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


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


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


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


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


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


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


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


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


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


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


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


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


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 4 4 6 12 15 15 20 23 25 31 31 34 35 37 37 39 44 46 47 50 56 56

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 ( 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

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.

FLAVODOXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

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).

2.


61

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.

62


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).

2.


63

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).

64


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).

2.

FLAVODOXINS AND ELECTRON-TRANSFERRING

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).

66


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).

2.

FLAVODOXINS A N D ELECTRON-TRANSFERRING FLAVOPROTEINS

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.


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


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.


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


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


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.


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.

2.


77

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.

2.


79

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


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.

2.

FLAVOWXINS AND ELECTRON-TRANSFERRING FLAVOPROTEINS

81

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.

2.


FLAVOPROTEINS

83

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' -(

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


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.


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


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


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


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


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


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


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


,

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


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.


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


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.


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 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.


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

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.


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.


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.


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).

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(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).

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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.

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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.

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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


V. akalescens (17 6 )


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

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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).

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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).

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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).

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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).

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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).

404

R. C. BRAY

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).

6.


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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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407

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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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.

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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),

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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

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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.


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.


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


(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] 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


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


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.


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


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


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


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


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


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.


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.


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


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


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


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.


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


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


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


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


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.


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


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


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


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.


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


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.


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


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.


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


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.


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.

474


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


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-

-

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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-

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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

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--

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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


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 , [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


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


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


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.

-

- -

-

-

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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


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

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-

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


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


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


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


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


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


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


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


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


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


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


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.


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


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. 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


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.


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


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


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


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


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).


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.


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.


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.


515

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.


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.


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.


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.


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.


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

529

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.


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.


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.

COPPER-CONTAINING

OXIDASES AND SUPEROXIDE DISMUTASE

535

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.


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.


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.


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


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.


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.


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.


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

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