Ogura, Y.,214, 432, 434, 435(48), 436(45), 440(45), 441(45), 448, 449(45), 451 (45), 452(45), 458(45), 462, 463, 465, 471, 476(48), 481(45), 482(82), 513, 518(39), 519(39), 521(39), 523(39), 525(39), 526(39), 560, 561(326), 564, 570(368), 571, 572(368) Ohama, H., 447,448

602 Ohesson, J. T., 470 Ohishi, N., 444, 448, 449, 450, 477t8.3) Ohnishi, T., 56 Ohta, Y., 223, 224 Ohyama, T., 538, 539(221), 540(221), 541 (2211, 542(221), 543(221), 544(221), 545(221, 2221, 546(221), 547(221) Oi, S., 469,470,525,526 Oka, T., 125 Okada, K., 158(227), 159 Okamoto, H., 226, 230 Oki, T., 516 Okuno, S., 130, 138(50), 139(50) Olander, B., 169, 184 183(351) Oleson, J. J., 410 Oliver, I., 359 Olomucki, A,, 203, 204 Olsen, B. R., 153, 155, 156, 162,292, 293 Olsen, J. S., 56 Olsen, K. W., 73, 447 Olson, A. C., 163 Olson, J. A., 126 Olson, J. S., 306, 313, 314(46), 326, 327 (461, 328(45), 331(46), 336(46), 337 (46), 338(46), 340(45), 341, 353(45), 355(45), 356(45, 461, 362(45), 364 (45), 365(45), 368, 369(46), 371, 372, 374(45, 461, 375(46), 376, 380, 381 (46), 382(46), 383, 384, 385, 386, 387 (46), 388, 399, 405(46) Olsson, B., 518, 520(76), 521(76), 525 Oltrik, R., 414 Omfeldt, M , 180, 183(351) Omura, T., 255, 259(12), 265, 267, 268 (591, 270, 271, 273, 275, 276(12, 13, 124), 561, 565 Ondega, B., 534 Ondrejickova, O., 164 Ono, K., 133, 140, 141(62, 851, 143 Ooshima, A., 153, 154(174) O’Regan, C., 316, 319(75b), 353(75b), 394(75b), 396(75b) O’Reilly, S., 575 Oreland, L., 468, 470, 471(153), 498(153) Orme-Johnson, N. R., 56 Orme-Johnson, W. H., 4, 10(66), 22, 23 (501, 24, 26, 32, 34(85), 39, 40, 44, 47, 49, 50, 261, 262(37), 263, 266, 269 (67), 280(37), 339, 341, 356, 369(156), 353(150), 534

AUTHOR INDEX

Orrenius, S., 270, 274, 277 Osaki, S., 559, 560, 575,576 Osamura, M., 477 Osborne, M. J., 531 Ottnad, M., 548 Owen, E. C., 357,359(210) Oyamburo, G. M., 358 Orawa, T., 455, 446, 479

P Pachowsky, H., 287 Packer, E. L., 44 Pankalainen, M., 155, 157, 293 Page, D. S., 440, 460, 461, 483 Page, M. I., 478 Palaith, D. H., 24 Palleroni, N. J., 128 Palmatier, R. D., 170, 171(316, 3211, 173, 174(329), 175(329), 176(316), 177 (316), 179(329) Palmer, G., 4, 6, 7,8, 10(66), 19, 20, 21, 22, 23, 24(23), 25(16), 26, 28(57), 29, 31, 32(37), 33(16), 39, 40, 41(102), 42 (102),43(102), 44(16), 52, 53,54(134), 55, 56, 90, 95(121), 96(120), 98, 100 (1211, 103, 105, 106(158, 1591,192,246 (2, 31, 247, 248, 263,301, 304, 305, 306, 307(35, 441, 308(35, 44), 310(34 44), 311(35, 44), 312, 313(35, 45), 314(35, 461, 315(44, 611, 317(44), 324(44), 325, 326(44, 46), 327, 328, 329, 331, 332(128), 336(46, 1281, 337(46, 110, 117), 338(46), 339(117, 128), 340(45), 341, 343, 344(128), 352(44), 353(35, 45), 354(61), 355, 356, 361(35), 362 (35, 4.9, 363(61, 110), 364(45, 611, 365(45), 367(61), 368(2, 35, 44, 46), 369, 371(46), 372(2, 31, 35, 45, 2631, 373(263), 374(35, 44-46, 61, 117, 128, 2631, 375(46, 128), 376(45, 128, 263), 377(128, 263), 378(46), 379(46), 380 (46), 381(35, 46), 382(44, 461, 383(45, 46, 263), 384(45, 46, 117, 201), 385(45, 46, 263), 386(46), 387(46, 611, 388(45, 46), 389, 391(283), 394(283), 395(162, 2831, 396(162, 283), 399(46, 2011, 405 (461, 422, 424, 425(13, 181, 444, 445, 446(13), 449, 477(13), 504, 533(147), 534, 553 Pan, S.S., 309,404,405(304-306)

AUTHOR INDEX

Pandey, J., 411 Parker, H. E., 527 Parker, R., 402 Parker, T. S., 173, 174, 175(329), 179 (329), 186(334) Parzen, S. D., 407 Paschen, W., 535, 536, 540(179) Patek, D. R., 467, 468, 469, 470, 471, 498 (139) Patel, K. B., 556 Patel, M. D., 143 Pateman, J. A., 402, 412 Patil, S. S, 296 Patrick, R. S., 162 Pawlik, R. T., 336, 389, 391(286), 303 (2861, 394(286), 395(286), 396(286) Pearson, A. J., 242, 244, 251(162) Pecht, I., 384, 399(271), 577, 578(432, 433) Peck, H. D., 32,60 Peel, J. L., 61, 65 Peisach, J., 2, 22, 139, 261, 262(37), 280 (37), 414, 529, 530(129), 574 Peltokallio, P., 154 Penton, Z., 565 Penzer, G. R., 424 Petering, D. H., 19, 22, 23, 26, 32(37) Peterkofsky, B., 153, 156, 157(103, 165), 160, 161(1651, 162, 185(157) Peters, J. M., 325 Peterson, E., 120, 254, 296(2) Peterson, J. A., 14, 130, 260, 261, 263, 264 (53), 282(38), 283(53) Peterson, T. C., 533(151), 534 Petersson, L., 37 Petrack, B., 239,291 Pettersson, E., 560, 561 (330) Pettersson, G., 517, 518, 519(72), 520, 521 (72, 79), 522, 523(72), 549(92), 525, 526, 567, 570, 575, 576(420) Pettersson, I., 575, 576(420) Pettersson, R., 304, 307(26), 309(26), 311 (26), 329, 341(26), 342(26), 361(26), 369(26), 370(26), 372(26, 30), 374 (26), 387(26) Pfab, F. K., 153, 154(168), 159, 164 Phillips, W. D., 6, 7(17), 9(17), 12(17), 25, 33, 43, 44, 46, 96, 341 Philpot, G. R., 400 Pho, D. B., 204 Pick, F. M., 310, 311(58), 315, 316(58),

603 323(58), 324, 325(98), 327, 329(98), 330(98), 331, 334, 337(58), 338, 355, 368 (691, 374 (58), 377 (69, 148a), 382 (98), 384(197), 388(58, 129, 148a), 534 Pickard, M. A., 559 Piette, L. H., 8, 12(26), 317, 576 Pionetti, J. M., 516 Pistorius, E. K., 150 Pitt, B. M., 181,428 Poe, M., 6, 7(17), 9(17), 12(17), 33, 43 Poillon, W. N., 128, 129, 562 Pollard, J. K., 163 Polyakova, N. A., 112 Pomales, R., 302 Pomerantz, S. H., 296, 297 Popenoe, E. A., 158, 165, 166, 167(281), lsS(219) Porter, D. J. T., 428, 429(33), 430(33), 431, 432, 434(46), 435(39), 437(33), 439(33), 442(33, 391, 443, 444(33), 445(33, 391, 449, 450(39), 451(33, 39), 452(39, 461, 453(39), 454(87), 456, 457(33), 459, 461(33, 531, 473, 476, 479(46), 480(30, 39, 531, 481(165), 482(46, 165), 483, 485, 486(39), 488 (39, 185), 490(39), 491(39, 185), 492, 494(165, l87), 495(39, 165, 187), 496 (39, 1651, 497(165), 501(165), 502 (165, 182) Porter, H., 533, 538(141) Postgate, J. R., 32, 34(86), 50, 51(119), 53(119) Poulson, L. L., 229 Poyer, J. L., 358 Prego, C. E., 358 Prema Kumar, R., 225,247 Previll, J. M., 153 Prichard, P. M., 162 Priest, D. G., 358(226), 359, 360(232), 361, 369(226) Priest, R. E., 145, 185 Prijs, B., 250(181), 251 Prinz, R., 538, 540(219), 541(226), 542 (219), 543(219, 2261, 545(219, 226), 546(219) Prockop, D. J., 152, 153, 154, 155, 156, 157, 158, 159, 160, 161, 162, 163(195), 164(185), 165, 166, 167, 292, 293, 294 Prodamov, E., 358 Proudfoot; R.;357, 359(210)

604

AUTHOR INDEX

Prynne, C. J., 153,162 Puget, K.,537 Purvis, J., 19

Q Que, L., Jr., 46,47

R Rabani, J., 356, 357, 541(295), 553, 554, 555, 556, 557 Rabinowita, J. C.,42, 44, 45, 46(114), 62, 65(41) Radda, G.K.,424, 461 Radhakyishnan, A. N., 152, 164, 499 Radlick, P., 536 Raheja, M. C.,183 Rajagopalan, K. V., 301, 309, 312, 316, 319, 320(78), 321, 322, 323, 324(96), 325, 326, 343, 350, 351(175), 353(75, 781, 354(78), 360(175), 361(175), 367, 368, 369(103), 388(87a), 389, 390(103, Bl),391(75, 103, 283,287), 392(282), 393(96, 103, 175, 258, 281), 394(75, 103, 175, 283), 395(162, 283, 287), 396 (75, 96, 103, 162, 175, 283), 397(103, 175, 282>, 398(103, 175), 399(103), 414, 415(52), 416(337), 417(8, 334, 3371,418(337), 533(144), 534,537 Ramaley, P. B., 153 Ramseyer, J., 269 Rando, R. R., 498 Rao, K. K.,8, 9(24, 27a), lO(24, 27a), 11(98), 12, 15, 17, 24, 35, 37 Rao, N. A., 124(9), 125, 452, 534 Rao, N. V., 164 Rao, S. T.,67 Rapaka, R. S., lSS(2291, 159 Rapp, U.,538,547(228) Rasmussen, C.,517, 525(71), 526(71) Raub, A., 514 Ravindranath, S. D.,124(9), 125, 225, 247(88), 534 Rawitch, A. B.,456 Rawlings, J., 24 Raymond, W. N., 14,15(31) Raynor, J. B.,329 Reed, D.J., 525, 526(104)

Reeves, S. G., 30 Reichard, P., 177 Reifsnyder, C. A.,234 Reindars, L.,537 Reinhammar, B., 552, 560, 561 (328-3301, 562, 563, 564(346, 347, 353), 565(346, 347, 3531, 566, 567(346, 347, 353), 568 (3461, 569(377), 570(287, 288, 347), 371(287, 288, 3461, 372(287, 288) Remmer, H., 271, 274, 275(119), 276(119) Rencovi, J., 157 Rensen, J., 235 Rever, B. M.,402 Reanik, E.M.,333,339(136) Rhoads, R. E., 154, 155(191), 157(191), 158, 159, 160, 161(191, 2411, 184, 292, 293 Ribbons, D. W.,223,224 Rich, A., 76,447 Richards, F.M.,77 Richardson, D.C.,542, 557 Richardson, J. S , 542, 557 Richardson, W.H., 184 Riehert, D.A.,303,320,410 Rigo, A., 550, 551 Rimai, L.,6 Rinker, R. G.,103 Rittenberg, S. C.,147 Riveito, C.,536 Riviero, C.,358 Rivkin, I., 238 Robb, D.A.,296,297(227) Roberts, D.B.,402 Roberts, N.,153 Roberts, P. B., 357, 547(294), 551, 552 (263), 553, 554(263, 283, 286, 294), 555(263, 2941, 556(294), 557(294) Robertson, W. van B., 156 Robinson, D.S.,241 Robinson, J., 427 Roche, B.,280 Roche, J., 456 Roder, A., 282 Ronnquist, O., 76, 82(89), 102(89), 424, 425(21) Roerig, D.L.,533(151), 534 Rogers, M.J., 465,483(121) Rogler, J. C.,527 Rokkanen, P.,154 Rochi, S., 31, 445, 446(59), 472,473(159)

605

AUTHOR INDEX

Roodyn, D., 537 Rorabacher, D., 31 Rosaria Bruzzesi, M., 447 Rosen, S. M., 531 Rosenbloom, J., 153, 159(148, 149) Rosenthal, O., 254, 255(8), 265, 266, 267, 268(59, 68, 69), 270, 274, 282(69), 283 (68) Rosenthal, S. M., 515, 516(53) Rosmus, J., 157 Rossmann, M. G., 67, 73, 77, 447 Roth, E. S., 180, 181(349, 352), 183(349) Rothberg, S., 120, 121, 128, 133(3), 134, 230 Rotilio, G., 125, 149(16), 357, 499, 500 (2021, 516, 517, 518(66), 519(66), 520 (661,521, 522, 523(89), 525, 526(106), 535, 536(178), 540, 542(241), 543, 545, 546, 547, 548, 549(241), 550, 551, 552, 553, 554(286, 2941, 555(277, 2941, 556 (255, 267, 292, 294), 557(294), 560, 566, 567 R.oulle, F., 556 Roulet, F., 513 Rowe, P. B., 388,410(276) Roea, M., 150 Rubin, B. H., 557 Rucker, R. B., 527 Ruf, H.-H., 186,286,287(161) Rummel, W., 270 Rundles, W. R., 401 Russell, G. A., 502 Ruzicka, F. J., 32 Ryan, D., 273 Ryan, J., 88, 94, 99(111, 131) Ryan, J. P., 312(65a, 661, 314 Ryan, K. J., 254 RydCn, L., 561, 562(340, 3421, 563 Ryhanen, L., 165, 166 Rynd, J. A., 500 Rytting, J. H., 362

S Sadava, D., 154, 163, 164 Saeki, Y ., 135 Saito, Y., 128 Saiton, Y., 230 Sakakibara, S., 153, 158(228), 159, 166, 294

Salach, J. I., 470 Salisbury, C. M., 543 Salmeen, I. T., 6, 10(66), 26 Salvador, R. A., 154 Saleman, L. A., 165 Samuel, D., 355 Samuelsson, B., 126 Sanadi, D. R., 325 Sanders, E., 265, 267, 268(59) Sands, R. H., lo(&), 20, 21, 22, 24, 26, 39, 40, 41(102), 42(102), 43(102), 195, 263, 341, 449 Sang, J. H., 410 Sanioto, D. L., 536 San Pietro, A., 29, 31, 61, 62, 79(44), 107, loS(44, 161a) Sasaki, N., 200 Sasame, H. A., 279 Satake, H., 271 Sato, H., 268 Sato, R., 255, 259(12), 270, 271, 274, 275, 276(12, 13, 124) Sauer, M. C. V., 184 Savage, J. E., 527 Sawada, S., 445 Sawada, Y., 538, 539(221), 540, 541(221), 542(221), 543(221), 544, 545(221, 2221, 546(221), 547(221), 553 Scandurra, R., 125, 148 Scarle, R. D., 336,337 Scazeocchio, C., 302, 406, 412, 413 Schaap, A. P., 536 Schacter, B. A., 273 Schadelin, J., 277, 278(130), 284(130) Schalet, A., 411 Schallies, A,, 538, 540(219), 542(219), 543 (219), 545(219), 546(219) Schandl, E. K., 174, 186(334) Scharf, R., 285 Scheinberg, I. H., 561, 565, 566, 571(376), 575 Schenkman, J. B., 270, 271, 274, 275(119), 276(119), 277 Schepartz, B., 146, 179 Scherbatyuk, L. I., 516, 518(60), 519(60), 520(60), 521(60), 522(60) Schimke, R. T., 128,172(32) Schimmel, P. R., 477 Schlaak, H.-E., 45 Schlegel, R. A., 531

606 Schleyer, H., 265, 266, 267, 268(68, 69), 282(69), 283(68) Schmidkunz, H., 358 Schnabel, K. H., 275, 276(126), 285 Schnactman, C. A., 230 Schnepf, E., 537 Schtillnhammer, G., 112, 480 Schoenheimer, R., 156 Schonbrunn, A., 199, 204(25, 26), 428, 451(31), 485(31), 488(31), 490(31), 498 Schoot Uiterkamp, A. J. M., 571 Schopfer, L., 213, 220 Schreffler, D. C., 559 SchumanJorns, M., 66, 105(63), 106, 108 (63, 160), 112, 113(181), 114(181), 115(181), 116(63, 1811, 192, 225(4), 244, 246(4), 399, 422, 424, 480(6), 482, 494(6), 498,501(6), 534 Schuta, G., 128, 129 Schwartz, B., 156 Schwara, H. A., 356, 357(206), 553, 556 (297) Schweigert, B. S., 145 Scola-Nagelschneider, G., 95 Seegmiller, J., 302 Seegmiller, J. E., 401 Segrest, J. P., 165 Seifried, H. E., 187 Seifter, S., 152, 165(139) Seiker, L. C., 367 Seng, R. L., 470 Senoh, S., 132, 133, 136, 141(62), 142, 143 (74), 151(93) Sesame, H., 274 Sevier, E., 173, 174(329), 175(329), 179 (329) Seybold, W. D., 314, 390(68), 407(68) Shaffer, P. M., 170, 171, 172, 173, 174 (3291, 175(328, 3291, 176(316, 322, 3281, 177(328), 178, 179 Shah, V. K., 50, 52(122) Shapiro, E. R., 566, 567(380) Sharaway, M. M., 162 Sharma, D. C., 255 Sharpless, K. B., 289 Sharrock, M., 264, 282(54) Sheppy, F., 239, 291 Shethna, Y. I., 59, 62(21), 64, 104(21) Shields, G. S., 533

AUTHOR INDEX

Shiga, K., 440, 447 Shiga, T., 440, 447 Shikita, M., 269 Shiman, R., 239, 240(143), 290, 291(179) Shimazono, N., 265 Shimomura, O., 188, 227 Shin, M., 15, 61, 62, 63, 79(44, 451, 108(44, 45) Shindler, J. S., 525 Shinkai, S., 496, 497, 500(191), 501, 502, 503( 193) Shinoda, T., 410, 411 Shiraishi, J., 445 Shiratori, T., 126 Shoeman, D. W., 271 Shokeir, M. H. K., 559 Shooter, K. V., 303, 305, 306, 312(40), 313(40, 43), 314(43) Shudo, K., 153, 166, 294 Siedow, J., 29, 31, 108 Sieker, L. C., 5, 32, 33, 37(93), 60, 66, 70(29, 641, 71, 74, 76(83), 78(64, 83), 79(64), 98(64), 114(64) Sih, C. J., 144, 151(101) Siiman, O., 8, 11(27b), 24, 35(27b), 37 (27b) Simmonds, H. A., 401,402 Simons, K., 155, 158(193), 292, 293 Simonyan, M. A., 551, 552(282), 554(282) Simpson, E. R., 19 Simpson, F. J., 125 Sinex, F. M., 166 Singer, T. P., 125, 149(18), 150(19), 301, 456, 470, 558, 559(309), 574(309) Sizer, I. W., 296, 456 Sjoerdsma, A.. 153, 241, 291, 302, 401 Slaughter, R. S., 172, 173(327), 175(328), 176(3281, 177(3281, 178(328) Sletten, K., 32 Smiley, I. E., 77 Smillie, R. M., 58, 59(4,5), 60, 61, 62(5), 63, 65(5), 82(5, 46), 91, 99(46, 1251, 100(125), 104(125), 537 Smith, A. J., 540 Smith, B. E., 50, 51(1191, 53, 54( 123), 55(131) Smith, J. R. L., 186 Smith, R. A., 515(55), 516, 522 Smith, R. S., 184 Smith, S. S., 59, 60(27), 66(27), 99(27)

AUTHOR INDEX

607

Smith, S. T., 350, 351(175), 360, 361, 389, Stein, G., 357 391(287), 393(175), 394(175), 395 Stein, H. O., 153 (2871, 396(175), 397(175), 398(175) Steinman, H. M., 537, 543(199), 557 Stensland, B., 76, 82(89), 102(89), 424, Smith, U., 58(17), 59, 62(17), 99(17) 425(21) Smith, W., 527, 531(119) Smith, W. W., 66, 70(66), 71(69), 72(66), Stenson, J. P., 368 73(66), 74(69), 75(69), 76(69), 77 Stern, R., 512 (69), 78(66), 79(69), 80(69), 81(69), Sternlicht, H., 44 82(69), 94(66), 98(66), 101(69), 102 Stetten, M. R., 156 Stevens, C. O., 145 (691, 114(66) Steward, F. C., 163 Snedden, W., 401,402(299) Stiefel, E. I., 334, 336, 368, 382, 418 Snell, E. E., 123, 127(7) Stirpe, F., 320, 321, 322, 366, 389(95), 402 Snyder, E. R., 522, 523(91) (95) Sobel, B. E., 4,6(9), 8, 14(9) Stoddard, D. D., 526 Sorbo, B., 125, 149 Sakoloff, R. L., 173, 174(329), 175(329), Stohrer, G., 352, 355(180, 182), 361 Stokes, A. M., 547 179(329) Stoltz, M., 166 Somerville, H. J., 63, 64(48) Stombaugh, N. A., 32, 34, 44, 47(108) Someya, J., 258 Stone, N., 156 Somogyi, A., 273 Song, P.-S., 78, 86(96), 92(127), 93, 94, Storm, C. B., 234, 235(123) 350, 351(178), 368(178), 424, 494(22) Strahs, G., 567 Strand, J. P., 270 Sonoda, Y., 258 Strecker, H. J., 499 Sote, H., 358 Strehler, B. L., 227 Sourkes, T. L., 467, 468 Strickland, S., 106, 108(160), 192, 213, 217, Soyama, T., 445 218(77), 220, 225(4), 246(4), 247, 534 Sparrow, L. G., 123, 127(7) Strittmatter, C. F., 397 Spector, S., 153, 154(174) Spector, T., 211, 212(62), 213, 214, 363 Strobel, H. W., 272, 273 Stromberg, C., 570, 574(392), 577, 578 Spencer, E., 574, 576(412) Sperling, O., 359 (392) Speyer, B. E., 569, 571(389) Strong, L. E., 560 Spiller, H., 58(15), 59, 62(15), 65(15), 82 Strong, L. H., 24 (151, 99(15) Strothkamp, K. G., 562 Spiro, R. G., 165 Struck, J., 456 Spiro, T. G., 23, 566, 567(380) Stubbs, C. S., 239 Sreeleela, N. S., 225 Stumpf, P. K., 456 Stabel, H. H., 45 Sturani, E., 108 Stangroom, J. E., 328, 341(122) Sturtevant, J. M., 352, 353(181), 369, 372, Stanier, R. Y., 128, 146, 211 373 Stansell, M. J., 533, 538, 539(216, 2171, Subba Rao, P. V., 225 541(216, 217), 543(217), 545(217) Suda, M., 146, 200 Stark, G. R., 559, 562, 563 Suga, K., 159 Stassen, F. L. H., 162 Staudinger, H. J., 186, 275, 282, 284, 286, Sugiura, N., 447,448 Suhara, I<., 255, 259(18), 260(18) 287 (162) Sullivan, P. A., 66, 105(63), 106, 108(63, Staudt, H., 279 1601, 116(63), 188, 192, 195, 196(23), Stauff, J., 358 Stavens, B. W., 24 197(23), 201(23), 203(23), 225(4), Steennis, P. J., 213 235,246(4,5), 248(5), 399,422,534

608 Sun, M.,78, 86(96), 92(127), 93, 424, 494 (22) Sund, H., 64, 90(51), 99(51), 102(51), 104 (51) Sundaram, T. K.,123, 127(7) Sutton, W. B., 194, 195, 198, 201(16) Suzuki, H., 513, 518(39), 519(39), 521(39), 523(39), 525(39), 526(39) Suzuki, I., 125 Suzuki, K.,19, 195, 206, 208, 210 Swallow, A. J., 31,90,328 Swann, J. C.,301, 302, 310, 311(58), 315(6, 57), 316(58), 323(58), 328(6), 329, 331, 332, 334(6), 337, 338(125), 339 (1251, 363(125), 369(6), 373(6), 374 (58, 1251, 375, 376, 381(125), 382 (125),387, 388(58) Sweat, M. L., 265 Sweeney, E.W.,128, 172(32) Sweeney, W. V.,45,46(114) Swindell, R., 523, 525, 526(95, 104) Swoboda, B. E.P., 427, 430(25), 434(25), 435(25), 436(25), 437(25), 438(25), 440(25), 441(25), 442(25), 449, 461, 462(25), 463(25), 465(251, 476(25), 477(25) Symons, M. C. R., 329 Szent-Gyorgyi, A., 558 Szybek, P.,514, 515, 519(50), 520, 521 (50),523(50), 525 Szymanska, J., 525

AUTHOR INDEX

Tamamoki, B. I.,268 Tamiya, N.,156, 158, 159(224), 292 Tamura, M.,130 Tanaka, M.,15, 17, 18, 66, 67(68, 'TO), 267) Tanaka, T., 129,182 Tang, S.-P. W., 23,566,567(380) Taniguchi, K.,180,181(348) Tanioka, H.,146 Taniuchi, H.,124(10), 125, 134, 135(63), 140 Tanphaichitr, V., 167 Tappel, A. L., 151, 154(133), 157, 158 (1331, 159(133), 161 Tappen, D.C.,178,179 Tarpley, W. B., 296, 558, 559(310), 574 (310) Tauscher, L., 102 Taylor, C. E.,517, 525(71), 526 Taylor, C.P.S., 24 Taylor, H.L.,560 Taylor, K.B.,295 Taylor, R. J., 239 Taylor, R. S., 517, 525(71), 526(71) Taylor, S. G., 77 Taylor, W.E.,19,268 Tedro, S.M.,32,35 Teitelbaum. H.D.,535 Teller, M. N.,352, 355(182), 361(182) Tel-Or, E.,26 Teng, N.,212 Terenzi, M.,550, 551(274) Teschke, R.,187 T Thayer, A. L.,536 Theorell, H., 83,150 Tabor, C. W., 515, 516 Thijsse, G. J. E., 258 Tabor, H.,515, 516 Thoai, N.V.,203,204,456 Tamvik, A.,537, 543(204) Thomas, G., 541(245), 542, 545(245), 546 Tagawa, K.,19,58,61 (245) Tagaya, N.,258 Thomas, K. A.,557 Tai, H.H., 144, 151(101) 163 Takahashi, M.,538, 539(223), 541(223), Thomas, M.D., 542(223), 544(223), 545(223), 547 Thomk-Beau, F., 204 Thompson, C. L.,8, 9(24, 27a), lO(24, (223), 556(223) 27a), 12(24), 35(27a) Takai, A., 444,448, 449(54), 450, 477(83) Takeda, H.,124(10), 125, 199, 200, 201 Thompson, J. F.,163 Thor, H.,270 Takeda, Y.,134,142,146 Takemori, S.,140, 146, 195, 206, 207, 208, Thorgeirsson, S., 279 Thorneley, R.N.F., 52 210, 255, 259(18), 260(lS) Takemoto, C., 268 Thornley, J. F.,22 Takeuchi, T., 153 Tiffany, M.L.,27

609

AUTHOR INDEX

Tillberg, O., 76, 82(89), 102(89), 424, 425 (21) Timpl, R., 166 Tipton, K. F., 468, 469, 471 TissiBres, A., 565 Tisue, T., 58(17), 59, 62(17), 99(17) Tofft, M., 167, 168, 169, 175, 176(336), 184(313), 293 Tokuyama, T., 132 Tollin, G., 59, 65, 66, 77, 82, 83, 84(61, 62, 100, 107, 108), 85, 86(61, 102, 1081, 87, 88, 90, 93(62), 94, 95, 99(61, 100, 131), lOO(23, loo), 102, 104, 107, 108 Tomida, I., 516 Tong, J. H., 240 Tong, L. K. J., 556 Torbjornsson, L., 76, 82(89), 102(89), 424, 425(21) Totter, J. R., 358,536 Trachevskii, V. V., 33, 336(138) Trautschold, I., 514, 519(48), 523(48) Travis, J., 32 Treadwell, C. R., 153 Treichel, P. M., 368 Trip, J. A. P., 561 Trittelvitz, E., 45 Trudgill, P. W., 259,263(29) Truscore, R., 511 Tsai, I., 154 Tsai, R. L., 22, 23(50), 259, 260(28), 261, 263 Tsibris, J. C. M., 4, 10(97b), 19, 22, 23(49, 501, 35, 263, 446 TSO,M.-Y., 50, 52(122), 55 Tsuji, T., 471 Tu, S. C., 445,446,447 Tul’chinskaya, L. S., 112 Turini, P., 125, 149(18) Tuttle, J. V., 389, 397(279a), 398(279a) Tyson, C. A., 260, 261, 264(31), 268(31)

Uitto, J., 153, 154 Ullrich, V., 186, 260, 261(33), 270, 274, 275, 276(123, 125, 1261, 277, 278(130), 279, 281(145), 282, 284, 285, 286, 287 (162) Ulmer, D. D., 8 Ultmann, J. E., 359 Ungar, H., 345, 346(165), 348(165), 349 (165) Uozumi, M., 317 Urano, M., 538, 539(223), 541(223), 542 (2231, 544(223), 545(223), 547(223), 556 (223) Urry, G. W., 502 Usprung, H., 315,389(68a) Uyeda, K., 62,65 (41) Uyeda, M., 130, 137, 138(49, 50, 771, 139 (50, 77)

V

Vanngbrd, T., 304, 317, 327,328(116), 329, 331, 332(116), 333, 334, 336, 337, 342 (251, 388, 509, 546, 547(254), 552, 562, 563, 564, 565(344, 347, 353, 361), 566 (11, 353, 361), 567, 569, 570, 571(287), 344, 358, 3641, 372(287, 344, 363, 364, 3931, 573, 574(364, 392, 3931, 575 (3851, 576, 577, 578, 579(407) Vaidyanathan, C. S., 124(9), 125, 225, 247(88), 534 Vainio, K., 154 Vaish, S. P., 102, 107 Valentine, R. C., 58, 59, GO, 61 (21, 62(24), 64(24, 25) Valerino, D. M., 345, 349 Vallee, B. L., 8 Vallogini, M. P., 521 van Arem, E. J. F., 104 Vance, P. G., 537 van Dam, J., 561 van den Hamer, C. J. A., 561 U van der Hoeven, T. A,, 272 Udenfriend, S., 151, 152, 153, 154, 155(144, Vanderlinde, R., 320 191), 156, 157, 158, 159, 160, 161, 162, van der Linden, A. C., 258 163, 165(187), 171(145), 174(145), VanEtten, R. L., 460, 483 184, 185, 186, 187(145), 188,219, 220, van Gelder, B. F., 534, 571, 572, 573 221(81), 232, 235, 238, 239(138), 240, Van Heuvelen, A., 56, 331, 332(128), 336 (128), 339 (128), 344 (128), 374(128), 254, 269(7), 284, 290, 291, 292, 293 375(128), 376(128), 377(128) Uehleke, H., 269

AUTHOR INDEX

van Leeuwen, F. X. R., 571, 573 van Lin, B., 59, 61(19), 62, 64(19), 65(19), 104119) Vanloon, E., 153(178), 154 Vanneste, M., 269 Van Slyke, D. D., 158, 166, 186(219) Varnon, J. E., 559 Vartanyan, L. S., 314 Veeger, C., 250, 424, 425(10), 448, 451, 454 Velcani, B. E., 64, llO(54) Veldink, G. A., 536 Veldsema, A., 571, 572, 573, 577(403) Venkatachalam, C. M., 72, SO(72) Vestling, C. S., 500 Vetter, H., Jr., 58(16), 59, 62(16), 64(16), 90(16), 91(16), 99(16), lOO(l6) Viscontini, M., 252 Vliegenthart, J. F. G., 536 Voelcker, G., 539, 543, 546(233, 248), 548 (2481, 549(248) Voelter, W., 311, 317, 328(60), 341(60), 538, 540(219), 541(245), 542, 543 (219),545(219,245), 5461219,245) Voet, D., 76, 447 Voet, J. G., 449, 454(87), 476, 481(165), 482(165), 483, 485, 488(185), 491 (185), 492, 494(165, 1871, 495(165, 1871, 496(165), 497(165), 501(165), 502(165, 182) Voetsch, W., 538, 540(219), 542(219), 543 (2191, 546(219) von Glehn, M., 76, 82(89), 102(89), 424, 425(21)

W Wada, H., 147 Wada, K., 15 Wade, T. D., 76 Wadeinski, I. M., 324 Wainer, A., 149 Waisman, H. A., 234 Walaas, O., 560, 576 Walker, G. A., 52 Walker, M., 55 Walker, W., 90 Walker, W. H., 248, 249,319, 328, 470, 501, 502(213) Wallace, W. J., 534

Walsh, C. T., 199, 204(24-26), 428, 431, 451, 485, 486, 488(31), 489, 490(30), 498 Wang, K. C., 144 Wang, P., 389, 397(279a), 398(279a) U'ang, T. M., 519 Warburg, O., 254, 461 Watanabe, K., 516,522 Watanabe, M. S., 170, 171(320), 175(320), 176(320), 187(320) Watari, H., 445, 476 Watenpaugh, K. D., 5, 60, 66, 70, 71, 72 (781,73(86), 74, 75, 76(83), 78, 79(64, 781, 98(64), 114(64) Waterman, M. R., 275, 282 Watson, J. G., 355, 367(190), 368(190) Watt, W. B., 388 Wattenberg, L. W., 270 Watts, D. C., 306, 310(41), 319, 352(41) Watts, R. W. E., 302, 401, 402 Waud, W. R., 309, 312, 321, 322,388(87a) Webb, E. C., 303 Weber, G., 89, 319, 320(79), 424 Weber, P., 275, 276(123), 277(123) Weber, U., 536, 557(190) Weibel, M. K., 427, 440, 465(27), 476(27), 483(27) Weiher, J. F., 6, 7(17), 9(17), 12(17), 25, 46(65) Weinhouse, S., 179 Weinstein, E., 166 Weir, E., 302 Weisbach, H., 165 Weisiger, R. A., 537, 538(200), 539(200), 540(200), 541(200), 542(200), 543 (200), 544, 545(200) Weissbach, H., 291 Wellner, D., 456: 457, 461(95), 480(95) Welton, A. F., 275 Wenk, M., 512 Werle, E., 513, 514, 519(48), 523(48) Werner, P.-E., 76, 82(89), 102(89), 424, 425 (21) Weser, U., 8, 9(27a), 10(27a), 35(27a), 510, 535, 536, 538, 539, 540(179, 219), 541(226, 245), 542, 543, 545(219, 226, 245), 546(219, 233, 245, 2481, 548, 549 (248), 557(190) West, C. A., 187, 280 West, S., 273

611

AUTHOR INDEX

Westerfeld, W. W., 303, 320, 410 Westhead, E. W., 577, 578(431) Westheimer, F. H., 463 Westlake, D. W. S., 559 Westley, J., 469, 470(145) Wever, R., 534, 571, 573 Wexler, S., 536 Whatley, F., 22 Whelan, D. T., 182 White, A. H., 53 Whiteley, H. R., 8, 12(26), 389 White-Stevens, R. H., 206, 207, 208, 213 (521, 225(53), 247 Whitfield, C. D., 110, 111, 112, 113, 114 (MI), 115(170, 178, 1811, 116(170, 178, 1811, 117(170, 178), 118(170), 480 Whitlock, H., Jr., 144 Whyman, R., 547 Wieher, J. F., 341 Wiley, R. D., 312 Wilkie, D., 537 Wilkinson, G., 339 Williams, C. H., Jr., 31, 273, 445, 446(59), 449,472,473(159) Williams, J. H., 410 Williams, L. G., 170, 173, 175(323), 178 Williams, R. F., 497, 502(193), 503 Williams, R. J. P., 282, 569 Williams, R. T., 269 Williams, V., 511 Williams, W. M., 12 Willson, R. L., 556 Wilson, D. F., 24 Wilson, G. S., 19, 263 Wilson, L. D., 255, 265 Wilson, P. W., 59, 62(21), 104(21) Wintrobe, M. M., 533, 538, 541(215), 548 (215) Witkop, B., 158, 160, 182, 219, 220, 221 (81), 235, 284 Wolf, J., 284 Wolkow, M., 179 Wong, S. H., 265 Wood, E. J., 539, 540(231), 541(231, 239), 543(231), 545(231). 546(239, 2501, 548(231), 549(231, 239, 250) Wood, H. C. S., 250, 424, 425(10) Wood, J. M., 143 Wood, N. L., 153(177), 154

Wood, P. M., 356, 400(204), 532, 556, 579 (140) Wood, W. A., 110 Woody, R. W., 4 Woolfolk, B. S., 389 Woolfolk, C. A., 389 Wyckoff, H. W., 77 Wyngarden, J. B., 388, 410(276)

X Xuong, Nguyen-huu, 32, 33(90), 35(90)

Y Yagi, K., 432, 434(44), 444, 445, 446, 447, 448, 449, 450, 452, 454(44), 455(44), 460(44), 476, 477, 479, 480(44), 482 (441, 483(44) Yamada, H., 513, 514(43), 515, 516, 517 (611, 518, 519, 520(79), 521, 522, 523, 525(39, 821, 526(39) Yamamoto, S., 127, 131,200, 201, 202,206, 207(49), 226, 230 Yamamoto, T., 6, 511 Yamano, H., 519, 520(83), 521(83) Yamano, M., 445 Yamano, T., 134, 136, 140, 143(74), 271, 272, 484, 505, 519, 520(79), 521(79) Yamasaki, E. F., 523, 525(95), 526(95) Yamasaki, I., 538, 539(221), 540(221), 541(221), 542(221), 544(221), 545(221, 222), 546(221), 547(221), 553, 505 Yamauchi, T., 201,202 Yamazaki, I., 130, 322, 354, 355, 356, 367 (186), 574, 576 Yano, K., 211, 212(63), 214 Yashimura, J., 448, 471, 482(82) Yasuda, H., 206, 207(50), 208 Yasunobu, K., 146, 182 Yasunobu, K. T., 8, 12(26), 13, 15, 17, 18, 66, 67(68, 70), 267, 467, 468, 469, 470(131, 144, 1451, 515, 516, 518, 519, 520(83), 521, 522, 523(77, 78, 82), 525, 526 (103) Yates, M. G., 52, 61, 62(36), 65(36), 109 Yen, T. T., 407, 408,410(309) Yoch, D. C., 59, 60, 62(24), 64(24, 25), 99(144), 100 Yoegtli, W., 524, 525(100)

612

AUTHOR INDEX

Yohro, T., 265 Yokoe, I., 499, 501, 502 Yokum, C. F., 29 Yonetani, T., 21 Yoo, B. Y., 470, 471(153), 498(153) Yoshida, H., 558 Yoshimura, T., 461 Yost, F. J., Jr., 534, 537 Youdin, M. B. H., 467 Young, S. N., 559, 669, 575, 576(422) Yousten, A. A., 535 Yu, C. A., 255, 259(18), 260, 261,262, 280 (371, 283(39)

YU, T. F., 302, 363(12), 401(12)

Z Zabel, A., 514 Zabinski, R., 143 Zach, D., 123, 127(7) Zahid, N. D., 280

Zahrradnik, R., 157 Zaltsman-Nirenberg, P., 219 Zamaraev, C. I., 516, 518(60), 619(60), 520(60), 521(60), 522(60)

Zancan, G. T., 531 Zanetti, G., 31, 472,473(159) Zannoni, V. G., 179, 180, 181, 182, 238 Zapesochnaya, L. G., 112 Zeller, E. A., 456, 470, 471, 498(154), 512, 513,516,521,523,524,525,527(86)

Zeszotek, E., 470 Zhuravleva, 0. S., 556 Ziegler, D. M., 229 Zimmerman, R., 535 Zito, R., 296, 297(230) Zubieta, J. A,, 32, 34 Zucker, M., 296 Zumft, W. G., 50, 52, 53(121, 126), 54 (134), 55, 58(15), 59, 62(15), 85(15), 82(15), 99(15)

Subject Index A

Acetylene, reduction of, 50,51, 53 Acetylenic compounds, flavoprotein oxiAbsorption spectra dases and, 498 amine oxidases, 518-519 N-Acetyl-o-galactosamine, galactose oxicamphor 5-ezo-monooxygenase, 260dase and, 531 261 N-Acetylimidazole, spinach ferredoxin cytochrome P-420, 270-271 and, 29 cytochrome P450,273-277,282-283 N7-Acetyl-5-methoxykynurenamine, foreight iron-sulfur protein, 45 mation of, 132 electron-transferring flavoproteins, 112, 3-Acetylpyridine adenine dinucleotide, 114 orcinol hydroxylase and, 224 flavodoxins, 88-91,99 Active site, amine oxidases, substrate insingle crystals, 91-93 teraction, 524-525 p-hydroxybenzoate hydroxylase, 212 Activity-flavin ratio, xanthine oxidase melilotate hydroxylase, 217, 218 and, 310 molybdenum hydroxylases, 393-394 Acylation, rubredoxin, 12 phenol hydroxylase, 223 Acyl coenzyme A dehydrogenase, flavoprotocatechuate 3,4-dioxygenase, protein and, 57, 109 137-139 Adenine pyrocatechase, 135 glucose oxidase and, 462 rubredoxin, 7-8 molybdenum hydroxylases and, 347 salicylate hydroxylase, 207 Adenosine, indoleamine 2,3-dioxygenase llg-steroid monooxygenase, 265266 and, 131 tryptophan 2,3-dioxygenase, 129, 130 Adenosine diphosphate, nitrogenase and, two iron-sulfur proteins, 19,23, 24-25 50, 52, 54 apoproteins, 26 Adenosine triphosphate, nitrogenase and, xanthine oxidase, 318, 328 50, 51, 52, 54, 55 Acetaldehyde Adenylate, flavin adenine dinucleotide molybdenum hydroxylases and, 346, binding and, 446 352 Adenylyl sulfate reductase, properties of, nitroethane oxidation and, 496 48 Acetate, galactose oxidase, 53 Adrenal mitochondria, Acetone cholesterol side chain cleaving monocytochrome P-450,.,, and, 262 oxygenase of, 268-269 3,4-dihydroxyphenylacetate2,3dioxy1Ip-steroid monooxygenase of, 265-268 genase and, 142 two iron-sulfur protein, ferredoxin and, 23 properties, 16 metapyrocatechase and, 140 amino acid sequence, 18 steroid oxygenase and, 144 613

614

SUBJECT INDEX

antibodies and, 26 Adrenochrome, formation of, 357 Adrenocorticotropic hormone, cholesterol utilization and, 266 Adrenodoxin, 2 11p-steroid monooxygenase and, 268, 283

Affinity chromatography, xanthine oxidase, 307, 310 Agkistrodon piseivorus, Gamin0 acid oxidase of, 456 Agmatine, amine oxidases and, 513, 514, 516, 517

Alanine n-amino acid oxidase and, 444,450,451, 452,453,455,480

lysine monooxygenase and, 202 spinach ferredoxin and, 28 Alanine residues, flavodoxins, 69 Aldehyde (s) amine oxidases and, 511-512, 526 bacterial luciferase and, 227,228 monoamine oxidase and, 469 Aldehyde oxidase deflavo form, 319 Drosophila, substrates, 407-408 A”-methylnicotinamide and, 362 molecular properties, 390-391 properties of, 48 substrate specificity, 345-352 Aldehyde oxidase mutant, chromosome locus, 407 Alfalfa, ferredoxin, amino acid sequence, 17

Alkaloids, biosynthesis of, 280 Alkanes, w-hydroxylation, rubredoxin and, 12 n-Alkylamines, amine oxidase and, 514 Allopurinol Aspergillus xanthine dehydrogenases and, 413 molybdeum hydroxylases and, 348 xanthine oxidase and, 302,354, 363-364, 365,375,383

affinity chromatography, 307 xanthinuria and, 401 Alloxanthine, xanthine oxidase and, 315, 354, 363, 387

Amine(s), amine oxidase spectra and, 518, 520-521

Amine oxidase (s), 509 active site, substrate interaction, 524-525

catalytic mechanism, 525-526 copper content, 519-521 definition and classification, 511-513 microsomal, properties of, 229-230 purification, molecular weight and substrate specificity, 513-518 Amino acid(s) adrenal iron-sulfur protein sequence, 267

amine oxidase composition, 515 D-amino acid oxidase composition, 446 blue oxidase composition, 562-563 camphor 5-eso-monooxygenase composition, 260 eight iron-sulfur protein sequences and, 38-39 ferredoxin sequences, adrenodoxin, 18 other plants, 17 putidaredoxin, 18 spinach, 15,17 flavodoxin composition, 59-60 sequences, 67-69 four iron-sulfur protein composition, 34

galactose oxidase composition, 528, 529 milk xanthine oxidase content, 310-312 rubredoxin sequences, Clostridium pasteurianum, 5 Pseudomonas oleovorans, 13 superoxide dismutase composition, 543-545

Amino acid oxidase(s) long wavelength intermediate, 479-482 reductive half-reaction, 432-434, 449-451, 476

n-Amino acid oxidase enzyme-monitored turnover, 440 flavin dissociation constants, 500 kinetic mechanism, 436437,444 lactate oxidase and, 198-199 model substrates and, 484-492 molecular properties and kinetic mechanism, 445-456 monitoring of, 428 nitroethane anion and, 476, 481

SUBJECT INDEX

steady-'state behavior, 430 L-Amino acid oxidase computer simulation, 442-443 enzyme-monitored turnover, 439-440 kinetic mechanism, 437, 444-445 molecular properties and kinetic mechanism, 456461 steady-state behavior, 430-431 o-Aminobenroate, n-amino acid oxidase and, 484 p-Aminobenroate, p-hydroxybenroate hydroxylase and, 213, 216 c-Aminocaproate, lysine monooxygenase and, 200,201 a-Amino-p-carboxymuconic semialdehyde, formation of, 145 a-Amino-p-chlorobutyrate, n-amino acid oxidase and, 486, 489 2-Amino-2-deoxy-~-galactose, galactose oxidase and, 531 Aminoguanidine, amine oxidases and, 523 2-Amino-4-hydroxypteridine, Drosophila xanthine dehydrogenase and, 407 2-Amino-4-hydroxypteridine-&aldehyde, xanthine oxidase and, 363 4-Amino-7-hydroxypteridine, molybdenum hydroxylase and, 349 Aminophenol(s), phenol hydroxylase and, 222 p-Aminosalicylate, salicylate hydroxylase and, 207,210 bAminovaleramide, formation of, 199-200 Ammeline, molybdenum hydroxylases and, 362 Ammonia amine oxidases and, 511-512, 526 D-aminO acid oxidase and, 480 flavoprotein oxidases and, 495 monoamine oxidase and, 469 thymine 7-hydroxylase and, 179 Amylamine, amine oxidase and, 516 Anacystis nidulans flavodoxin, 58, 60-61 absorption maxima, 99 activity of, 62 dithionite and, 104 redox potentials, 99, 100 1,5-Anhydrogalactitol, galactose oxidase and, 531

615 Aniline, eytochrome P-450 and, 277 Anions, sulfite oxidase and, 417, 418 Anthranilate D-amino acid oxidase and, 441, 446, 452-453,477,478,487-488,489 formation of, 147 Anthranilate hydroxylase(s), reactions and cofactors, 124, 125 Antibodies cytochrome P-450 and, 285 electron-transferring flavoproteins and, 113 ferredoxin apoproteins and, 26 flavodoxins and, 63,64 prolyl hydroxylase and, 162, 163, 292-293 rubredoxins and, 13-14 Aphanothece sacrum, ferredoxin of, 15 Apoflavodoxins preparation of, 65 and properties, 82-83 Apomyoglobin, cytochrome P-450,,, and, 262 Apoproteins two iron-sulfur proteins, reconstitution, 25-28 Arene oxide, aromatic hydroxylations and, 219-221, 235,237,252 Arginine D-amino acid oxidase and, 444 lysine monooxygenase and, 200,201, 202 Arginine decarboxylase, see Arginine monooxygenase Arginine monooxygenase, properties, 203-204 Arginine residues eight iron-sulfur proteins, 38-39 flavodoxin, 79 Arsenite sulfite oxidase and, 418 xanthine oxidase and, 325-326 Arthrobacter, melilotate hydroxylase of, 217 Arthrobacter pascens, uricase of, 511 Ascaris lumbricoides, prolyl hydroxylase of, 154, 164 Ascorbate ybutyrobetaine hydroxylase and, 168 copper protein-linked, monooxygenases

616 and, 122 ceruloplasmin and, 574-575 dopamine p-monooxygenase and, 294 5-formyluracil dioxygenase and, 175 homogentisate oxygenase and, 146 5-hydroxymethyluracil dioxygenase and, 175 p-hydroxyphenylpyruvate hydroxylase and, 180, 181 indoleamine 2,3-dioxygenase and, 131 a-ketoglutarate dioxygenases and, 185-186 lysyl hydroxylase and, 166,294 prolyl hydroxylase and, 157, 158, 162-163, 164, 185, 293 pyrimidine deoxyribonucleoside 2’hydroxylase and, 177 thymine 7-hydroxylase and, 174 tryptophan 2,3-dioxygenase and, 129 Ascorbate oxidase, 509 copper of, 564465, 570-571 discovery of, 558 function of, 559 molecular properties, 561 purification of, 560 specificity, 574 Asparagine residues, flavodoxin, 76, 78 Aspartate, histidine degradation and, 226 Aspartate residues flavodoxins, 69,79 oxidation, state and, N,81 Aspergillus flavus, quercitine dioxygenase of, 125 Aspergillus nidulans molybdenum hydroxylases of, 412414 nitrate reductase, molybdenum hydroxylase “common cofactor” and, 402-406 Aspergillus niger amine oxidase, 512 catalytic mechanism, 526 copper content, 519,520 inhibitors, 523 purification, molecular weight and substrate specificity, 513-514 pyridoxal phosphate and, 521 spectral properties, 518-519 glucose oxidase of, 461,462, 463 m-hydroxybeneoate-4-hydroxylase of, 225

SUBJECT INDEX

Aspirin, prostaglandin synthetaae and, 127 6-Azathymine deoxyribonucleoside, 2’hydroxylase and, 178 Aeide amine oxidases and, 523, 527 blue oxidases and, 569,575 galactose oxidase and, 530,531 superoxide dismutase and, 547-548,552, 556 Azotobacter, nitrogenase of, 52 Azotobacter chroococcum flavodoxin, 62 nitrogenase and, 109 Azotobacter vinelandii flavodoxin, 59, 82 absorption spectra, 89, 90,91, 99 activity of, 62-63 circular dichroism, 95 disproportionation, 102 dithionite and, 104 electron nuclear double resonance, 96 flavin binding and, 83-85,88 flavin derivatives and, 84,85-87 fluorescence, 93-94 iron and, 64 oxygen and, 107 reduction of, 82 Azotoflavin, 59 Azurin, copper of, 566,567

B Bacillus polymyxa four iron-sulfur protein, 32, 33 properties of, 34 Bathocuproine, tryptophan 2,3-dioxygenase and, 130 Benzene oxygenase, reaction and cofactors, 124, 125 Benzoate D-amino acid oxidase and, 440-441, 445, 451,477,478,488 p-hydroxybenzoate hydroxylase and, 213 salicylate hydroxylase and, 207,209, 210 Benzoate oxygenase, reaction and cofactors, 124, 125

617

SUBJECT INDEX

Benzoin, oxidation, flavin and, 501 Benzoquinone, xanthine oxidase and, 322 4-Benzoylamido4’-aminostilbene-2,2’disulfonate, D-amino acid oxidase and, 447 Benzphetamine, demethylation of, 273 Benzpyrine, cytochrome P-450 and, 275, 277 Benzylamine amine oxidases and, 468, 514, 516, 518, 527 substituted, monoamine oxidase and, 470 Benzylamine oxidase, 466 discovery of, 517 Benzyl cyanide, monoamine oxidase and, 469 Benzyl mercaptan, iron-sulfur protein model compound and, 46 2,Y-Bipyridyl, see Dipyridyl Bis(aminoethy1) benzene, amine oxidase and, 517 N,N’-Bis(3-aminopropyl)-1,2-diaminoethane, amine oxidase and, 516 N,N‘-Bis(3-aminopropyl~-l,6diaminohexane, amine oxidase and, 516 Bis [u-xylyldi thiolato-prsulf idoferrate (III)], ferredoxins and, 25 Blood, ceruloplasmin preparation from, 560-561 Borohydride amine oxidase and, 521, 522 D-aminO acid oxidase and, 455, 456 L-amino acid oxidase and, 461 flavoprotein oxidases and, 495 xanthine oxidase and, 352-353,368 Bovine liver, uricase of, 511 Bovine plasma amine oxidase, 512 catalytic mechanism, 526 copper content, 519,520 purification, molecular weight and substrate specificity, 514-516 pyridoxal phosphate in, 522 spectral properties, 518-519 Bovine serum albumin galactose oxidase and, 532 lysyl hydroxylase and, 166 prolyl hydroxylase and, 158 Bradykinin, prolyl hydroxylase and, 159

Brevibacterium juscum, pyrocatechase of, 134-135 Bromide galactose oxidase and, 530 glucose oxidase and, 465 3-Bromoallylamine, monoamine oxidase and, 498 N-Bromosuccinamide flavodoxins and, 88 rubredoxin and, 12 5-Bromouracil deoxyribonucleoside, 2’hydroxylase and, 178 Butanol, phenylalanine hydroxylase and, 235 Buttermilk, xanthine oxidase isolation from, 305 Butylamine, amine oxidases and, 516,517 t-Butyl hydroperoxide, a-ketoglutarate and, 184 Butylmercaptane, cytochrome P-450 and, 28 1 Butyrate, formation, flavodoxin and, 61 y-Butyrobetaine hydroxylase catalytic properties, 168-169 mechanism, 183-184 purification, 167 Butyryl coenzyme A dehydrogenase, electron-transferring flavoprotein and, 110, 113, 116-117

C Csdaverine, amine oxidases and, 514, 517 Calcium chloride, xanthine oxidase and, 318 Camphor, cytochrome P-450,,, spectrum and, 261 Camphor Bezo-monooxygenase, properties, 259-264 Canavanine, arginine monooxygenase and, 203 Cancer, xanthine oxidase and, 302 Candida tropicalis, monooxygenase of, 280 Candida utilis DPNH-ubiquinone reductase, 49 uricase of, 511 Carbohydrates amine oxidases and, 516, 518 blue oxidases and, 561

618 Carbon dioxide 5-formyluracil oxidation and, 176 p-hydroxyphenylpyruvate hydroxylase and, 183 internal monooxygenases and, 194 lysyl hydroxylase and, 166 prolyl hydroxylsse and, 161, 185 pyrimidine deoxyribonucleoside 2'hydroxylase and, 178 salicylate hydroxylase and, 206 thymine 7-hydroxylase and, 175 Carbonic anhydrase cobalt enzyme, 550 xanthine oxidase and, 366 Carbon monoxide adrenal cortex microsomes, 265 mitochondria, 265 cholesterol cleaving cytochrome P-450 and, 269 cytochrome P-450 and, 275,276,277, 278,282,283,284 cytochrome P-450,., and, 260,261,262 dopamine 8-monooxygenase and, 295 microsomal hydroxylases and, 254, 258 xenobiotic monooxygenase and, 270, 271 @-Carboxymuconicacid, formation of, 136 Carboxypeptidase (9) eight iron-sulfur protein and, 45 spinach ferredoxin and, 28 Carnitine, formation of, 167 &Carotene, dioxygenase and, 123, 126 Carrot, prolyl hydroxylase of, 163-164 Catalase r-butyrobetaine hydroxylase and, 168 flavoprotein oxidases and, 427 galactose oxidase and, 530 p-hydroxyphenylpyruvate hydroxylase and, 181 lysyl hydroxylase and, 166,294 orcinol hydroxylase and, 223-224 prolyl hydroxylase and, 158 pyrimidine deoxyribonucleoside 2'hydroxylase and, 177 superoxide dismutase and, 535, 556 thymine 7-hydroxylase and, 174 tryptophan pyrrolases and, 131, 292 tyrosine hydroxylase and, 239, 291

SUBJECT INDEX

Catechol metapyrocatechase and, 141, 142 phenol hydroxylase and, 221-222 protocatechuate 3,4-dioxygenase and, 137 Catecholamine, monoamine oxidase and, 466 Catran, amine oxidase and, 522 Cerebrocuprein, 533 Ceruloplasmin, 509 copper of, 563, 565, 570, 571 discovery of, 558 function of, 559 molecular properties, 561 oxidation-reduction potentials, 572-573 purification of, 560 specificity, 574-575 Charge-transfer complex, D-amino acid oxidase and, 482 Cheese, monoamine oxidase and, 466 Chelex galactose oxidase and, 529 nitrogenase and, 52 Chemiluminescence, superoxide dismutase and, 536 Chicken liver superoxide dismutase of, 539, 541, 544 xanthine dehydrogenase, catalytic properties, 396, 398, 399 molecular properties, 390-391 Chlorella fusca flavodoxin, 59 absorbance spectra, 91, 99 activity of, 62 redox potentials, 99 iron deficiency in, 65 Chloride galactose oxidase and, 531 glucose oxidase and, 465 p-hydroxybenzoate hydroxylase and, 213 phenol hydroxylase and, 222-223 8-Chloroalanine amino acid oxidase and, 199 n-amino acid oxidase and, 431, 484-491 B-Chloro-a-amino acid(s), n-amino acid oxidase and, 446, 484-492 4-Chlorocatechol metapyrocatechase and, 141 pyrocatechases and, 134

SUBJECT INDEX

Chloroform-ethanol, superoxide dismutase and, 537, 538 p-Chlorolactate, lactate oxidase and, 199 p-Chloromercuribenroate, superoxide dismutase and, 543 Chlorophenol(s), phenol hydroxylase and, 222 4-Chlorophenylalanine, phenylalanine hydroxylase and, 234 Chloropyruvate, n-amino acid oxidase and, 485 Cholesterol side chain cleaving monooxygenase, properties, 268-269 Chromatium high potential iron-sulfur protein, 32 Mossbauer parameters, 10 nitrogenase of, 53 Chromium laccase and, 578 rubredoxin and, 14 Chromohacterium violaceum, tryptophan 5-monooxygenase in, 291 Chymotrypsin flavodoxins and, 67 phenylalanine hydroxylase and, 236, 289 tyrosine hydroxylase and, 239 xanthine oxdase and, 321 Circular dichroism apoferredoxin, 26, 27 apoflavodoxins, 82, 83, 85 blue oxidases, 566 camphor 5-ezo-monooxygenase, 264 flavodoxins, 94-95 galactose oxidase, 530-531, 532 p-hydroxybenzoate hydroxylase and, 212 metapyrocatechase, 141-142 4-methoxybenroate O-demethyl-monooxygenase, 286, 287 pyrocatechase, 136 superoxide dismutase, 545-546, 550 xanthine oxidase, 328, 341 Claviceps purpurea, cytochrome PA50 of, 280 Clorgyline, monoamine oxidase and, 469 Clostridium, flavodoxins of, 60 Clostridium acidi-urki ferredoxin of, 38-39, 42, 44 pyruvate dehydrogenase of, 48

619 Clostridium cylindrosporum xanthine dehydrogenase molecular properties, 390-391, 393 oxidizing substrates, 398 reversibility, 360 substrate specificity, 345-352 Clostridium M P flavodoxin, 83 absorption spectra, 89, 99 amino acid sequence, 67-69 circular dichroism, 94, 95 distribution of residues, 79 dithionite and, 103 flavin binding site, 74-78, 87-88, 109 flavin derivatives and, 86-87 fluorescence, 93, 94 iron and, 64 molecular conformation, 71-73 nuclear magnetic resonance, 96-97 oxidation state and structure, 79-82 oxygen and, 107 redox potentials, 99, 100, 101, 102 single crystal absorption, 91-93 structure determination, 70-71 three-dimensional structure, 67 Clostridium pasteurianum eight iron protein of, 34 absorption spectrum, 45 Mossbauer spectra, 37 nuclear magnetic resonance, 44 flavodoxin, 58, 61, 65 absorption spectra, 88, 89 activity of, 62 dithionite and, 104 nuclear magnetic resonance, 96-97 partial sequence of, 66, 68, 69 redox potentials, 99, 100, 101 regulation by iron, 63-64 iron protein, Mossbauer parameters, 11 nitrogenase of, 48, 52, 53, 56 rubredoxin, 4 amino acid sequence, 5 Mossbauer parameters, 10 three-dimensional structure, 5-6 Clostridium tartarivorum, ferredoxin of, 46 Clostridium thermosaccharolyticum, ferredoxin of, 46 Cobalt, superoxide dismutase and, 549, 550-551

620 Coenzyme Q p-hydroxyphenylpyruvate hydroxylase and, 180 xanthine oxidase and, 367 Collagen amine oxidase and, 527 peptides, extent of hydroxylation, 159-160 thermal stability, 152-153 Complementation, maroon-like and rosy mutant extracts, 411 Comproportionation, flavodoxins, 102103 Computer, flavoprotein oxidases m d , 442-443 Conformation, flavodoxin, oxidation state and, 79-82 Connective tissue, amine oxidase of, 527 Copper amine oxidase, 527 content, 519-521 spectra and, 518,519 blue oxidases, 561-562 magnetic and spectroscopic properties, 563-571 type 1, 566567,568 type 2, 567, 569-570 type 3, 570-571 dioxygenases and, 125, 129-130 dopamine p-monooxygenase and, 295 galactose oxidase and, 529-530,532 monoamine oxidases and, 466,468 phenol o-monooxygenase and, 297 superoxide dismutase and, 533, 537, 545,546-548, 549-550, 553-556 redox properties, 551-552 Copper chelates, superoxide and, 536,557 Copper-containing oxidase (s) blue biological distribution and function, 558-559 catalytic properties, 574-579 forms of copper, 563-571 history, 557658 oxidation-reduction properties, 571474 oxygen and, 508 purification and molecular properties, 560-563 catalytic properties

SUBJECT INDEX

reducing substrates, 575-578 reduction of oxygen, 578-579 specificity, inhibition and steadystate kinetics, 574575 historical background, 507-511 Copper protein(s) ascorbate-linked, monooxygenases and, 122 dioxygenases and, 122 Corynebacter-ium, cytochrome P-450 monooxygenase of, 280 Cream, xanthine oxidase isolation from, 305 Creatine-creatine kinase, nitrogenase and, 54 m-Cresol, orcinol hydroxylase and, 223-224 Crotalus adamanteus, L-amino acid oxidase of, 456 Crotonyl coenzyme A, electron-transferring flavoprotein and, 116 Cucumis sativus, ascorbate oxidase of, 560 Cuprizone, amine oxidase and, 522,523, 524 Cyanate, superoxide dismutase and, 548 Cyanide aldehyde oxidases and, 393 amine oxidases and, 523, 527 L-amino acid oxidase and, 461 blue oxidases and, 569,575 dopamine p-monooxygenase and, 295 flavoprotein oxidases and, 427,492-493 galactose oxidase and, 529,530,531 laccase and, 565, 570 phenol o-monooxygenase and, 297 sulfite oxidase and, 418 superoxide dismutase and, 537,546-547, 548,551,552, 556 xanthine oxidase and, 307, 323,324,325, 326, 352,365, 383 5-Cyanoethyl-1,5-dihydroflavin adenine dinucleotide, formation of, 493-494, 496 Cyanogen bromide, flavodoxins and, 67 4-Cyanophenol, monoamine oxidase and, 469 6-Cyanopurine, molybdenum hydroxylases and, 347 Cyclohexane, cytochrome P-450 and, 275,

621

SUBJECT INDEX

278, 279 Cysteamine amine oxidase and, 518 dioxygenase and, 125 Cysteamine oxygenase, properties, 148149 Cysteine dioxygenase and, 125 homogentisate assay and, 183 metapyrocatechase and, 140 nonheme iron protein and, 289 protocatechuate 4,5-dioxygenase and, 143 Cysteine oxygenase, properties, 149-150 Cysteine residues adrenal iron-sulfur protein, 267 D-amino acid oxidase, 446 blue oxidases, 566-567 eight iron-sulfur proteins, 37-38, 43, 45 flavodoxins, 60,70 distribution of, 78-79 modifitation of, 78, 87-88 four iron-sulfur proteins and, 32 monoamine oxidase, 470 putidaredoxin, 20, 263 rubredoxin, 6, 12 spinach ferredoxin, 15, 22 superoxide dismutase, 546 Cysteine sulfinic acid, formation of, 125, 149 Cysteine trisulfide, ferredoxin and, 26 Cytochrome(s), iron deficiency and, 65 Cytochrome b,, cytochrome P-450 and, 270, 272, 279 Cytochrome c cytochrome P-450 and, 272-273 EPR spectrum. 24 4-methoxybenzoate O-demethylmonooxygenase and, 286 molybdenum hydroxylases and, 397 reduction, flavodoxin and, 63, 108 11p-steroid monooxygenase and, 268 sulfite oxidase and, 417, 418 xanthine oxidase and, 354,357,359,366 Cytochrome c reductase cytochrome P-450 and, 273 nitrate reductase and, 403, 405 Cytochrome oxidase, ceruloplasmin and, 559

copper and, 507-508 Cytochrome P-420 formation of, 270 spectral characteristics, 271 Cytochrome P-450 absorption spectra, 273-277, 282-283 cholesterol side chain cleaving monooxygenase and, 268.269 hydroxylases and, 255, 258-259 inhibitors of, 284-285 oxygen activation, 283-284 llp-steroid-monooxygenase and, 265, 268 structure, 280-282 substrate binding by, 282-283 xenobiotic monooxygenases,and, 270 Cytoeuprein, 533

D Dactylium dendroides, 527 5-Deaaaflavin, flavoprotein oxidases and, 497 Dehydrogenase(s1, flavoprotein, flavodoxins and, 66, 73 Deoxycholate, cytochrome P-450 and, 270, 271-272 Deoxycorticosterone, steroid 118-monooxygenase and, 266, 267 2-Deoxy-~-galactose, galactose oxidase and, 531 2-DeoxygIucose, glucose oxidase and, 436, 444, 462, 463, 465, 466 2’-Deoxyriboflavin adenine dinucleotide, D-amino acid oxidase and, 447 Deoxyribose, salvage of, 171-172 Deoxyuridine conversion to uridine, 176, 178 2’-hydroxylase and, 173 Desuljovibrio flavodoxins of, 60 sulfate reduction in, 61 Desulfovibrio desulfuricans, high potential iron protein, 32 Desulfovibrio gigas adenylyl sulfate reductase of, 48 flavodoxin, properties, 99 high potential iron protein, 32 hydrogenase of, 48 Desulfovibrio vulgaris flavodoxin, 66

622 absorption spectra, 90, 99 amino acid sequence, 67-69 circular dichroism, 94-95 distribution of residues, 79 flavin binding site, 74-78,88, 109 flavin derivatives and, 85-87 fluorescence, 93 molecular conformation, 71-73 reduction potential, 99, 101 structure determination, 70 three-dimensional structure, 66,81 Detergents cytochrome P-450 and, 270-273 cytochrome P-450,., and, 262 Deuterium oxide, flavin adenine dinucleotide and, 328 Diamine oxidase(s), characteristics of, 512 2,4-Diamino-6-hydroxy-s-triazine,molybdenum hydroxylases and, 362 2,8-Diaminooctanoate, lysine monooxygenase and, 201 3,5-DichlorocatechoI, metapyrocatechase and, 141 2,6-Dichlorophenolindophenol electron-transferring flavoprotein and, 110, 113, 117 p-hydroxyphenylpyruvate hydroxylase and, 180 4-methoxybenzoate O-demethyl-monooxygenase and, 286 molybdenum hydroxylases and, 397, 398 sulfite oxidase and, 417 xanthine oxidase and, 322,353-354 Diethyldithiocarbamate amine oxidases and, 519,522,523-524, 527 ceruloplasmin and, 565 galactose oxidase and, 529 tryptophan 2,3dioxygenase and, 130 Diethylpyrocarbonate, arginine monooxygenase and, 204 Digitonin A, monoamine oxidase and, 467 Dihydrobiopterin, quinonoid, reduction of, 232-233, 238 7,8-l)ihydrobiopterin, reduction of, 233 l,&Dihydroflavin, formaldehyde reduction and. 502

SUBJECT INDEX

Dihydrofolate reductase, 7,gdihydrobiopterin and, 233-234 Dihydroorotate dehydrogenase, properties of, 48 Dihydropteridine reductase, tyrosine hydroxylase and, 238, 290 Dihydropyridines, oxidation of, 497498 Dihydroxyacetone, galactose oxidase and, 531 Dihydroxybenzene (s) , phenol hydroxylase and, 222 Dihydroxybenzoate (s) p-hydroxybenzoate hydroxylase and, 212, 213, 214, 215, 216 salicylate hydroxylase and, 207 2,3-Dihydroxyindole 2,3-dioxygenase, properties, 147-148 3,4-Dihydroxymandelate, protocatechuate 3,4-dioxygenase and, 137 3,4-Dihydroxyphenylacetate,protocatechuate 3,4-dioxygenase and, 137, 138 3,4-Dihydroxyphenylacetate 2,3-dioxygenase, properties of, 142-143 3,4-Dihydroxyphenylalanine, formation of, 238, 290 2,3-Dihydroxyphenylpropionate, formation of, 217 3,4-Dihydroxyphenylpropionate,protocatechuate 3,4-dioxygenase and, 137, 138, 139 3,4-Dihydroxyphenylpyruvate,p-hydroxyphenylpyruvate hydroxylase and, 181 6,&Dihydroxypurine excretion, allopurinol and, 401402 molybdenum hydroxylases and, 346 2,5-Dihydroxypyridine oxygenase, properties, 146-147 4,6-Dimercapto-allopurine, xanthine oxidase and, 364-365 Dimethoxythane, xanthine oxidase and, 353 4-Dimethylaminobutydate, y-butyrobetaine hydroxylase and, 168 4,5-Dimethylcatechol, formation of, 120 3,10-Dimethyl-8-cyanoisoalloxazine, nitroalkane oxidation and, 501-502 Dimethyl formamide, adrenodoxin and, 23

SUBJECT INDEX

e-N,N-Dimethyl-L-lysine, lysine monooxygenase and, 201 Dimethyloctylamine, microsomal amine oxidase and, 229-230 3,4-Dimethylphenol, oxidation of, 120 Dimethyl-p-phenylenediamine, ceruloplasmin and, 575, 576 Dimethyl sulfoxide adrenodoxin and, 23 superoxide solutions and, 540 6,7-Dimethyltetrahydropteridine phenylalanine hydroxylase and, 235, 237 tyrosine hydroxylase and, 239,240 Dinitroethane, formation of, 496 2,4-Dinitrofluorobenzene, xanthine oxidase and, 317 2,4-Dinitrophenylhydrazones,a-keto acid monitoring and, 428 Dioxygenase (8) , 2 classification, 121, 122 biological function and general properties, 123-127 heme-containing, 127-132 history and definition, 120-121 a-ketoglutarate, mechanism, 183-189 nonheme iron-containing, 132-151 Diphenols, tyrosinase and, 508 9,10-Diphenylanthracene-2,3-dicarboxylate, singlet oxygen and, 536 Diphosphopyridine nucleotide-ubiquinone reductase, properties of, 49 Dipyridyl amine oxidases and, 523 metapyrocatechase and, 141 nitrogenase and, 52 prolyl hydroxylase and, 161,163 steroid oxygenase and, 144 tryptophan hydroxylase and, 241 tyrosine hydroxylase and, 239 4,9-Diseco-3-hydroxyandrosta-l(10),2diene-5,9,17-trion-4-oicacid, formation of, 143-144 Dithiobis(2-nitrobezoate) flavodoxins and, 88 nitrogenase and, 52 xanthine oxidase and, 312 Dithioerythritol, xanthine oxidase and, 321

623 Dithionite adrenodoxin and, 267, 268 amine oxidases and, 518, 521 eight iron-sulfur protein and, 45 electron-transferring flavoprotein and, 116 external flavoprotein monooxygenases and, 206 flavodoxin and, 63, 103-104 flavin derivatives and, 86 four iron-sulfur proteins and, 33 p-hydroxybenzoate hydroxylase and, 212 4-methoxybenzoate Odemethyl-monooxygenase and, 287 molybdenum hydroxylases and, 395 monoamine oxidase and, 467 nitrogenase and, 50, 52, 53, 54, 55 protocatechuate 3,Cdioxygenase and, 137, 138 pyrocatechase and, 134, 135, 136 salicylate hydroxylase and, 208 sulfite oxdase and, 415, 417, 418 two iron-sulfur proteins and, 19, 29 xanthine oxidase and, 326-327, 332, 337-338,352, 360, 361, 369-371,375, 378-379, 388 Dithothreitol apoferredoxin reconstitution and, 27 2,5-di hydroxypyridine oxygenase and, 147 lysyl hydroxylase and, 166 prolyl hydroxylase and, 155-156, 157, 158, 163 thymine oxidation and, 175 Dodecyl sulfate amine oxidase and, 518 2,5-dihydroxypyridine oxygenase and, 147 electron-transferring flavoprotein and, 111 phenol hydroxylase and, 221 prolyl hydroxylase and, 155 superoxide dismutase and, 542 Dopamine amine oxidase and, 518 metapyrocatechase and, 141 Dopamine p-monooxygenase, properties of, 294-295 Double bonds, dioxygenases and, 123-125

SUBJECT INDEX

Double stopped-flow measurements, flavoprotein oxidases and, 440-442, 450451 Drosophila, xanthine oxidase of, 314 Drosophila melanogaster molybdenum hydroxylase of, 352 genetics of, 406412 molecular properties, 390-391 Drugs,hydroxylation of, 254, 257, 283

E Earthworm, prolyl hydroxylase of, 164 Echinocystis macrocarpa, cytochrome P-450 of, 280 Effective magnetic moment, rubredoxin, 6-7 Electron acceptors, flavoproteins and, 192 Electron density maps, flavodoxin, 67 Electron nuclear double resonance adrenodoxin and, 24 ferredoxins and, 21-22 ffavodoxins, 96 putidaredoxin, 263 Electron paramagnetic resonance amine oxidrtses and, 519-521 ascorbate oxidase, 567, 568 blue oxidases, 567-570, 576 ceruloplasmin, 562, 563, 567 cholesterol monooxygenase, 269 cytochrome P-420, 271, 272 cytochrome P-450,272,275,280-282 cytochrome P-45OC.,, 261-262 eight iron-sulfur proteins, 34-43, 45 ferredoxins, 2 flavin adenine dinucleotide, 327428 flavodoxin, 91, 95-96 four and eight iron-sulfur proteins, 34 galactose oxidase, 529-530, 532 high potential iron-sulfur proteins, 2 laccase, 563, 564, 567 4-methoxybenzoate O-demethyl-monooxygenase, 286 molybdenum hydroxylmes, 391, 394, 399 nitrogenase, 50-54 nonheme iron protein, phenylalanine 4-monooxygenase, 288-289 phenol o-monooxygenme, 297

phenylalanine hydroxylase, 237 putidaredoxin, 263 rubredoxins, 3, 7 118-steroid monooxygenase, 266 sulfite oxidase, 414,415-417,418 superoxide dismutase, 546, 547, 549, 550-551, 552, 553, 554 two iron-sulfur proteins, 19-20 perturbant effects, 23-25 xanthine oxidase, 315-316, 317, 322, 324-325, 363 flavin adenine dinucleotide and, 327-328 iron-sulfur systems, 339-342 molybdenum and, 32M39, 365 pre-steady-state kinetic studies, 372-374 reducing agents and, 369-371,374375, 378-381, 387 reoxidation and, 383-385 Electron spin resonance cysteamine oxygenase, 148-149 3,4-dihydroxyphenylacetate 2,3-dioxygenase, 142-143 protocatechuate 3,4-dioxygenase and, 139-140 pyrocatechase, 135-136 pyrocatechuate 4,5-dioxygenase, 143 Electron transfer reactions, two iron-sulfur proteins, 29-31 Electron-transferring flavoprotein catalytic properties, 116-118 function, 57, 109-110 molecular properties, 111-116 occurrence, 109-110 Endoplasmic reticulum prolyl hydroxylase and, 162 xenobiotic monooxygenases in, 269-270 Enolates, flavoprotein oxidases and, 428-129 Epinephrine oxidation, superoxide and, 533-534, 540 xanthine oxidase and, 357 Epinine, dopamine 8-monooxygenase and, 294 Equisetum, ferredoxin of, 15 Erythrocuprein, 533 function of, 510 Erythrocytes, superoxide dismutase of, 539,541,544,547

625

SUBJECT INDEX

Esche)ichia coli flavodoxin, 59 absorption maxima, 99 iron and, 64 redox potentials, 99 sulfite reductase of, 48 superoxide dismutases of, 537 two iron-sulfur protein of, 19 Ethanol ferredoxin and, 23 protocatechuate 4,bdioxygenase and, 143 7-Ethoxycoumarin, cytochrome P-950 and, 275,276 Ethylamine, amine oxidases and, 521 Ethylenediaminetetraacetate L-amino acid oxidase and, 457 cysteine oxygenase and, 149 electron-transferring flavoprotein and, 115 external flavoprotein monooxygenases and, 205, 206, 208 galactose oxidase and, 529 p-hydroxybenzoate hydroxylse and, 211 superoxide dismutase and, 548 thymine 7-hydroxylase and, 174 xanthine oxidase and, 308, 323 Ethylene glycol adrenodoxin and, 23 apoflavodoxin and, 82 Ethylisocyanide, cytochrome P-950,., and, 261 N-Ethylmaleimide flavodoxins and, 88 xanthine oxidse and, 368 Ethylmercaptan, flavoprotein oxidases and, 495 Ethylmorpholine, galactose oxidase and, 531 Ethylprotocatechuic acid, pyrocatechase and, 134, 135, 136 Ethyl pyruvate, model reactions and, 503 Exopeptidases, ferredoxins and, 28

F Fatty acids excretion of, 110 phenylalanine hydroxylase and, 235

Ferredoxin(s) bacterial background, 37-39 chemical properties, 4 4 4 6 physical properties, 3 W 4 characteristics of, 2 flavodoxins and, 58,60 molybdenum hydroxylases and, 397, 398 two irons per center background, 15-20 chemical properties, 25-31 perturbants of EPR spectra, 23-25 physical properties, 20-23 Ferredoxin-nicotinamide adenine dinucleotide phosphate-reductse flavodoxins and, 62,03,79,98,108 ionic strength and, 31 Ferric oxide, MGssbauer parameters, 11 Ferric tris-pyrollidyl dithiocarbamate, Mossbauer parameters, 11 Ferricy anide n-amino acid o x i d m and, 452 electron-transferring flavoprotein and, 116 ferredoxin and, 22,26 flavodoxins and, 107-108 four iron-sulfur proteins and, 33 galactose oxidase and, 530, 532 4-methoxybenzoate Odemethyl-monooxygenase and, 286 molybdenum hydroxylases and, 397, 398 nitrogenase and, 53 Old Yellow Enzyme and, 473 protocatechuate 3,ldioxygenase and, 138 sulfite oxidase and, 417, 418 xanthine oxidase and, 320,353,364,359, 364, 367 Ferricytochrome c, superoxidase dismutase assay and, 539,552-553 Ferriprotoporphyrin IX, cytochrome P-450,., and, 262 Ferritin, 2 Ferrocyanide laccase and, 572 superoxide dismutase and, 548, 551-552 Ferroxidase activity, 509 blue oxidases and, 576

626 Flavin(s) chemical mechanism of reduction chemical model systems, 498-503 inhibitors and coenayme analogs, 497-498 model substrates, 484-497 models, 474-475, 484 physiological substrates, 475 electron-transferring flavoprotein and, 111-113 monoamine oxidase and, 467 Flavin adenine dinucleotide amino acid oxidase and, 446-447,480 L-amino acid oxidase and, 456, 457 arginine monooxygenaae and, 204 camphor 5-em-monooxygenase and, 263 cytochrome c reductase and, 273 electron-transferring flavoprotein and, 111, 113, 115, 117 flavodoxins and, 84 protein modifications affecting binding, 87-88 flavoprotein oxidases and, 436 glucose oxidase and, 461-462 m-hydroxybenzoate-4-hydroxylaseand, 225 p-hydroxybenzoate hydroxylaae and, 211, 212, 213-214, 216 imidazolylacetate monooxygenase and, 226 kynurenine-3-hydroxylaseand, 230-231 lysine monooxygenase and, 200 melilotate hydroxylase and, 217,218 microsomal amine oxidase and, 229 molybdenum hydroxylases and, 399 monoamine oxidase and, 470, 471 orcinol hydroxylase and, 223,224 phenol hydroxylase and, 221, 223 salicylate hydroxylase and, 206, 207, 208, 210 11p-steroid monoogygenase and, 267 xanthine oxidase and, 308,310, 312, 313, 315, 316, 322, 352, 353 alkylation of, 319-320 magnetic and optical properties, 327428, 342-343,363, 364 oxygen and, 356, 384,385,386 pre-steady-state kinetics and, 372373

SUBJECT INDEX

reduction and, 375, 376, 379 removal of, 318-319 substrate and, 365, 367 Flavin coenzyme (s) , structure, and chemistry, 423-425 Flavin mononucleotide bacterial luciferase and, 227, 228 cytochrome c reductase and, 273 derivatives, binding by flavodoxins, 84, 85-87 flavodoxins and, 59, 65-66,70 binding site, 74-78 thermodynamics and kinetics of binding, 83-85 lactate oxidase and, 194 4-methoxybenroate O-demethyl-monooxygenase and, 286 redox potentials, 99 xanthine oxidase and, 319 Flavodoxins composition and molecular weight, 59-60. 99 discovery and nomenclature, 58-59 distribution, 59 flavin mononucleotide in, 65-66 derivatives, 84, 85-87 protein modifications in binding, 87-88 thermodynamics and kinetics of binding, 83-85 function, 60-63 oxidation-reduction potentials, 98-102 purification, 60 reactivity comproportionation, 102-103 dithionite, 103-104 oxygen and ferricyanide, 105-108 redox proteins and, 108-109 regulation by iron, 63-65 spectroscopic properties circular dichroism, 94-95 fluorescence, 93-94 magnetic resonance, 95-98 optical absorption spectra, 88-91,W single crystal absorbance, 91-93 structure, 66-67 determination and comparison of chemical sequences, 67-70 flavin binding site, 74-78 molecular configuration, 71-74

SUBJECT INDEX

oxidation state and, 79-82 residue distribution, 78-79 structure determination, 70-71 Flavoprotein camphor 5-ezo-monooxygenase and, 263 dioxygenases and, 122 as electron donors other systems, 279-280 xenobiotic monooxygenases, 269-279 pyridine nucleotide-linked, monooxygenases and, 122 reaction with oxygen, 192 llp-steroid monooxygenase and, 265, 267 Flavoprotein oxidase(s) chemical mechanism 'flavin reduction, 474-503 oxidation by oxygen, 503-505 definition of, 421-423 deuterium kinetic isotope effect, 482 function of, 423 kinetics computer simulation, 442443 mechanism, 435442 steady-state, 426-432 strategy, 425-426 summary, 443-445 transient-state : the half reaction, 432-435 long wavelength (purple) intermediate, 479-482 reductive half-reaction, 475-479 Fluorenylcarbanion, cytochrome P-450 and, 276 Fluorescence n-amino acid oxidase, 446, 447 apoflavodoxins, 82 electron-transferring flavoproteins, 115 flavodoxin, 93-94,102 flavin binding and, 83, 85, 87 p-hydroxybenzoate hydroxylase, 212 salicylate hydroxylase, 207-208, 209 Fluoride blue oxidases and, 569, 573, 577-578 cytochrome P-420 and, 271 galactose oxidase and, 530 glucose oxidase and, 465 superoxide dismutase and, 548 Fluorophenol(s), phenol hydroxylase

627 and, 222 Fluorophenylalanine (s) , phenylalanine hydroxylase and, 234, 235 Fluorophenylpyruvate, p-hydroxyphenylpyruvate hydroxylase and, 181 Formaldehyde molybdenum hydroxylases and, 396 reduction of, 502403 xanthine oxidase and, 324-325,332, 382 Formate, galactose oxidase and, 531 Formylkynurenine, formation of, 128, 130, 131 5-Formyluracil, formation of, 170,173, 187 5-Formyluracil dioxygenase, properties, 175-176 N-Formylmaleamate, 2,5-dihydroxypyridine oxygenase and, 147 Free radical(s), blue oxidases and, 575-576 Free radical flavoprotein, 59 Fumarate, dopamine p-monooxygenase and, 294,295 Furoin, oxidation, 0avin and, 501 Fusarium oxysporum, superoxide dismutase of, 547

Galactohexodialdose, galactose oxidase and, 530 n-Galactosamine, galactose oxidase and, 531 Galactose galactose oxidase and, 530-531 glucose oxidase and, 436, 444, 462 Galactose oxidase, 509 chemical and physical properties, 528-529 copper of, 529-530 discovery and purification, 527-528 inhibitors, 531 mechanism, 532-533 optical properties, 530-531 specificity, 531 superoxide and, 557 Gibberellin, formation of, 280 n-Glucal, glucose oxidase and, 465-466 Glucocorticoids, tryptophan 2,3dioxygenase and, 128

628 n-Glucono-&lactone, glucose oxidase and, 466 Glucose m a y of, 428 deuterated, glucose oxidase and, 463-464 glucose oxidase and, 436,443-444,462 oxidation, thermodynamics, 499 Glucose oxidase enzyme-monitored turnover, 437439 integrated steady-state equation, 431-432 kinetic mechanism, 436,443-444 molecular properties and kinetic mechanism, 461-466 nitroalkanes and, 492-497 reductive half-reaction, 434,476 turnover of, 430 Glutamate residues, flavodoxin, 79,81 Glutathione dioxygenase and, 125 homogentisate oxygenase and, 146 thymine oxidation and, 174, 175 Glycerol cytochrome P-450 and, 272 ferredoxin and, 23 Glycine lysyl hydroxylase and, 166 prolyl hydroxylase and, 158 Glycine residues flavodoxins, 69,76 oxidation state and, 80 Glycollate oxidase, flavin binding site, 114 Glyoxal, n-amino acid oxidase and, 446 Glyoxylate oxidase, flavin and, 422 Gold, ferredoxin structure and, 70 Gout, molybdenum hydroxylase genetics and, 400-402 Guanidinium chloride amine oxidases and, 513,516,518 electron-transferring flavoprotein and, 113 flavodoxin and, 82 prolyl hydroxylase and, 155 superoxide dismutase and, 542, 543 xanthine oxidase and, 314, 362 7-Guanidinobutyramide, formation of, 203 Guran, galactose oxidase and, 531

SUBJECT INDEX

H Half-reactions flavoprotein oxidases oxidative, 435 reductive, 432-434 Halides blue oxidases and, 569,575 glucose oxidase and, 465, 466, 483 a-Helix flavodoxins, 67,71,72,73,76 metapyrocatechase, 141 phenol o-monooxygenase, 297 pyrocatechase, 136 Heme amine oxidases and, 518 sulfite oxidase and, 415,418,419 Hemerythrin, 2 Hemin, monoamine oxidase and, 468 Hemocuprein, 533 function of, 510 Hemoglobin, superoxide and, 534 Hemopexin, cytochrome P-450,., and, 262 Hemoprotein (s) dioxygenases and, 122, 127-132 flavin-linked, monooxygenases and, 122 iron-sulfur protein-linked, monooxygenases and, 122 Hepatocuprein, 533 Heptylamine, amine oxidages and, 514, 516 Hexammineruthenium(II), rubredoxin and, 14-15 n-Hexane, cytochrome P-450 and, 279 Histamine, amine oxidases and, 513, 514, 517, 518, 522 Histidine, catabolism of, 2252226 Histidine residues arginine monooxygenase and, 204 bacterial ferredoxins and, 46 blue oxidases, 570 flavodoxins, 60 superoxide dismutase and, 547, 552 Homoarginine, arginine monooxygenase and, 203 Homogentisate, formation of, 179-180, 181-182 Homogentisate oxygenase, properties, 145-146

SUBJECT INDEX

Hydrazine amine oxidases and, 512,519, 521, 522, 523 galactose oxidase and, 531 monoamine oxidase and, 471 Hydride abstraction, flavoprotein oxidases and, 475, 482, 491 Hydrocarbons, perfluorinated, cytochrome P-450 and, 279 Hy drbgenase flavodoxin and, 61,63,65,98 properties of, 48 Hydrogen atom abstraction, flavoprotein oxidases and, 475,482, 491 Hydrogen bonding flavodoxins, 72-73, 80 flavin and, 74, 77,78, 81,86 Hydrogen chloride, n-amino acid oxidase and, 485, 489,491, 497 Hydrogen peroxide amine oxidases and, 511412,526 D-amino acid oxidase and, 452, 453 bacterial luciferase and, 228 blue oxidases and, 570, 578, 579 copper-containing oxidases and, 509, 511-533 double stopped-flow measurements and, 441442 flavin monooxygenases and, 192, 194, 205 flavin oxidation and, 502 flavoprotein oxidases and, 427, 428, 435 galactose oxidase and, 530,531, 532 glucose oxidase and, 438,463, 405,496 p-hydroxyphenylpyruvate hydroxylase and, 181 imidazolylacetate monooxygenases and, 226 indoleamine 2,3-dioxygenase and, 131 lactate oxidase and, 198 lysine monooxygenase and, 202 metapyrocatechase and, 141 4-methoxybenaoate O-demethyl-monooxygenase and, 287 monoamine oxidase and, 469 monooxygenases and, 24P243, 245, 247 phenylalanine hydroxylase and, 234, 236 phenylalanine 4-monooxygenme and, 289

629 salicylate hydroxylase and, 210 sulfite oxidase and, 417 superoxide dismutase and, 551, 552, 554, 555 superoxide radical and, 357, 358 tryptophan 2,3dioxygenase and, 129, 130 tyrosine hydroxylase and, 239,291 xanthine oxidase and, 302-303,325,326, 355, 369, 384 Hydrogen sulfide, galactose oxidase and, 529 Hydroperoxide, monooxygenases and, 242, 243 Hydrophobic interactions, amine oxidases, 524-625 4-Hydroxy-6-aminopyrimidine,molybdenum hydroxylases and, 348 3-Hydroxyanthranilate oxygenase, properties, 144-145 4-Hydroxy-7-azapteridine, xanthine oxidase and, 349,351 Hydroxybenzoate(s), salicylate hydroxylase and, 207 m-Hydroxybenzoate-4-hydroxylase,247 properties, 225 m-Hydroxybenzoate-6-hydroxylase,properties, 224-225 p-Hydroxybenzoate hydroxylase, 247 properties, 211-216 L-2-Hydroxy3-butynoic acid, lactate oxidase and, 199, 498 4-Hydroxycatechol, pyrocatechase and, 134 20a-Hydroxy cholesterol cholesterol monooxygenase and, 269 cytochrome P-450 and, 266 6-Hydroxy-7,8-dimethyl-lO-(ribityld’ADP)-isoalloxazine, electron-transferring flavoprotein and, llP115, 116, 117 l-(2-Hydroxyethyl) -2-methyl-nitroimidaaole, xanthine oxidase and, 366 L-a-Hydroxyisovalerate, lactate oxidase and, 197 Hydroxylamine amine oxidases and, 519,523,527 cysteamine oxygenase and, 148 cysteine oxygenase and, 149 flavoprotein oxidases and, 495

SUBJECT INDEX

galactose oxidase and, 531 Hydroxyl radicala superoxide and, 535 xanthine oxidase and, 358 5-Hydroxylysine, function of, 165 p-Hydroxymercuribenzoate, cysteine oxygenase and, 149 Hydroxymethylhydroperoxide, xanthine oxidase and, 325 5-Hydroxymethyluracil, formation of, 170, 173, 175 5-Hydroxymethyluracil deoxyribonucleoside, 2’-hydroxylase and, 178 5-Hydroxymethyluracil dioxygenase, properties, 17&176 L-a-Hydroxy-p-methylvalerate,lactate oxidase and, 197 a-Hydroxymuconic r-semialdehyde, formation of, 140 l-Hydroxy-2-naphthoate, salicylate hydroxylase and, 207 6-Hydroxynicotinate, p-hydroxybenzoate hydroxylase and, 213, 214 2-HydroxyS-nitrobensylbromide,rubredoxin and, 12 p-Hydroxyphenylpyruvate hydroxylase, 179-180 catalytic properties, 180-183 purification and molecular properties, 180 Hydroxyproline, formation of, 152 4-Hydroxyprazolo (3,4-d) pyrimidine, 8ee Allopurinol 2-Hydroxypyrimidine, molybdenum hydroxylases and, 347 Hy droxyquinol orcinol hydroxylase and, 223 phenol hydroxylase and, 222 &Hydroxy quinoline amine oxidases and, 523 steroid oxygenase and, 144 p-Hydroxysalicylate, salicylate hydroxylase and, 210 7-Hydroxy-(1,2,5)-thiadiarolo (3,4&pyrimidine, molybdenum hydroxylases and, 348 &Hydroxytryptophan, indoleamine 2,3dioxygenase and, 132 Hypotaurine formation of, 125, 148

xanthine oxidase and, 324 Hypoxanthine allopurinol and, 302 molybdenum hydroxylases and, 346, 350, 359 oxidation, xanthinuria and, 401

I Imidazolylacetate monooxygenase, properties, 225-226 Imino acid(s) o-amino acid oxidase and, 452,455456, 480, 487, 489, 491 L-amino acid oxidase and, 461 Imino compound(s), monoamine oxidase and, 469 Indigo disulfonate flavodoxins and, 98 4-methoxybenaoate O-demethyl-monooxygenase and, 287 Indoleamine 2,3-dioxygenase historical, 130-131 molecular and catalytic properties, 131-132 Indomesathine, prostaglandin synthetase and, 127 Indophenol, see Dichlorophenolindopheno1 Iodine, flavodoxin and, 88 Iodoacetamide, xanthine oxidase and, 316, 319-320 Iodoacetate cysteine oxygenase and, 149 rubredoxin and, 12 Infrared spectra, two iron-sulfur proteins, 21 Iron apoferredoxin reconstitution and, 27 v-butyrobetaine hydroxylase and, 168 ceruloplasmin and, 558, 575, 576 cysteamine oxygenase and, 148-149 3,4-dihydroxyphenylacetate 2,3-dioxygenase and, 142 dioxygenases and, 123, 124, 150-151 flavodoxin regulation by, 63-65 5-formyluracil dioxygenase and, 175 homogentisate oxygenase and, 146 3-hydroxyanthranilate oxygenase and, 145

631

SUBJECT INDEX

5-hydroxymethyluracil dioxygenase and, 175 p-hydroxyphenylpyruvate hydroxylaae and, 180,181 a-ketoglutarate dioxygenases and, 151, 185-186 lipoxygenase and, 150 lysylhydroxylase and, 166, 294 metapyrocatechase and, 140-141, 142 monoamine oxidase and, 468 phenylalanine hydroxylase and, 237, 251, 289 prolyl hydroxylase and, 157, 158, 163, 164, 185, 293 protocatechuate 3,4dioxygenase and, 137, 138, 139-140 protocatechuate 4,5-dioxygenase and, 143 pteridine-linked monooxygenases and, 193, 231 putidaredoxin and, 263 pyrimidine deoxyribonucleoside 2'hydroxylase and, 177, 178 pyrocatechases and, 134, 135, 136 removal from rubredoxin, 12 steroid oxygenase and, 144 superoxide dismutase and, 537 thymine 7-hydroxylase and, 174 tryptophan hydroxylase and, 241, 292 tyrosine hydroxylase and, 239, 291 xanthine oxidase and, 310,315 Iron proteins, classes of, 2 Iron-sulfur enzymes, 49-50 mitochondrial, 56 nitrogenase system, 5056 xanthine oxidase, 56 Iron-sulfur proteins camphor Bezo-monooxygenase and, 263 conjugated, characteristics, 3 eight irons background, 37-39 chemical properties, 44-46 physical properties, 39-44 four irons per center background, 31-33 high potential proteins, 35-37 low potential proteins, 34 4-methoxybenzoate Odemethyl-monooxygenase and, 284

model compounds, 46-47 high potential, characteristics of, 2 llp-steroid monooxygenase and, 265, 267, 268 Iron-su lfur systems molybdenum hydroxylases and, 394, 396, 399 xanthine oxidase, 367, 372-374, 375, 376, 378, 379, 383, 385,386 magnetic and optical properties, 339-342, 343, 363, 364 Isoalloxazine flavin binding and, 447 flavodoxins and, 74-75, 76,77-78,79 glucose oxidase and, 462 Isoamylalcohol, galactose oxidase and, 529 Isoascorbate, thymine 7-hydroxylase and, 174 Isocaproic aldehyde, cholesterol cleavage and, 268 Isoleucine residues, flavodoxins, 69,71 Isonicotinic acid hydrazine, amine oxidases and, 523, 527 Isopropylcatechol(s) , steroid oxygenase and, 144 Isotope effects D-amino acid oxidase, 448, 450, 477, 479,482,485,486437,488,489, 491 L-amino acid oxidase and, 460-461 flavin reduction and, 482, 485, 486 glucose oxidase and, 463-464 xanthine oxidase, 376-377, 382 Isozymes, L-amino acid oxidase, 456

K Kaur-IB-ene, monooxygenase and, 280 Keto acids o-amino acid oxidase and, 480 formation, monitoring of, 428 Ketoadipates, pyrimidine deoxyribonucleoside 2'-hydroxylase and, 177 a-Ketobutyrate, deuterium incorporation into, 486-487 a-Ketoglutarate y-butyrobetaine hydroxylase and, 168, 169 dioxygenases and, 121, 151-152, 183-189

632

SUBJECT INDEX

5-formyluracil and, 175 5-hydroxymethyluracil dioxygenase and, 175 lysyl hydroxylase and, 166, 294 prolyl hydroxylase and, 157,158, 164, 185, 293 pyrimidine deoxyribonucleoside 2’-hydroxylase and, 177, 178 thymine 7-hydroxyhe and, 174,175 Kidney, xenobiotic monooxygenases in, 270 Kinetic constants lactate oxidase, 197 melilotate hydroxylase, 218-219 Kbbsiella pneumoniae, nitrogenase of, 52, 53, 55 Kynuramine, amine oxidases and, 516, 518 Kynurenine, formation of, 127-128 Kynurenine-3-hydroxylase, properties, 230-231

1 Laccase, 509 anaerobic reduction, 577 copper of, 561,564,565,566,570 copper and, 508 discovery of, 557-558 distribution and function, 558-559 molecular properties, 561 oxidation-reduction potentials, 572, 573 purification of, 560 specificity, 574 Lactate o-amino acid oxidase and, 477 fermentation, iron deficiency and, 64 prolyl hydroxylase and, 163 Lactate dehydrogenase electron-transferring flavoprotein and, 110, 115, 116-117 Ravodoxin and, 73, 77 Lactate oxidase inhibition of, 498 nature of, 187-188 properties, 194-199 Lactose, galactose oxidase and, 531 Leuceana glauca, ferredoxin, amino acid sequence, 17

Leucine D-amino acid oxidase and, 444, 450, 454-465 deuterated, L-amino acid oxidase and, 460-461 spinach ferredoxin and, 28 Leucine aminopeptidase, spinach ferredoxin and, 28 Leucine residues, flavodoxins, 69, 71,97 Lignin, decomposition, laccaae and, 559 Lipase, cytochrome P-450 and, 273 Lipids, superoxide and, 535 a-Lipoate, singlet oxygen and, 536 Lipoxygenase properties, 150 reaction and cofactors, 127 Liver xanthine oxidase of, 315 xenobiotic monooxygenases in, 26S270 Low pyridoxal oxidase mutant, chromosome locus, 407 Low xanthine dehydrogenase mutant, chromosome locus, 407 Luciferase, bacterial, properties, 226-229, 248 Lumiflavin ferricyanide and, 108 flavodoxins and, 84,87 N-methyl dihydronicotinamide and, 473 as model compound, 423-424 Luminescence, xanthine oxidase and, 358 Luminol, superoxide and, 536-537 Lung, xenobiotic monooxygenases in, 269-270, 271 Lysine, amine oxidase and, 514 Lysine hydroxamate, lysine monooxygenase and, 201 Lysine monooxygenase, properties, 199203,293-294 Lysine residues flavodoxins, 69, 79 rubredoxin, 12 Lysine-vasopressin, amine oxidase and, 527 Lysolecithin, phenylalanine hydroxylaae and, 234, 235, 236, 289 Lysophosphatidylethanolamine, monooxygenase and, 280 Lysyl hydroxylase

633

SUBJECT INDEX

catalytic properties, 166-167 purification and molecular properties, 165 Lysyl oxidase, 466

M Magnesium, nitrogenase and, 5(M2 Magnetic circular dichroism, superoxide dismutase, 545-546, 550 Magnetic field, rubredoxin and, 9 Magnetic optical rotatory dispersion, superoxide dismutaae, 545-546 Magnetic susceptibility, two iron-sulfur proteins and, 21 Maleylacetoacetate, formation of, 145 Mammals, electron-transferring flavoprotein of, 115 Mandelic ester, oxidation, flavin analog and, 500-501 Manganese spinach ferredoxin and, 28 superoxide dismutase and, 537 Mannose, glucose oxidase and, 436,444, 462, 464 Maroon-like mutant chromosome locus, 407 maternal effects, 410 Melatonin, dioxygenases and, 123, 132 Melibiose, galactose oxidase and, 531 Melilotate hydroxylase, 247 properties, 217-221 Membranes, xanthine oxidase and, 322 Mercaptoethanol amine oxidases and, 513, 516, 518 flavoprotein oxidases and, 495 galactose oxidase and, 528 proiyf hydroxylase and, 155 superoxide dismutase and, 542 thymine 7-hydroxylase and, 174 Mercurials cytochrome P-450,., and, 262 flavodoxins and, 8 2 , s spinach ferredoxin and, 21-22 xanthine oxidase and, 311 p-Mercuribenzoate 3,4dihydroxyphenylacetate 2,3-dioxygenase and, 142 imidazolylacetate monooxygenase and, 226

phenol hydroxylase and, 223 prolyl hydroxylase and, 158 Mercury, superoxide dismutase and, 549 Mersalyl ferredoxins and, 26 flavodoxins and, 87-88 Mescaline, amine oxidase and, 516, 518 Metals, nitrogenase and, 51 Metapyrocatechase circular dichroism, 141-142 molecular and catalytic properties, 140-141 Methanol ferredoxin and, 23, 24 molybdenum hydroxylases and, 396 oxidation, thermodynamics, 499 xanthine oxidase and, 324-325,332 Methional, xanthine oxidase and, 357-358 Methionine residues flavodoxins, 69, 78, 101-102 putidaredoxin, 20 rubredoxin Clostridium pasteurdanzlm, 12 Pseudomonas oleovorans, 13 superoxide dismutase, 545 4-Methoxybenzoate Odemethyl-monooxygenase, properties, 285-287 4Methoxycatecho1, pyrocatechaae and, 134 Methylcatechols metapyrocatechase and, 141 protocatechuate 3,4-dioxygenam and, 137 pyrocatechases and, 134, 135 N-5-Methyl-1,5-dihydroflavin, ethyl pyruvate and, 503 N-Methyl dihydronicotinamide, lumiflavin and, 473 Methylene blue indoleamine 2gdioxygenase and, 131 sulfite oxidase and, 417 xanthine dehydrogenase and, 409 xanthine oxidase and, 353 Methyl ethyl ketone, monoamine oxidase and, 468 Methyl-n-galactopyranosides, galactose oxidase and, 531 N-Methylglutamate synthetase, 6-deazariboflavin and, 498 l-Methyl-2-hydroxypurine, molybdenum

634 hydroxylases and, 347 2-Methyl-3-hydroxypyridine-5-carboxylate, dioxygenase and, 123, 127 7-Methyl-8hydroxy-lO-( ribityl-5’ADP)-isoalloxadne, electron-transferring flavoprotein and, 112-115, 116, 117 3-Methylhypoxanthine, molybdenum hydroxylases and, 347, 352 N’-Methylnicotinamide, molybdenum hydroxylases and, 348 351, 362 Methyloctylsulfide, cytochrome P-450 and, 276 Methylphenol(s), phenol hydroxylase and, 222 ZMethylpropane diol, ferredoxin and, 23 SMethylsalicylate, salicylate hydroxylase and, 207 6-Methyl tetrahydropteridine phenylalanine hydroxylase and, 235 tyrosine hydroxylase and, 239 7-Methyltetrahydropteridine, phenylalanine hydroxylase and, 235 @-Methylthiopropionaldehyde, xanthine oxidase and, 357-358 Methyl viologen, flavodoxins and, 104 1-Methylxanthine, molybdenum hydroxylases and, 347, 374, 375, 376-378 Metmyoglobin, cytochrome P-450 and, 280, 281 Metyrapone cytochrome PA50 and, 276 cytochrome P-450,,,,, and, 260-261 Micrococcus aerogenes ferredoxin of, 44 oxypurine dehydrogenase of, 389 rubredoxin of, 13 Micrococcus denitrificcans, nitrate reductase of, 48 Micrococcus lactylyticus ferredoxin, electron paramagnetic resonance, 4 W 2 iron-sulfur protein of, 32 Microsomes, hydroxylation reactions in, 254 Milk xanthine oxidase activation energy, 361 assays for, 358-360 catalytic properties mechanism of action, 365-388

SUBJECT INDEX

reactions catalyzed, 344-365 deflavo form, 318-319,352, 353, 354, 367, 381 allopurinol and, 364 demolybdo form, 306-309, 317 desulfo form, 307-308,316-317,322324, 352,383 electron paramagnetic resonance, 332,337,338 reduction of, 327,371, 387 historical background, 303-304 inhibition, 362-365 excess reducing substrate and, 386387 isotope effects, 376377 molecular properties chemical modification, 317-326 composition and physical properties, 310-317 magnetic interactions of redox groups, 342-344 purification, 304-310 redox group magnetic and optical properties, 326342 one- and two-electron reduction by, 355-358 overall catalytic process, 386 pre-steady-state kinetic studies, 372-374 rates of internal electron transfer processes, 385-386 redox-active groups, “additional,” 326-327 magnetic and optical properties, 326-342 magnetic interactiom and locations, 342-344 reducing substrates and, 368 reduction of, 374-382 slow phases, 387-388 reoxidation by oxygen, 382385 reversibility of reaction, 360-361 sites of substrate and inhibitor interaction, 365-366 specific activity, 309-310, 311 specificity oxidizing substrates, 353-355 reducing substrates, 344-353 steady-state kinetic analysis, 368369 titration with reducing agents, 369372

635

SUBJECT INDEX

turnover number, 361 Mitochondria iron-sulfur centers, properties, 49,56 kynurenine-3-hydroxylase of, 230 monoamine oxidase and, 467 Ilp-steroid monooxygenase of, 265-268 superoxide dismutase of, 537 Model systems, chemical, flavoprotein oxidases and, 498-503 Mossbauer spectra high potential iron proteins and, 10, 35, 37 nitrogenase, 53, 55 putidaredoxin, 263 rubredoxin, 8-12 two iron-sulfur proteins, 21 xanthine oxidase, 341-342 Molybdenum isotopes, electron paramagnetic resonance and, 329-330 maroon-like mutant and, 410 molybdenum hydroxylases and, 394395, 396397 nitrate reductase and, 404,406 sulfite oxidase and, 414, 418 xanthine oxidase, 388 allopurinol and, 363, 364 content, 307-309,310, 315 magnetic and optical properties, 328-339, 343 pre-steady-state kinetics and, 372374 substrate and, 365, 367, 376,378, 379382, 386 Molybdenum hydroxylases assays for, 358-360 distribution and biological importance, 301303 general considerations, 300-301 genetic studies Aspergillus nidulans, 412414 Drosophila melanogaster, 406-412 nitrate reductase and, 402-406 xanthinuria and gout in man, 400-402 other, 388-389 catalytic properties, 394-400 molecular properties, 389-394 oxidizing substrates, 397400 reduction mechanism, 394-397 Monoamine oxidase(s)

characteristics of, 512 inhibitors of, 470-471, 498 intracellular localization, 467 molecular properties and kinetic mechanism, 466-471 physiological role, 466 types of, 466 Monooxygenase (9) classification of, 121-123 copper-containing, 294-297 external, 204-206 flavoprotein, electron donors and, 192 heme-containing flavoproteins and, 269-280 general mechanisms, 280-285 iron-sulfur proteins as electron donors, 259-269 history and definition, 120-121 internal, 193-204 iron- and copper-containing functions, 257-258 historical aspects, 253-255 nomenclature, 256-257 role in oxygen activation, 256 iron-containing, heme, 258-285 nonheme, 285-294 model studies and poasible mechanisms, 241-252 pterin-linked, 231 reactions catalyzed, 257 Monophenols, laccase and, 574 cis,cis-Muconic acid, formation of, 120, 133 Mushroom, phenol 0-monooxygenase of, 296-297 Mycobacterium phlei, lactate monooxygenase of, 194-195 Mycobacterium smegmatis, lactate monooxygenase of, 195-196 Myoglobin, xanthine oxidase and, 366

N a-Naphthoquinone, metapyrocatechase and, 141 Naphthylamines, microsomal amine oxidase and, 229 Neocuproine, monoamine oxidase and, 527 Neurospora, superoxide dismutase of,

636 539, 541, 544, 547 Neurospora crassa nitrate reductase, 403-405 thymidine salvage in, 169-172, 178-179 Nicotinamide adenine dinucleotide Aspergillus xanthine dehydrogenases and, 412 bacterial luciferase and, 227 camphor 5-eso-monooxygenase and, 259, 262263 cytochrome P-420 and, 271 bdearaflavin and, 497-498 demolybdo xanthine oxidase and, 309 desulfo xanthine oxidase and, 319, 324, 338 dihydrobiopterin reductase and, 233 dioxygenaaes and, 124 Drosophila xanthine dehydrogenase and, 409 electron-transferring flavoprotein and, 110, 113, 116, 117 flavoprotein oxidaaes and, 428 m-hydroxybenzoateShydroxy1we and, 224 imidazolylacetate monooxygenase and. 226 kynurenine-3-hydroxylase and, 230 melilotate hydroxylase and, 217-218 microsomal monooxygenases and, 279 molybdenum hydroxylases and, 348, 352, 353,397, 398,399-400 Old Yellow Enzyme and, 471473 orcinol hydroxylase and, 223-224 salicylate hydroxylase and, 207,208, 209, 210 sulfite oxidase and, 416 tyrosine hydroxylase and, 238 xanthine dehydrogenase and, 320, 321, 322,345,361,365,386,381,387 Nicotinamide adenine dinucleotidedihydropteridine reductase, monooxygenases and, 231 Nicotinamide adenine dinucleotide phosphate cysteine oxygenase and, 149 cytochrome c reductase and, 273 cytochrome P-420 and, 271 cytochrome P-450 and, 272,277-278, 279

SUBJECT INDEX

7,8dihydrobiopterin reduction and, 233 ferredoxin reduction and, 31 gibberellin formation and, 280 m-hydroxybenzoate-4-hydroxylaae and, 225 m-hydroxybenzoate-6-hydroxylase and, 224 p-hydroxybenroate hydroxylase and, 211, 212, 213-214 microsomal amine oxidase and, 229 Old Yellow Enzyme and, 471473 orcinol hydroxylase and, 224 phenol hydroxylase and, 221,222,223 salicylate hydroxylase and, 207, 208 llp-steroid monooxygenase and, 265, 267, 268 tryptophan bmonooxygenase and, 292 yeast cytochrome P-450 and, 280 Nicotinamide adenine dinucleotide phosphate-cytochrome c reductase, nitrate reductase and, 403, 405 Nicotinamide N-oxide, xanthine oxidase and, 355,367 Nicotinic acid, xanthine dehydrogenases and, 412 NIH shift aromatic hydroxylations and, 219-221 phenylalanine hydroxylase and, 234-235 tyrosine hydroxylase and, 240 Nitrate reductase molybdenum hydroxylase “common cofactor,” 402406 properties of, 48 Nitric oxide ceruloplasmin and, 571 cytochrome P-450 and, 276 Nitrite reductase, iron deficiency and, 65 Nitroalkanes flavoprotein oxidases and, 492497 glucose oxidase and, 483 0-Nitrobenzoate, salicylate hydroxylase and, 207 p-Nitrobenzyl chloride, alkylation of, 502 Nitro blue tetrazolium superoxide dismutase assay and, 539-540 xanthine oxidase and, 354,357,359 4-Nitrocatechol, phenol 0-monooxygenase

637

SUBJECT INDEX

and, 297 Nitroethane anion D-amino acid oxidase and, 476, 481, 492 glucose oxidase and, 4 W 9 7 Nitrogen molybdenum bonding and, 337, 339 reduction, flavodoxins and, 61 Nitrogenase fiavodoxin and, 108, 109 iron deficiency and, 65 properties of, 48, 50-56 Nitromethane, glucose oxidase and, 483, 492 o-Nitrophenol, metapyrocatechase and, 141 2-Nitropropane, flavoprotein oxidases and, 495 2-Nitropropyl anion, alkylation by, 502 Nocardia restrictus, steroid oxygenase of, 144 Nonheme iron protein(s) dioxygenases and, 122, 132-151 phenylalanine 4-monooxygenase and, 288-239 superoxide and, 533 Norepinephrine amine oxidases and, 466, 514, 516 formation of, 294 tyrosine 3-monooxygenase and, 290 Norvaline, D-arniflO acid oxidase and, 450, 477, 478, 482 Nuclear magnetic resonance eight iron-sulfur proteins, 43-44 flavodoxins, 96-98 rubredoxin, 12 superoxide dismutase, 547,551 Nutrition, xanthine dehydrogenase and, 410

0 %-Octane, monooxygenase and, 280 Octylamine, cytochrome P-450 and, 276 Octylmercaptane, cytochrome P-450 and, 276 Old Yellow Enzyme flavin dissociation constants, 500 molecular properties and kinetic mechanism, 471473

Optical rotatory dispersion blue oxidases, 566 xanthine oxidase, 341 Orcinol hydroxylase, properties, 223-224 Ornithine arnine oxidase content, 514 lysine monooxygenase and, 201 Oxalate oxidase, flavin and, 422 Oxenoid mechanism, dioxygenases, 184-185 Oxidase, definition of, 422 Oxipurinol, see Alloxanthine Oxygen adrenodoxin and, 267 aldehyde oxidase and, 409 amine oxidases and, 517, 518, 526 n-amino acid oxidase and, 452, 453, 454, 485 L-amino acid oxidase and, 430-431, 437,439,444445,457,459 apoferredoxin reconstitution and, 27 arginine monooxygenase and, 204 bacterial luciferase and, 228 blue oxidases and, 578-579 y-butyrobetaine hydroxylase and, 169, 170 cytochrome P-450 and, 279 activation, 283-284 dopamine p-monooxygenase and, 295 electron-transferring flavoprotein and, 113, 116 external flavoprotein monooxygenases and, 205, 206 flavodoxins and, 66, 105-107 flavoproteins and, 192 functions of, 253-254 glucose oxidase and, 438, 462 p-hydroxybenzoate hydroxylase and, 212, 214, 215 p-hydroxyphenylpyruvate hydroxylase and, 180, 181-182 imidazolylacetate monooxygenase and, 226 iron proteins and, 26 kynurenine-3-hydroxylaseand, 230 laccase and, 573, 577 lactate monooxygenase and, 195, 199 lysyl hydroxylase and, 166, 200 melilotate hydroxylase and, 217,218 microsomal amine oxidase and, 229

SUBJECT INDEX

molybdenum hydroxylases and, 395, 397, 398, 399-400 monoamine oxidase and, 469, 471 nitroalkane oxidation and, 493, 496 nitrogenase and, 50, 51, 52, 53 Old Yellow Enzyme and, 471-472 orcinol hydroxylase and, 223-224 oxidoreductases and, 422 oxygenases and, 120-121 phenol hydroxylase and, 223 phenol o-monooxygenase and, 254, 296, 297 phenylalanine hydroxylase and, 232, 238 prolyl hydroxylase and, 156, 158, 160, 164, 185 pyrimidine deoxyribonucleoside 2'hydroxylase and, 178 pyrocatechuate 3,4dioxygenase and, 137, 138 reduced flavin oxidation mechanism, 503-505 salicylate hydroxylase and, 206, 208, 209, 210 singlet, superoxide dismutase and, 535-537 solubility, 427 llp-steroid monooxygenase and, 267, 268 sulfite oxidase and, 417,418, 419 superoxide dismutase and, 555-556 tetrahydropteridines or dihydroflavins and, 193, 231,242,245-246,251 thymine 7-hydroxylase and, 174, 175, 179 tryptophan hydroxylase and, 241 tyrosinase and, 508-509 tyrosine hydroxylase and, 238 uracil 5-carboxylate and, 176 xanthine oxidase and, 353, 354,355358, 363, 364, 365, 368-369, 372, 374, 382-385, 386 Oxygen electrode(s), flavoprotein oxidases and, 426-428 Oxypurine dehydrogenase, specificity, 389

P Pargyline, monoamine oxidaee and, 470471, 498

Parsley ferredoxin antibodies and, 26 properties of, 16 Pea, superoxide dismutase of, 539, 541, 544, 547 Pea seedlings amine oxidase, 512 catalytic mechanism, 526 copper content, 519,520 inhibitors of, 523 purification, molecular weight and substrate specificity, 514 pyridoxal phosphate and, 522-523 spectral properties, 518-519 substrate binding site, 525 Penicilliurn amagasakiense, glucose oxidase of, 461, 463 Penicillium notatum, glucose oxidase of, 464-465 Peptidyllysine, amine oxidase and, 527 Peptococcus aerogenes ferredoxin, X-ray diffraction, 37-38 Peptostreptococcus elsdenii electron-transferring flavoprotein, 110 catalytic properties, 116-118 molecular properties, 111-116 ferredoxin of, 46 flavodoxin, 61 absorption spectra, 89, 90, 91 circular dichroism, 95 comproportionation, 102 dithionite and, 103-104 electron nuclear double resonance, 96 flavin and, 77, 83, 85, 88 flavin derivatives and, 84, 85-87 fluorescence, 93 nuclear magnetic resonance, 96-98 oxygen and, 105-107 reduction potential, 98,99, 100, 101, 102 regulation by iron, 64, 65 sequence of, 67,69 Perox id ase galactose oxidase and, 532 horseradish n-amino acid oxidase and, 453 glucose amay and, 428 tyrosine 3-monooxygenase and, 291

SUBJECT INDEX

Peroxydihydro intermediate, monooxygenases and, 243-244, 247 Persuccinate, dioxygenases and, 184 Persulfide group aldehyde oxidases and, 393 nitrate reductase cofactor and, 405 spinach ferredoxin and, 22 xanthine oxidase, 323-324, 325, 365, 381382 Petroleum, conversion t o animal food, 258 PH n-amino acid oxidase and, 450, 451 L-amino acid oxidase and, 460461, 483 apoferredoxin reconstitution and, 27 flavin reduction and, 483-484 flavodoxins and, 65,83,90,104 reduction potential, 98, 100-101 glucose oxidase and, 464465,466,483 laccase and, 573574 monoamine oxidase and, 470 sulfite oxidase and, 416-417, 418 superoxide dismutase and, 546, 547-548 xanthine oxidase and, 314,351,361362, 374, 382, 386 m-Phenanthroline, metapyrocatechase and, 141 l,l0-Phenanthroline amine oxidases and, 523, 527 y-butyrobetaine hydroxylase and, 168 cysteine oxygenase and, 149 2,3-dihydroxyindole 2B-dioxygenase and, 148 3-hydroxyanthranilate oxygenase and, 145 p-hydroxyphenylpyruvate hydroxylase and, 181 lipoxygenase and, 150 metapyrocatechase and, 141 monoamine oxidase and, 468 nonheme iron protein and, 289 phenylalanine hydroxylase and, 237 protocatechuate 3,4-dioxygenase and, 137 pyrocatechase and, 134 rubredoxin and, 12 steroid oxygenase and, 144 tryptophan hydroxylase and, 241

tyrosine hydroxylase and, 239 Phenazine methosulfate, xanthine oxidase and, 354, 363,364,367, 378 Phenethylamine, amine oxidases and, 514, 518 Phenethylhydrazine, monoamine oxidase and, 471 Phenobarbital, cytochrome P-450 and, 271, 274, 275, 277-278, 279, 283 Phenolase, oxygen and, 120 Phenol o-monooxygenase oxygen and, 254 properties of, 221-223, 296-297 Phenyl acetate, xanthine oxidase and, 320 Phenylalanine L-amino acid oxidase and, 457,459, 460,461, 477 oxidation, kinetics, 430-431, 444,449 substituted, D-amino acid oxidase and, 454 tyrosine hydroxylase and, 239-240 Phenylalahine hydroxylase, properties, 232-238, 287-289 Phenylalanine hydroxylase stimulating factor, effects, cofactors and, 234, 236-237, 289 Phenylalanine residues flavodoxins, 69, 79 superoxide dismutase and, 546 2-Phenylcyclopropylamine, amine oxidases and, 512 Phenylethylamine, dopamine p-monooxygenase and, 294 Phenylglycine, substituted, D-amino acid oxidase and, 454 Pheuylhydrazine, amine oxidases and, 471,519, 522, 523,527 N-Phenylimidazole, cytochrome P450,., and, 260-261 10-Phenylisoalloxazine, substrate oxidation by, 500-501 p-Phenyl-L-lactate, lactate oxidase and, 197 Phenylpyruvate D-amino acid oxidase and, 456 p-hydroxyphenylpyruvate hydroxylase and, 181 7-Phenyltetrahydropteridine,phenylalanine hydroxylase and, 235 Phosphate

SUBJECT INDEX

flavodoxin and, 77, 87 galactose oxidase and, 531 lactate monooxygenase and, 195-196 Phosphatidylcholine, xenobiotic monooxygenase and, 273 Phospholipid, monoamine oxidase and, 468 Phosphoroclastic reaction ferredoxins and, 34,45 flavodoxin and, 61,62,63 Phosphotransacetylase, flavodoxin and, 61 l-Phospho-2,8,9-trioxadamantane ozonide, singlet oxygen and, 536 Photobacterium fischeri, luciferase of, 227 Photobacterium leiognathi, superoxide dismutase of, 537-538 Photooxidation n-amino acid oxidase, 446 xanthine oxidase, 326 Photosynthesis, flavodoxins and, 61,62, 63 Photosystem I, ferredoxin and, 29-30 Phytoflavin, 58 Picohate, 3-hydroxyanthranilate oxygenase and, 145 4-Picolylamine, amine oxidase and, 518 Pig kidney amine oxidase, 512 active site, 524 catalytic mechanism, 526 copper content, 519,520 purification, molecular weight and substrate specificity, 516-517 pyridoxal phosphate in, 521-522 spectral propertiea, 518 Pig liver aldehyde oxidase, molecular properties, 390-391 monoamine oxidase of, 526-527 uricase of, 511 Pig plasma amine oxidase, 512 catalytic mechanism, 525-526 copper content, 519, 520 purification, molecular weight and substrate specificity, 517-518 pyridoxal phosphate in, 522 spectral properties, 518-519

A-1-Piperidine-2-carboxylate D-amino acid oxidase and, 484,489 lysine monooxygenase and, 200-201 Planteose, galactose oxidase and, 531 Plasma, monoamine oxidase of, 466 Pleated sheet, flavodoxins, 67,69, 71-72, 73 Polyglutamic acid, apoferredoxins and, 26-27 Poly (glycylalanylproline) , prolyl hydroxylases and, 164 Poly (glycylprolylalanine), prolyl hydroxylases and, 164 Poly (glycylprolylglycine) , prolyl hydroxylase and, 160 Polyporus circinatus, galactose oxidase of, 527-528 Polyporus versicolor, laccase of, 560 Poly (L-proline) , prolylhydroxylase and, 159, 160, 164 Poly (prolylglycylproline), prolyl hydroxylase and, 160 Potassium bromide, flavodoxins and, 65, 82 Potassium iodide, xanthine oxidase and, 316, 319 Potentiometric titrations, photosystem I, 30 Pregnenolone formation of, 268 118-steroid monooxygenase and, 267 Proline, n-amino acid oxidase and, 455, 481 Prolyl hydroxylase, 152-154 assays, 161 catalytic properties, 156-160 mechanism, 184-185 nonvertebrate, 163-165 purification and molecular properties, 154-156, 292-293 regulation, 161-163 Propanol ferredoxin and, 24 phenylalanine hydroxylase and, 2% Propionitrile, cytochrome P-450 and, 276 Propylamine amine oxidases and, 514, 517 lysine monooxygenase and, 202 Prostaglandin synthetaae, oxygenaees and, 126-127

SUBJECT INDEX

Proteolysis, xanthine oxidase preparation and, 305, 312, 320-322 Pro tocatechualdehyde metapyrocatechase and, 141 protocatechuate 3,4dioxygenase and, 137 Protocatechuate formation of, 225 metapyrocatechase and, 141 Protocatechuate 3,4-dioxygenase electron spin resonance properties, 139-140 molecular and catalytic properties, 136-137 spectral properties, 137-839 properties, 143 Protocollagen lysine 5-monooxygenase and, 293,294 proline 4-monooxygenase and, 292 Proton(s) molybdenum interactions, 334, 336-337 nitrogenaae and, 50, 51 Proton abstraction, flavoprotein oxidases and, 475,478-479,480,482,483484, 491492 Proton relaxation enhancement, amine oxidases, 520 Proton transfer, amine oxidases and, 524 Protoporphyrin IX, tryptophan 2,3-dioxygenase and, 128-129 Pseudomonas y-butyrobetaine hydroxylase of, 167-169 melilotate hydroxylase of, 217 tryptophan 2,3-dioxygenaae of, 128 Pseudomonas aeruginosa m-hydroxybenzoate-6-hydroxylase of, 224-225 protocatechuate 3,4dioxygenase of, 137 Pseudomonas arvilla rnetapyrocatechase of, 140 pyrocatechase of, 133-134, 135 Pseudomonas desmolytica, p-hydroxybenaoate hydroxylase of, 211, 214 Pseudomonas elsdenii, rubredoxin of, 13 Pseudomonas jluorescens homogentisate oxygenase of, 146 p-hydroxybenzoate hydroxylase of, 211-212, 214

641 lysine monooxygenase of, 199-200 Pseudomoms okovorans rubredoxin, alkanes and, 6 chemical properties, 12-13 Pseudomonas ovalis, 3,4dihydroxyphenylacetate 2,3-dioxygenase of, 142 Pseudomonas putidu camphor 5-ezo-monooxygenase of, 259-264 cytochrome P-450 of, 255 2,5-dihydroxypyridine oxygenase of, 146-147 p-hydroxybenzoate hydroxylase of, 211 iron protein, Mossbauer parameters, 10 4-met hoxy U-de methyl-monooxygenase of, 285-287 orcinol hydroxylase of, 223-224 salicylate hydroxylase of, 206, 207, 210 two iron-sulfur protein, amino acid sequence, 18 antibodies and, 26 properties, 16 Pseudomonas testosteroni m-hydroxybenzoate-4-hydroxylase of, 225 protocatechuate 4,5-dioxygenase of, 143 Pteridine(s) molybdenum hydroxylases and, 349, 358, 409 monooxygenases and, 122 Pulse radiolysis, superoxide dismutase and, 553 Purine, molybdenum hydroxylases and, 346, 350, 351 Purine-6-aldehyde, xanthine oxidase and, 363 Purine N-oxides, xanthine oxidase and, 354-355 Putidaredoxin, 2 camphor hydroxylation and, 264 properties, 263 Putrescine, arnine oxidases and, 514, 517, 521 Putrescine oxidase, sulfite and, 422 Pyridine, metapyrocatechase and, 141 Pyridine nucleotides external flavoprotein monooxygenases and, 204-205

642

SUBJECT INDEX

flavodoxin and, 61, 63 phenylalanine hydroxylase and, 232 Pyridoxal, copper-containing oxidases and, 509, 519 Pyridoxal oxidase, Drosophila, nature of, 408 Pyridoxal phosphate, amine oxidases and, 466, 521-523, 524, 527 Pyrimidines, dioxygenase reactions of, 169-179 Pyrimidine deoxyribonucleoside 2'hydroxylase catalytic properties, 176-178 properties of, 174 regulatiop of, 178-179 Pyrocatechase circular dichroism, 136 electron spin resonance properties, 135-136 isolation and properties, 120 molecular and catalytic properties, 133-135 spectral properties, 135 Pyrogallol metapyrocatechase and, 141 phenol hydroxylase and, 222 protocatechuate 3,4-dioxygenase and, 137 pyrocatechases and, 134 Pyruvate n-amino acid oxidase and, 479-480, 484-486 lactate monooxygenase and, 195, 196, 198 reduction, model systems and, 503 Pyruvate dehydrogenase flavodoxin and, 61, 65 properties of, 48 Pyruvate oxidase energy and, 423 flavin and, 422 Pyruvate synthase, flavodoxins and, 61

Q Quercitine dioxygenase, cofactor of, 125 Quinol, phenol hydroxylase and, 222 Quinolinate, 3-hydroxyanthranilate oxygenase and, 145 Quinoline, metapyrocatechase and, 141

Quinoline-8-thiol, molybdenum and, 333-335, 339 Quinone(s) phenol o-monooxygenase and, 296, 297 xanthine oxidase and, 353, 356

R Rabbit liver aldehyde oxidase, catalytic properties, 395, 396, 397, 398 molecular properties, 390-391, 393, 394 Raffinose, galactose oxidase and, 531 Raman spectroscopy, blue oxidases, 566-567 Rate constants, camphor 5-monooxygenase, 264 Rate equation, steady-state, flavoprotein oxidases, 429432 Redox-active groups xanthine oxidase magnetic and optical properties, 326-342 magnetic interactions and locations, 342-344 potentials, 379 Redox dyes, cysteamine oxygenase and, 148 Redox states, flavins, 424-425 Reduction potential cytochrome P-450, 282 eight iron-sulfur proteins, 44 flavodoxins, 65, 98-102 flavin derivatives and, 86 galactose oxidase and, 532533 4-methoxybenzoate O-demethyl-monooxygenase, 286 nitrogenase, 51, 52, 53 rubredoxins, 14 superoxide radical, 356 two iron-sulfur proteins, 19,23 Regreening factor, Old Yellow Enzyme and, 473 Resorcinol orcinol hydroxylase and, 223-224 phenol hydroxylase and, 221,222 Retinal, dioxygenase and, 126 Rhizobium japonicum, cytochrome P-450

643

SUBJECT INDEX

of, 280 Rhodospirillum gelatinosa, high potential iron protein, 32 R hodospirillum rubrum flavodoxin, 59, 66, 87 absorption maxima, 99 circular dichroism, 95 dithionite and, 104 fluorescence, 93-94 redox potentials, 99 Rhus uernicifera, laccase of, 560 Ribityl phosphate, flavodoxin and, 75 Ribityl side chain, flavodoxins and, 77, 79, 86-87 Riboflavin derivatives, binding by flavodoxins, 84 flavodoxin and, 77, 87 Ribonuclease phosphate environment, 77 superoxide and, 534 Rosy mutant chromosome locus, 407 electrophoretic variants, 410-411 Rubredoxin (8) characteristics of, 2-3 chemical properties, 1 2 1 5 historical background, 4-6 iron deficiency and, 65 physical properties, 6-12 superoxide and, 533

s Salicylaldehyde, molybdenum hydroxylases and, 346, 375, 376 Salicylamide, salicylate hydroxylase and, 207 Salicylate, xanthine oxidase and, 305, 308, 319, 323, 362 Salicylate hydroxylase catalytic pathway, 208 properties, 2W-211 Samarium, flavodoxin structure and, 70-71, 79 Santalum album, prolyl hydroxylase of, 164 Sarcosine dehydrogenase, flavopsotein and, 109 Scenedesmw, ferredoxin, amino acid sequence, 17

Schiffbase, amine oxidases and, 525 Selenide desulfo xanthine oxidase and, 323 putidaredoxin and, 19,20, 22-23, 263 Semicarbazide, amine oxidases and, 512, 522, 523, 527 Semiquinones flavodoxins and, 65,66, 71, 79-81, 83, 90, 95L96, 98, 100, 101 flavin derivatives, 86 oxygen and, 105-107 reduction of, 103 Serine residues, flavodoxins, 69, 73, 76, 78 Serotonin amine oxidases and, 514, 516,518 dioxygenases and, 123, 132 Shethna flavoprotein, 59 Skin, xenobiotic monooxygenases in, 270 Small intestine, xenobiotic monooxygenases in, 269-270 Solvent, flavodoxins and, 72-73, 76, 77 Spermidine, amine oxidases and, 513, 516, 527 Spermidine oxidase, 466 Spermine, amine oxidase and, 516, 527 Spermine oxidase, discovery of, 514-515 Spinach ferredoxin amino acid sequence, 17 antibodies and, 26 chemical modification, 28-29 EPR perturbants and, 23 exopeptidases and, 28 properties, 15, 16 118-steroid monooxygenase and, 268 iron protein, Massbauer parameters, 10 Spinach leaves, superoxide dismutase of, 539, 541, 544, 547 Spirochaeta aurantia, high potential iron protein, 32 Spirohydantoin, monooxygenases and, 244-245 Squash, ascorbate oxidase of, 560 Stachyose, galactose oxidase and, 531 Staphylococcal nuclease, phosphate environment, 77 Steapsin, microsomal cytochromes and, 272 Stellacyanin, copper of, 566, 567

SUBJECT INDEX

Steroid(s), hydroxylation of, 254, 255, 257, 258 Steroid oxygenase, properties, 143-144 118-Steroid monooxygenase, properties, 265-268,282283 Steroid 2l-monooxygenase, cytochrome P-450 and, 282283 Streptomyces antibioticus, prolyl hydroxylase of, 164-166 Streptomyces glaucescens, phenol omonooxygenase of, 297 Streptomyces griseus, arginine monooxygenase of, 203-204 Subunits blue oxidases, 562 p-hydroxyphenylpyruvate hydroxylase, 180 prolyl hydroxylase, 155-156 superoxide dismutase, 542-543 xanthine oxidase, 314-317 Succinate y-butyrobetaine hydroxylase and, 169, 170 5bormyluracil oxidation and, 176 a-ketoglutarate dioxygenases and, 184 lysyl hydroxylase and, It% prolyl hydroxylase and, 160 pyrimidine deoxyribonucleoside 2’hydroxylase and, 178 thymine 7-hydroxylase and, 175 Succinate-ubiquinone reductase, properties of, 49 Sulfate, reduction, flavodoxin and, 61 Sulfhydryl groups D-amino acid oxidase, 446 protocatechuate 3,4-dioxygenase and, 139 superoxide dismutase, 542-543,549 Sulfhydryl reagents, lysine monooxygenase and, 202 Sulfide apoferredoxin reconstitution and, n cysteamine oxygenase and, 148, 149 desulfo xanthine oxidase and, 323,324 flavodoxin and, 61 superoxide dismutase and, 551 Sulfite flavins and, 501, 502 flavodoxins and, 66 flavoprotein oxidases and, 422

Sulfite oxidases liver catalytic properties, 417-419 molecular properties, 414-417 Sulfite reductase, properties of, 48 Sulfur acid-labile, xanthine oxidase and, 310, 324 molybdenum ligands and, 333, 339 oxidation of, 125 Sulfur dioxide, toxicity, sulfite oxidase and, 414 Superoxide dismutase assay methods, 539-540 catalytic mechanism, 552667 crystallization, 542 determination of purity and concentration, 540 flavodoxin oxidation and, 105-106, 108 functionof, 510, 533-537 galactose oxidase and, 530, 532 historical, 533 m-hydroxybenzoate-4-hydroxylase and, 225 modification of, 549-551 molecular properties apoenzyme, 548-549 native enzyme, 542-548 reconstitution, 549651 redox properties, 551-552 monooxygenases and, 247 optical properties, 541, 545-548 apoenzyme, 549 prokaryotic and mitochondrial, 537-538 purification, 538-539 xanthine oxidase and, 357, 359, 366, 384 Superoxide free radicals, xanthine oxidase and, 302-303,354, 355-358,359, 366,367,373,384,385 Superoxide ion dioxygenases and, 186 electron paramagnetic resonance spectrum, 327 galactose oxidase and, 530, 532 oxidoreductases and, 422 tryptophan 2,3-dioxygenase and, 129, 131 Synechococcus lividus

645

SUBJECT INDEX

ferredoxin of, 26 flavodoxin of, 59 absorption maxima, 99 electron paramagnetic resonance, 96 nuclear magnetic resonance, 96-97 photosynthesis and, 109 redox potentials, 99

T n-Talose, galactose oxidase and, 531 Taro, ferredoxin, amino acid sequence, 17 Temperature, apoferredoxin reconstitution and, 27 Tetradecane, monooxygenase and, 280 Tetraethylthiuram disulfide, xanthine oxidase and, 366 Tetrahydrobiopterin phenylalanine hydroxylase and, 232, 236 tryptophan hydroxylase and, 241 tyrosine hydroxylase and, 239, 240 Tetrahydrofolate y-butyrobetaine hydroxylase and, 168 p-hydroxyphenylpyruvate hydroxylase and, 180 Tetrahydropteridines monooxygenases and, 192-193, 231 phenylalanine hydroxylase and, 238, 288,289 prolyl hydroxylase and, 157 tryptophan hydroxylase and, 241,292 tyrosine hydroxylase and, 238, 291 Tetranitroformate, spinach ferredoxin and, 28 Tetranitromethane D-aminO acid oxidase and, 452 flavodoxins and, 88 spinach ferredoxin and, 28-29 superoxide dismutase and, 552,555 xanthine oxidase and, 357 Tetrazolium reductase inhibitor, xanthine oxidase and, 366 Tetrazolium salts, xanthine oxidase and, 359 Thermolysin, flavodoxins and, 67 p-2-Thienylalanine, phenylalanine hydroxylase and, 234 Thiobacillus thiooxiduns, dioxygenase of, 125

Thiocapsa pfennigii, high potential iron protein, 32 Thiocyanate galactose oxidase and, 530 superoxide dismutase and, 548 xanthine oxidase and, 319,323,339, 362 Thiols, p-hydroxybenzoate hydroxylase and, 211 Thiol groups phenylalanine hydroxylase, 236 superoxide and, 534 xanthine oxidase, 311412, 320-322, 354, 365, 366 Thiophenol, oxidation, flavin and, 501 6-Thioxanthine, molybdenum hydroxylases and, 347 Threonine, spinach ferredoxin and, 28 Threonine residues, flavodoxins, 76 Thymidine, salvage of, 169-172 Thymidylate, pyrimidine deoxyribonucleoside 2’-hydroxylase and, 177 Thymine 7-hydroxylase catalytic properties, 174-176 purification and molecular properties, 172-174 regulation of, 178-179 Tiron ferredoxins and, 26 tryptophan hydroxylase and, 241 xanthine oxidase and, 366 Transferrin, 2 Trichloroacetic acid ferredoxins and, 26 flavodoxin and, 82 rubredoxin and, 12 Trichosporon cutaneum, phenol hydroxylase of, 221-223 4-Trifluoromethylbenzoate, 4-methoxybenzoate O-demethyl-monooxygenase and, 287 Trifluoroperacetic acid, cytochrome P450 and, 284 Trihydroxybenzene(s), phenol hydroxylase and, B 2 2,3,4-Trihydroxybenzoate, formation of, 214, 216 3-Trimethylaminopropionate, y-butyrobetaine hydroxylase and, 168 3-Trimethylaminopropylsulfonate,y-

646 butyrobetaine and, 169 Triton X-100, monoamine oxidase and, 467, 468 Trypsin cytochrome P-450 and, 273 flavodoxins and, 67 p-hydroxyphenylpyruvate hydroxylase and, 180 tyrosine hydroxylase and, 239, 291 Tryptamine amine oxidases and, 514, 518 indoleamine 2,3-dioxygenase and, 132 Tryptophan dioxygenases and, 123 phenylalanine hydroxylase and, 234, 235 L-Tryptophan 2,Bdioxygenase catalytic properties, 129-130 historical, 127-128 mechanism, 186 molecular properties, 128-129 Tryptophan hydroxylase, properties, 240-241, 291-292 n-Tryptophan pyrrolase, see Indoleamine 2,3-dioxygenase Tryptophan pyrrolase, see Tryptophan dioxygenase Tryptophan residues flavin binding and, 447 flavodoxins, 60,73,77, 78,82,97, 108 distribution of, 79 flavin and, 88 fluorescence and, 94 modifications of, 78, 88 oxidation state and, 80, 101-102 salicylate hydroxylase, 207-208 superoxide and, 534 superoxide dismutase content, 543, 646 Tumors, tryptophan 5-monooxygenase in, 291 Tungsten liver xanthine oxidase and, 309 sulfite oxidase and, 415 Turkey liver xanthine dehydrogenase activation energy, 361 catalytic properties, 395, 396397, 398, 399 dismutation reaction, 360-361

SUBJECT INDEX

molecular properties, 390-391, 393, 394 substrate specificity, 345-352 Tyramine amine oxidases and, 514, 516, 518 cheese and, 466 dopamine p-monooxygenase and, 294 Tyrosinase copper and, 508 substrates of, 508-509 Tyrosine, phenylalanine 4-monooxygenase and, 289 m-Tyrosine phenylalanine hydroxylase and, 235-236 tyrosine hydroxylase and, 240 Tyrosine hydroxylase, properties, 238240, 290-291 Tyrosine residues n-amino acid oxidase, 446 bacterial ferredoxin, 39, 45-46 flavin binding and, 447 flavodoxins, 78,82,97, 101-102 distribution of, 79 flavin and, 88 fluorescence and, 94. modification of, 78 metapyrocatechase, 142 rubredoxin, 12 spinach feriedoxin, 28-29 superoxide dismutase, 543, 546

U Ubiquinone-cytochrome c reductase, properties of, 49 Uracil-5-carboxylate, formation of, 173, 187 Urea amine oxidase and, 522 n-amino acid oxidase and, 448 cytochrome P-450,,, and, 262 galactose oxidase and, 528 3-hydroxyanthranilate oxygenase and, 145 metapyrocatechase and, 141 nitrogenase and, 52 two iron-sulfur proteins and, 23, 26 xanthine oxidase and, 314,326,362 Uric acid, xanthine oxidase and, 333,

647

SUBJECT INDEX

350, 354, 355, 359, 36CL361, 367 Uricase, copper and, 509, 511

V Valine, n-amino acid oxidase and, 448, 450, 454-455 Valine residues, flavodoxins, 69 Vanadous ions, rubredoxin and, 14, 15 Veillonella alcalescens xanthine oxidase of, 336,343,350 catalytic properties, 395, 396, 398, 399 molecular properties, 390-391, 393, 394 reversibility, 360 Vibrio tyrosinaticus, phenol o-monooxygenase of, 297 Vitamin A, dioxygenase and, I23

W Water, flavodoxins and, 72-73, 76,90,98 Wheat germs, superoxide dismutase of, 539, 541, 544 Wilson’s disease, ceruloplasmin and, 559

utilization as nitrogen source, 301-302 Xanthine dehydrogenase conversion to oxidase, 320 molecular properties, 390-391 nitrate reductase and, 405-406 Xanthine oxidase, see ako Milk xanthine oxidase genetic deficiency of, 302 indoleamine 2,3-dioxygenase and, 131 properties of, 48, 56 superoxide and, 534,539,553 Xanthinuria, human, molybdenum hydroxylase genetics and, 400-402 Xenobiotic monooxygenase systems, flavoproteins and, 269-270 X-Ray crystallography flavodoxins, 70,71, 102 superoxide dismutase, 557 Xylose, glucose oxidase and, 436,444, 462, 477 a-Xylyl-a,a’-dithiol, iron-sulfur protein model compound and, 46

Y Yeast, superoxide dismutase of, 539,541, 544, 547

X

Z Xanthine dismutation of, 360 molybdenum hydroxylases and, 346, 350,353, 359, 361-362,368-369, 372, 374-382, 386, 387, 395, 396

Zinc, superoxide dismutase and, 533, 537, 548, 549, 550 Zymobacterium oroticum, dihydroorotate dehydrogenase of, 48

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