Flavins and Flavoproteins 1990: Proceedings of the Tenth International Symposium, Como, Italy, July 15–20, 1990 9783110855425, 9783110123739

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Flavins and Flavoproteins 1990

Flavins and Flavoproteins 1990 Proceedings of the Tenth International Symposium Como, Italy, July 15 - 20,1990 Editors B. Curti • S. Ronchi • G. Zanetti

W DE

G Walter de Gruyter • Berlin • New York 1991

Editors Bruno Curti, M.D., Ph.D. • Professor of Biochemistry Dipartimento di Fisiologia e Biochimica Generali Severino Ronchi, Ph.D. • Professor of Biochemistry Istituto di Fisiologia Veterinaria e Biochimica Giuliana Zanetti, Ph.D. • Professor of Biochemistry Dipartimento di Fisiologia e Biochimica Generali Università Statale di Milano Via Celoria 26 1-20133 Milano Italia © Printed on acid free paper which falls within the guidelines of the ANSI to ensure permanence and durability. Library of Congress Cataloging-in-Publication Data International Symposium on Flavins and Flavoproteins (10th : 1990 ; Como, Italy) Flavins and flavoproteins : proceedings of the Tenth International Symposium on Flavins and Flavoproteins, Como, Italy, July 15-20.1990 / editors: B. Curti, S. Ronchi, G. Zanetti, p. cm. Includes bibliographical references and index. ISBN 0-89925-666-X 1. Flavoproteins--Congresses. 2. Flavins-Congresses. I. Curti, B. (Bruno). 1931- . II. Ronchi, S. (Severino), 1935- . III. Zanetti, G. (Giuliana), 1941- . IV. Title. QP552.F54I57 1990 91-128 574.19'258-dc20 CIP CIP-Kurztitelaufnahme der Deutschen Bibliothek Flavins and flavoproteins . . . : proceedings of the . . . international symposium . . . - Berlin ; New York : de Gruyter. 1990. Proceedings of the tenth international symposium, Como, Italy, July 15-20,1990. -1991 ISBN 3-11-012373-8 Cover illustration by courtesy of SCIENCE Polypeptide chain folding and domain structure of ferredoxin-NADP + reductase. Stylized Cabackbone of the whole protein: the FAD domain (purple) and the NADP+ domain (green) are shown with models and van der Waals surfaces for FAD (yellow) and 2'-phospho-AMP (blue). The cleft where ferredoxin may bind faces the viewer. P. A. Karplus et al. in: SCIENCE, Vol. 251 (1991), @ A A A S Fig. 2,60 Copyright © 1991 by Walter de Gruyter& Co., D-1000 Berlin 30. All rights reserved, including those of translation into foreign languages. No part of this book may be reproduced in any form - by photoprint, microfilm or any other means - nor transmitted nor translated into a machine language without written permission from the publisher. Printing: Gerike GmbH, Berlin. Binding: Heinz Stein, Berlin. - Printed in Germany.

PREFACE

This volume is a collection of the proceedings of the Tenth International Symposium on Flavins and Flavoproteins, held in Como at the Villa Olmo, from the 15th to the 20th of July 1990. Since the first meeting held in Amsterdam in 1965, these Symposia have aimed at focusing on the latest developments in the area of flavin and flavoprotein research and at providing a discussion forum for the scientists working in this field. The success of these meetings is certainly due, first of all, to the interaction of various disciplines which has been slowly reinforced over 25 years: protein chemists, kineticists, crystallographers and organic chemists have since then started to speak a common language about flavins. Secondly, as pointed out by Vince Massey and Charles Williams in their Preface to the proceedings of the Ann Arbor Symposium in 1981, "...This mixture of disciplines, meeting in a relaxed atmosphere where everyone can listen and participate, makes these Symposia of great value, and has been instrumental in promoting the rapid advances in the field". Judging from the lively discussions among the 210 participants (with a large proportion of young scientists) which went on late into the summer evenings, we believe that the Como Symposium followed a similar pattern. This volume gives a clear indication of the advances made in flavin research since the last Symposium. To mention just one of the recent achievements in the field, seven new threedimensional structures of flavoproteins were presented at the Symposium and many of them were complemented by an impressive wealth of data obtained from the most advanced chemicophysical and molecular biology techniques. At the Symposium

VI

there was also evidence that protein engineering is now routinely employed in flavoprotein research in addition to flavin analogs as a means of elucidating the way each apoprotein modulates the chemistry of its flavin coenzyme. The Symposium witnessed also the great progress which has been made in the field of flavin chemistry and in the understanding at molecular level of the action mechanism of single flavoproteins; but, in our opinion, the major advances have been made in the elucidation of similarities and differences among the various types of flavoproteins. This mass of new data made it more difficult to arrange the Table of Contents, due to the fact that although it gave a clear definition of the action mechanism of a great number of flavoproteins, it also weakened any strict border lines among their classes. We are entirely responsible for the choices made and we only hope that these choices will not hinder the consultation of the volume but rather, that reading these proceedings will stimulate new ideas and further progress in the field of flavin research. We would like to thank the members of the Organizing Committee, the Chairpersons of the Lecture and Discussion Sessions and all those who contributed in various ways to the organization of this Symposium: in particular, the University of Milano and its Rector, Professor Paolo Mantegazza, the International Union of Biochemistry, the Italian Biochemistry Society, the Italian Research Council, the pharmaceutical and equipment companies whose names are listed at the front of the volume. Thanks also to all those colleagues and students in our laboratories whose enthusiastic cooperation brought about the realization of this Symposium. We owe a considerable debt to our colleague Maria Antonietta Vanoni for her help in making this volume a reality. A special word of gratitude to the members of the Secretariat at the University and at Villa Olmo, who did a splendid job in assisting the people attending

VII

the conference. Our thanks also go to Ms Evelyne Glowka of de Gruyter Publishers for editorial assistance. We are sure that in the coming years flavin and

flavoprotein

research will continue to grow and we hope that the results of the

Como

Symposium will

appointment

will

on

and

Flavins

be part

be at the

of

"Eleventh

Flavoproteins"

such

a growth.

International

in Japan,

in

1993,

Our

next

Symposium where

host will be Professor Kunio Yagi. Arrivederci! Bruno Curti Severino Ronchi Giuliana Zanetti Milano, October, 1990

our

ORGANIZING COMMITTEE B. Cuiti, S. Ronchi and G. Zanetti (Milano) Co-chairpersons D.E. Edmondson (Atlanta), P. Engel (Sheffield), S. Ghisla (Konstanz), F. Lederer (Paris), V. Massey (Ann Arbor), S. Mayhew (Dublin), D.B. McCormick (Atlanta), Y. Miyake (Osaka), G. Schulz (Freiburg), C. Veeger (Wageningen), C. Walsh (Cambridge, U.S.A.), C.H. Williams (Ann Arbor), K. Yagi (Mitake) LOCAL COMMITTEE A. Negri, M.S. Pilone, M.A. Vanoni

ACKNOWLEDGEMENTS The Organizing Committee would like to express its gratitude to the following for their valuable help and support in making this symposium possible: Università' degli Studi di Milano International Union of Biochemistry Consiglio Nazionale delle Ricerche Società' Italiana di Biochimica EniChem Agricoltura - Agrimont (ENIMONT) E. I. du Pont de Nemours & Company Kontron Instruments Millipore - Waters Istituto G. Donegani Beckman Analytical S.p.A. Heraeus Pharmacia Amicon Grace Italiana Applied Biosystems Merck Sharp & Dome Serono Eneo Lepetit

CONTENTS

INTRODUCTORY LECTURE Flavins and flavoproteins - past, present, and future K. Yagi FLAVIN CHEMISTRY AND PHYSICO-CHEMICAL STUDIES OF FLAVOPROTEINS Spectral properties of cyanoalloxazines J. Koziol, M. M. Szafran, A. Koziolowa, and H. Szymusiak

19

Phototautomerism of alloxazines in the presence of acetic acid in different media A. Koziolowa, M. Stroinska, and M. M. Szafran

23

Ionisation properties of reduced, 1,5-dihydroflavin, rates of N(5)-H exchange with solvent. S. Ghisla, P. Macheroux, C. Sanner, H. Riiteijans, and F. Muller

27

Photoproduct of 8-hydroxyflavin K. Matsui, S. Kasai, R. Miura, and S. Fujii

33

The breakdown of C(4a)-flavin-peroxides into flavin and a hydroperoxide in different environments G. Mer6nyi, and J. Lind

37

Fluorescence spectra of crystalline alloxazine and its methyl derivatives M. M. Szafran, J. Koziol, and P. F. Heelis

41

Comparative fluorescence study of phototautomerism of lumichrome in dodecylammonium propionate reversed micelles B. Tyrakowska, A. Koziolowa, P. I. H. Bastiaens, and A. J. W. G. Visser

45

Picosecond dynamics reduced flavins A. J. W. G. fluorescence Visser, S. Ghisla, and J.ofLee

49

Reaction mechanism of flavin-photosensitized oxidation and reduction of c-type, cytochromes M. Roncel, M. Hervas, J. A. Navarro, M. A. De la Rosa, and G. Tollin

55

A simple method for the determination of redox potentials V. Massey

59

Electron spin resonance spectral studies of 15 N

- 2 H flavin coenzyme semiquinones in flavoenzymes D. E. Edmondson, F. Muller, F. Schaub, and Y. Nisimoto

67

X Interflavin energy transfer as a tool to determine geometrical parameters in flavoproteins P. I. H. Bastiaens, and A. J. W. G. Visser

73

BIOSYNTHESIS OF FLAVINS AND FLAVOPROTEINS Cloning and molecular characterization of the riboflavin synthase-encoding gene of Saccharomyces cerevisiae J. L. Revuelta, M. A. Santos, and J. J. Garcia-Ramirez

81

FAD synthetase from Brevibacterium ammoniagenes:progress report on cloning and expression in Escherichia coli P. A. Ellison, and E. F. Pai

85

Rat brain flavokinase: purification, properties, and comparison to the enzyme from liver and small intestine H. Nakano, and D. B. McCormick

89

Flavin-dependent expression and modification of 6-hydroxy-D-nicotine oxidase R. Brandsch

93

Covalent flavinylation of apo-6-hydroxy-D-nicotine oxidase K. Decker, H. Nagursky, V. Bichler, L. Mauch, and R. Brandsch

101

OXIDASES The structure of glycolate oxidase Y. Lindqvist

107

Two-photon reactions of dihydroflavin mononucleotide in glycolate oxidase, studied by nanosecond laser photolysis L. Lindqvist

115

Characterization glycolate and active site mutant P. Macheroux, V.ofMassey, D. oxidase J. Thiele, E.an Söderlind, and Y. Lindqvist

119

L-lactate monooxygenase, proposed active site structure and mechanism S. Ghisla, and V. Massey

123

Lactate oxidase: mutagenesis and expression of the mycobacterial gene U. Müh, D. A. Giegel, V. Massey, and C. H. Williams,Jr.

131

Molecular biological studies on structure-function relationship of D-amino acid oxidase Y. Miyake, K. Fukui, K. Momoi, F. Watanabe, M. Tada, M. Miyano, and S. Takahashi

135

Genomic organization and expression of D-amino acid oxidase gene K. Fukui, K. Momoi, F. Watanabe, M. Tada, M. Miyano, and Y. Miyake

143

XI

Expression of mouse kidney D-amino acid oxidase in Escherichia coli: purification and characterization of the recombinant protein M. Tada, K. Fukui, M. Miyano, and Y. Miyake

147

Purification of recombinant mutant porcine D-amino acid oxidases M. Miyano, K. Fukui, F. Watanabe, M. Tada, S. Takahashi, and Y. Miyake

151

Limited proteolysis studies on the apo-, holo-, holoenzyme-benzoate forms of pig kidney D-amino acid oxidase G. Torn Tarelli, M. A. Vanoni, A. Negri, and B. Curti

155

Reinvestigation of the role of lysine residues in D-amino acid oxidase T. Simonic, L. Mannucci, A. Negri, G. Tedeschi, and S. Ronchi

159

Structural change of a ligand upon complexation with D-amino acid oxidase Y. Nishina, K. Sato, and K. Shiga

163

D-amino acid oxidase expressed under induction conditions is enzymatically active in microperoxisomes of Rhodotorula gracilis M. Pilone Simonetta, M. E. Perotti, and L. Pollegioni

167

On the recombination process of apo-D-amino acid oxidase from Rhodotorula gracilis P. Casalin, L. Pollegioni, B. Curti, and M. Pilone Simonetta

171

FAD analogues as active site probes of Rhodotorula gracilis D-amino acid oxidase L. Pollegioni, M. Pilone Simonetta, and S. Ghisla

175

Structural studies on beef kidney D-aspartate oxidase A. Negri, G. Tedeschi, F. Ceciliani, T. Simonic, and S. Ronchi

179

Modification of substrate specificity of D-aspartate oxidase chemically modified by phenylglyoxal G. Tedeschi, A. Negri, P. A. Biondi, C. Secchi, and S. Ronchi

189

Kinetic isotope effects on flavoprotein oxidases P. F. Fitzpatrick, C.-T. Hsieh, J. M. Denu, and J. J. Emanuele

193

Structural and kinetic analysis of flavine adenine dinucleotide modification in alcohol oxidase from methylotrophic yeasts L. V. Bystrykh, R. M. Kellogg, W. Kruizinga, L. Dijkhuizen, W. Harder, J. Vervoort, and W. J. H. van Berkel

197

Studies on yeast-derived and mutant glucose oxidases S. Chakraborty, V. Massey, J. Stratton-Thomas, and S. Rosenberg

201

Effect of deglycosylation on the activity and stability of glucose oxidase from Aspergillus niger H. M. Kalisz, R. D. Schmid, H. J. Hecht, and D. Schomburg

205

and -NMR investigations of glucose oxidase from Aspergillus niger C. Sanner, P. Macheroux, H. Riiteijans, F. Miiller, and A. Bacher

209

XII

Effect of hydrogen peroxide on baker's yeast pyridoxaminephosphate oxidase under aerobic and anaerobic conditions H. Tsuge, K. Bando, S. Uchida, K. Shinohara-Asai, and Y. Hattori

213

MONOOXYGENASES Chemical functions of amino acid residues in the active site of para-hydroxybenzoate hydroxylase as studied by site directed mutagenesis and biophysical techniques B. Entsch, B. A. Palfey, D. P. Ballou, and V. Massey

219

Site-directed mutagenesis of para-hydroxybenzoate hydroxylase from Pseudomonas fluorescens A. H. Westphal, K. Eschrich, W. M. A. M. van Dongen, J. A. E. Benen, A. de Kok, and W. J. H. van Berkel

231

Partial sequences of phenol hydroxylase and similarities with other flavoenzymes H. Y. Neujahr, and T. Sejlitz

235

2-aminobenzoyl-CoA monooxygenase/reductase, a novel type of flavoprotein hydroxylase B. Langkau, S. Ghisla, V. Massey, and G. Fuchs

239

Structural studies on the porcine liver multisubstrate flavin-containing monooxygenase K. K. Korsmeyer, L. L. Poulsen, and D. M. Ziegler

243

The cofactor dependent interaction of molecular oxygen with phenylalanine hydroxylase S. W. Bailey, J. P. Crow, and J. E. Ayling

247

FLAVIN DEPENDENT BIOLUMINESCENCE Mechanisms of bacterial luciferase and aromatic hydroxylases S.-C. Tu, H. I. X. Mager, R. Shao, K. W. Cho, and L. Xi

253

Random and site directed mutagenesis of bacterial luciferase L. J. Chlumsky, L. H. Chen, C. Claik, H. Abu-Soud, M. M. Ziegler, F. M. Raushel, and T. O. Baldwin

261

Xenorhabdus luminescens luciferase: cloning, sequencing, and overexpression of the encoding genes and substrate inhibition of the enzyme L. Xi, K. W. Cho, and S.-C. Tu

265

On the mechanism of bacterial luciferase. 4a,5-dihydroflavins as model compounds for reaction intermediates J. W. Eckstein, and S. Ghisla

269

XIII

Kinetic and mechanistic investigation of the bacterial luciferase reaction F. M. Raushel, H. M. Abu-Soud, L. S. Mullins, W. A. Francisco, and T. O. Baldwin

273

Electron (charge) transfers in flavin-mediated luminescence H. I. X. Mager, and S.-C. Tu

277

On the mechanism of dithionite/HoOo-induced bacterial bioluminescence S.-C. Tu, and K. W. Cho

281

Structure of FP39Q including its prosthetic group (Q-flavin): physiological significance of light emitting reaction in luminous bacteria S. Kasai, S. Fujii, R. Miura, S. Odani, T. Nakaya, and K. Matsui

285

DEHYDROGENASES AND ELECTRON TRANSFERASES

Crystallographic studies of medium chain acyl-CoA dehydrogenase from pig liver mitochondria J. J. P. Kim

291

Modulation C. Thorpe of flavin reactivity in the acyl-CoA dehydrogenases

299

Deuterium solvent isotope effects on the dissociation of the charge transfer product complex of medium chain acyl-CoA dehydrogenase and FPCoA L. Niu, J. T. McFarland, and B. A. Feinberg

307

Aromatic substrate analogues as mechanistic probes for medium-chain acyl-CoA dehydrogenases S. Engst, and S. Ghisla

311

Inactivation of medium-chain acyl-CoA dehydrogenase from pig kidney by methylenecyclopropyl-acetyl-CoA: identification of a new type of flavin-inhibitor adduct H.-D. Zeller, and S. Ghisla

315

Some properties of Glu-376-Gln active site mutant of human medium-chain acyl-CoA dehydrogenase S. Engst, P. Brass, J. Stiemke, A. Schieber, A. W. Strauss, D. P. Kelly, I. Rasched, and S. Ghisla

319

On the role of Glu-376 in catalysis of acyl-CoA dehydrogenases K. Ankele, K. Melde, S. Engst, P. Brass, S. Ghisla, and A. W. Strauss

325

Rat liver medium-chain acyl CoA dehydrogenases directed by complementary DNAs differing in their 5'-region T. Inagaki, N. Ohishi, K. Yagi, N. Tsukagoshi, S. Udaka, and S. Ghisla

329

XIV

Purification and some properties of five distinct acyl-CoA dehydrogenases from bovine liver K. Melde, and S. Ghisla The involvement of a green FAD-containing enzyme in the dehydration of 5-hydroxyvaleryl-CoA to 4-pentenoyl-CoA in Clostridium aminovalericum U. Eikmanns, and W. Buckel Is glutaryl-CoA dehydrogenase from Paracoccus denitrificans regulated by substrate and product binding? C. M. Byron, M. T. Stankovich, and M. Husain Two flavo-dehydiogenases catalyzing desaturation of A-ring of 34cetosteroids: 3-ketosteroid-A -dehydrogenase and 3-ketosteroid¿ 4 -dehydrogenase from Nocardia corallina E. Itagaki, T. Hatta, and H. Matsushita The cloning of old yellow enzyme K. Saito, M. Davio, O. Lockridge, and V. Massey Crystallization and characterization of old yellow enzyme K. M. Fox, S. M. Jacques, and P. A. Karplus Studies on old yellow enzyme reconstituted with the active site probe, 8-fluoro-8-demethyl FMN. Covalent modification and 19F-NMR R. Miura, S. Fujii, K. Kuroda, and S. Kasai Two-dimensional NMR studies of Desulfovibrio vulgaris and Megasphaera elsdenii flavodoxin J. Vervoort, C. P. M. van Mierlo, and J. LeGall l^C-NMR studies on cyanylated Megasphaera elsdenii flavodoxin G. M. Doherty, R. Motherway, S. G. Mayhew, and J. P. G. Malthouse Flavin binding site geometry in Anabaena 7120 flavodoxin. Progress in determining the flavodoxin solution structure B. J. Stockman, A. M. Krezel, J. B. Olson, E. S. Mooberry, and J. L. Maikley NMR study of NADPH-adrenodoxin reductase reconstituted with C-enrichedFAD S. Fujii, Y. Nonaka, M. Okamoto, and R. Miura 13

Covalent phosphorus groups in flavoproteins: the phosphodiester linkage in Azotobacter (OP) flavodoxin D. E. Edmondson, M. Boylan, and M. Taylor Differences in flavin motional dynamics in oxidized and reduced clostridial flavodoxin as assessed by molecular dynamics simulations and fluorescence anisotropy R. Leenders, and A. J. W. G. Visser

XV

A working proposal for the role of the apoprotein in determining the redox potential of the flavin in flavoproteins: correlations between potentials and flavin pKs L. M. Schopfer, M. L. Ludwig, and V. Massey

399

Polarized absorption spectra of flavin mononucleotide in flavodoxin crystals L. K. Hanson, G. W. Christoph, J. Hofrichter, and M. L. Ludwig

405

Structural analysis of fully reduced A. nidulans flavodoxin C. L. Luschinsky, W. R. Dunham, C. Osborne, K. A. Pattridge, and M. L. Ludwig

409

The site-directed mutagenesis of bacterial flavodoxins R. P. Swenson, G. D. Krey, and M. Eren

415

Structural characterization of site of clostridial flavodoxin M. L. Ludwig, K. A. Pattridge, M.mutants Eren, and R. P. Swenson

423

Redox properties of wild-type and mutant flavodoxins from Desulfovibrio vulgaris (Hildenborough) G. P. Curley, M. C. Carr, P. A. O' Farrell, S. G. Mayhew, and G. Voordouw

429

Characterization flavodoxin the eukariote Chlorella fusca M. L. Peleato, S. of Ayora, and C.from Gomez-Moreno

437

Common structural features of a red algal flavodoxin and cyanobacterial ferredoxins H. Matsubara, K. Fukuyama, and L. J. Rogers

441

Cloning of the ferredoxin-NADP+ oxidoreductase gene and overexpression of a synthetic flavodoxin gene from the cyanobacteria Anabaena PCC 7119 M. F. Fillat, W. E. Bonias, and P. J. Weisbeek

445

Structure/function of spinach ferredoxin:NADP+ oxidoreductase P. A. Karplus

449

Spinach ferredoxin-NADP+ reductase: characterization of the enzymes expressed in Escherichia coli A. Aliverti, B. Curti, G. Zanetti, S. Ronchi, T. Jansen, and R. G. Herrmann

457

Purification and characterization of ferredoxin-NADP+ oxidoreductase from non-photosynthetic tissues S. Morigasaki, and K. Wada

461

+

Ferredoxin binding site of ferredoxin-NADP reductase as explored by limited proteolysis and mutagenesis G. Gadda, A. Aliverti, G. Zanetti, and S. Ronchi

465

Ferredoxins and ferredoxin-NADP+ reductases from Anabaena PCC 7119 and spinach: electrostatic effects on intracomplex electron transfer M. C. Walker, J. A. Navarro, J. J. Pueyo, C. Gomez-Moreno, and G. Tollin

469

XVI Covalently stabilized electron-transfer complexes: a redox active complex between spinach ferredoxin-NADP+ reductase and Desulfovibrio vulgaris flavodoxin G. Zanetti, M. Colombo Pirola, F. Monti, B. Curti, and S. G. Mayhew

475

Structural analysis of the requirements for the formation of an electron transfer complex M. Medina, C. Blancas, M. L. Peleato, and C. Gomez-Moreno

479

Intramolecular electron transfer between ferredoxin-NADP+ reductase and flavodoxin: a laser photolysis study J. J. Pueyo, M. C. Walker, G. Tollin, and C. Gomez-Moreno

483

Evaluation of secondary structure predictions for enzymes related to ferredoxin reductase C. M. Bruns, and P. A. Karplus

487

Roles of four cysteine residues and lysine 110 in human NADH-cytochrome be reductase studied by site-directed mutagenesis K. Shirabe, T. Yubisui, Y. Fujimoto, T. Nagai, T. Nishino, and M. Takeshita

491

DISULFIDE REDUCTASES Pyridine nucleotide-disulfide oxidoreductases-Overview of the family and some properties of thioredoxin reductase altered by site directed mutagenesis: C135S and C138S C. H. Williams, Jr., A. J. Prongay, B. W. Lennon, and J. Kuriyan

497

Structural changes on binding FAD-analogues and on site-directed mutagenesis of glutathione reductase G. E. Schulz, and U. Ermler

505

Exploration of the coenzyme and substrate specificity of glutathione reductase and its subunit assembly R. N. Perham, N. S. Scrutton, A. Berry, and M. P. Deonarain

513

Subunit interactions in the glutathione reductase from Escherichia coli A. Berry, M. P. Deonarain, N. S. Scrutton, and R. N. Perham

521

Active-site mutants of the glutathione reductase from Escherichia coli M. P. Deonarain, N. S. Scrutton, A. Berry, and R. N. Perham

525

Yeast glutathione reductase: determination of the redox potential using NADH and NAD+ L. D. Arscott, D. M. Veine, and C. H. Williams, Jr.

529

Glutathione reductase photoreduction by one-electron donors: stabilization of FAD semiquinone species by complexation J. A. Navarro, M. Roncel, and G. Tollin

533

XVII

Mouse glutathione reductase cDNA. Alignment of the deduced amino acid sequence with the structural domains of the human enzyme M. Tutic, D. Werner, and R. H. Schirmer

537

High level expression of human glutathione reductase cDNA in Escherichia coli based on multiple mutations in the traslation initiation region U. S. Bucheler, D. Werner, and R. H. Schirmer

541

FAD properties of brain glutathione reductase N. L. binding Acan, and E. F. Tezcan

545

The three-dimensional crystal structure of lipoamide dehydrogenase from Azotobacter vinelandii at 2.2 A resolution A. Mattevi, A. J. Schierbeek, G. Teplyakova, and W. G. J. Hoi

549

Site-directed mutagenesis studies on lipoamide dehydrogenase from Azotobacter vinelandii J. A. E. Benen, W. J. H. van Berkel, and A. de Kok

557

Lipoamide dehydrogenase from Azotobacter vinelandii Kinetic studies on wild type and mutant enzymes J. A. E. Benen, N. A. H. M. Dieteren, W. J. H. van Beikel, and A. de Kok

565

Binding studies of the dihydrolipoamide dehydrogenase component (E3) in the pyruvate dehydrogenase complexfromAzotobacter vinelandii E. Schulze, J. A. E. Benen, A. H. Westphal, W. J. H. van Beikel, and A. de Kok

569

Properties of lipoamide dehydrogenase from E. coli modified by site-directed mutagenesis: K53R and I184Y K. Maeda-Yorita, V. Massey, C. H. Williams, Jr., N. Allison, G. C. Russell, and J. R. Guest

573

Replacement of the active site base in E. coli lipoamide dehydrogenase: spectral and kinetic characterization C. H. Williams, Jr., L. D. Arscott, D. Gamm, N. Hopkins, N. Allison, and J. R. Guest

577

Characterization of the active site mutants (C44S, C49S) of Escherichia coli lipoamide dehydrogenase N. Hopkins, C. H. Williams Jr., G. C. Russell, and J. R. Guest

581

FAD-induced dimerization of apo-lipoamide dehydrogenase from Azotobacter vinelandii and Pseudomonas fluorescens W. J. H. van Berkel, and M. C. Snoek

585

Pig heart lipoamide B. N. Leichus, and J.dehydrogenase: S. Blanchaid solvent kinetic isotope effects

589

Kjp effects associated with site-directed mutations thatreducethe velocity of one half reaction of a ping-pong mechanism R. G. Matthews

593

XVIII

Human dihydrolipoamide dehydrogenase: expression and functional aspects of normal and modified enzymes M. S. Patel, H. Kim, and J. E. Jentoft

599

Structure, function and evolution of multiple lipoamide dehydrogenases of Pseudomonas putida J. A. Palmer, K. Hatter, and J. R. Sokatch

603

Dihydrolipoamide dehydrogenase from Halobacterium volcanii: purification, characterisation and molecular cloning N. Vettakkorumakankav, K. J. Stevenson, and M. J. Dan son

607

Electron-transferring flavoproteins from glycine-metabolizing anaerobic bacteria D. Dietrichs, M. Meyer, A. Uhde, W. Freudenberg, and J. R. Andreesen

611

The three-dimensional structure of mercuric ion reductase from Bacillus sp. strain RC607 N. Schiering, K. Fritz-Wolf, W. Kabsch, M. J. Moore, M. D. Distefano, C. T. Walsh, and E. F. Pai

615

On the mechanism of mercuric reductase: an alternating site hypothesis S. M. Miller, V. Massey, D. P. Ballou, C. H. Williams, Jr., M. J. Moore, and C. T. Walsh

627

The streptococcal NADH peroxidase and NADH oxidase: structural and mechanistic aspects A. Claiborne, S. A. Ahmed, P. Ross, and H. Miller

639

Sequence fingerprints for the disulfide reductases: application to the streptococcal NADH peroxidase S. A. Ahmed, P. Ross, H. Miller, and A. Claiborne

647

The structure of Schulz, NADH S. peroxidase from T. Stehle, G. E. A. Ahmed, andStreptococcus A. Claiborne faecalis

651

Peroxide modification of monoalkylated glutathione reductase: evidence for stabilization of an active-site cysteine-sulfenic acid intermediate H. Miller, and A. Claiborne

655

Artificial flavins as probes of the active-site environment and redox behavior of the streptococcal NADH peroxidase S. A. Ahmed, and A. Claiborne

659

Reactivity of the streptococcal NADH oxidase reconstituted with artificial flavins S. A. Ahmed, and A. Claiborne

663

NADH peroxidase: profiles and kinetic isotope effects V. S. Stoll, and J. S.pH Blanchard

667

XIX

COMPLEX FLAVOPROTEINS An active site probe (6-N3-FAD) study of milk xanthine oxidase T. Saito, and V. Massey

673

Electron transfer in xanthine oxidase containing chemically modified flavins D. Shardy, and R. Hille

677

Studies electron transfer in xanthine oxidase R. Hille,ofand R. F. Anderson

681

Evidence for participation of the phosphoseryl residue of xanthine oxidase in the hydroxylation event during enzyme catalysis S. C. D'Ardenne, and D. E. Edmondson

68S

ENDOR studies of the molybdenum centre of xanthine oxidase B. D. Howes, B. Bennett, D. J. Lowe, and R. C. Bray

691

Milk xanthine J. Hunt, and V.dehydrogenase Massey Molecular cloning of cDNA for rat liver xanthine dehydrogenase and its primary structure Y. Amaya, K. Yamazaki, M. Sato, K. Noda, T. Nishino, and T. Nishino

695 699

Cysteine residues responsible for dehydrogenase-oxidase conversion of rat liver xanthine dehydrogenase T. Nishino, Y. Amaya, K. Noda, and T. Nishino

703

Towards an expression system for site-directed mutagenesis studies of xanthine dehydrogenase (Drosophila melanogaster rosy gene) R. C. Bray, R. K. Hughes, W. A. Doyle, J. R. S. Whittle, J. F. Burke, and A. Chovnick

707

Fumarate reductase from Escherichia coli: molecular approaches to the understanding of the function of its prosthetic groups G. Cecchini, B. A. C. Ackrell, M. K. Johnson, M. T. Werth, I. Schroder, D. J. Westenberg, and R. P. Gunsalus

711

Site directed mutagenesis of the putative ligands of the (3Fe-4S) cluster of fumarate reductase of Escherichia coli A. Manodori, G. Cecchini, M. T. Werth, M. K. Johnson, I. Schroder, and R. P. Gunsalus

719

Site-directed mutagenesis of the active site of Escherichia coli fumarate reductase B. A. C. Ackrell, B. Cochran, G. Cecchini, I. Schroder, and R. P. Gunsalus

723

XX

Cloning of the flavo-subunit of human succinate dehydrogenase M. Malcovati, L. Marchetti, T. Zanelli, M. L. Tenchiiii, L. Benatti, T. Simonic, and M. Soria

727

The flavoprotein fraction of NADH-ubiquinone reductase from bovine mitochondria: relationship to a bacterial NAD-reducing hydrogenase S. J. Pilkington, J. M. Skehel, and J. E. Walker

731

Two related forms of the respiratory chain NADH dehydrogenase (Complex I) in Neurospora mitochondria T. Friedrich, G. Hofhaus, G. Tuschen, and H. Weiss

735

Intramolecular electron transfer in trimethylamine dehydrogenase from bacterium W3A1 R. J. Rohlfs, and R.

ffille

739

Structure determination an iron-sulfur flavoprotein C. C. Correll, and M. L. of Ludwig

743

Glutamate synthase from Azospirillum brasilense: role of flavins and iron sulfur centers M. A. Vanoni, D. E. Edmondson, G. Zanetti, and B. Curti

749

Kinetic and structural studies on glutamate synthase from Azospirillum brasilense M. A. Vanoni, L. Nuzzi, M. Rescigno, M. Visentin, P. Accomero, R. Pelanda, M. Pilone Simonetta, G. Zanetti, and B. Curti

755

Sequence motifs in the flavin domain of NADH: nitrate reductase W. H. Campbell

761

Thermodynamics electron in nitrate reductase Kay, and L. transfer P. Solomonson M. J. Barber, C. J.and Redox interconversion of nitrate reductase activity catalyzed by photoexcited flavins M. A. De la Rosa, M. Roncel, and J. A. Navarro

765 769

On some aspects of the catalysis of lactate dehydrogenation by flavocytochrome b2 F. Lederer

773

Substitution of Tyrl43 by Phe in flavocytochrome b2 affects electron transfer from flavin to acceptors C. S. Miles, S. K. Chapman, G. A. Reid, S. A. White, F. S. Mathews, N. Rouvifcre, and F. Lederer

783

Isolation and characterization of the flavin domain of flavocytochrome b2 expressed independendy in E. coli R. L. Pallister, G. A. Reid, C. E. Brunt, C. S. Miles, and S. K. Chapman

787

XXI

Direct measurement of intramolecular electron transfer between the FMN and heme cofactors of yeast flavocytochromes b2 by flash photolysis: evidence for product-mediated conformational gating M.C. Walker, and G.Tollin

791

Cloning and sequencing of the Hansemda anomala flavocytochrome \>2 gene M. Gervais, M. Tegoni, and Y. Risler

797

Hansemda anomala flavocytochrome b^- loss of electron donor reactivity of the flavosemiquinone by pyruvate binding M. Tegoni, F. S. Mathews, and F. Labeyrie

801

Crystal structure of p-cresol methylhydroxylase at 3.0 A resolution F. S. Mathews, Z.-w. Chen, and W. S. Mclntire

805

Interactions in multi-subunit redox proteins: NMR study of the cytochrome subunit of p-cresolmethylhydroxylase W. S. Mclntire, F. S. Mathews, S. Bagby, J. A. Charman, H. A. O. Hill, P. C. Driscoll, and G. L. McLendon

809

FOLATE DEPENDENT FLAVOPROTEINS Stereochemistry of reduction of methylenetetrahydrofolate to methyltetrahydiofolate catalyzed by mammalian methylene-tetrahydrofolate reductase M. A. Vanoni, S. Lee, H. G. Floss, and R. G. Matthews

815

Reconstitution of apophotolyase with pterin and/or flavin derivatives M. S. Joins, B. Wang, S. P. Jordan, and L. P. Chanderkar

819

Chemical modification of Corynebacterium sarcosine oxidase with iodoacetamide H. Suzuki, and Y. Kawamura-Konishi

827

FLAVOPROTEINS OF MEDICAL RELEVANCE Molecular studies of trypanothione reductase: an antiparasitic target enzyme C. Walsh, M. Bradley, S. Sobolov, and F. X. Sullivan

833

Lipoamide dehydrogenase and trypanothione reductase from Trypanosoma cruzi, the causative agent of Chagas'disease R. L. Krauth-Siegel, H. Lohrer, K.-D. Hungerer, and T. Schollhammer

843

Binding of FAD to BCNU-treated apoglutathione K. Becker, T. Schollhammer, and R. H. Schirmer reductase

847

XXII

The EGRAC as a measure of the riboflavin status in man. Titration of hemolysate FAD with apoglutathione reductase K. Becker, and R. H. Schinner

851

Studies on the role of serine-127 in NADH-cytochrome b j reductase by site-directed mutagenesis T. Yubisui, K. Shirabe, and M. Takeshita

8S5

Adrenodoxin reductase of mitochondrial cytochrome P 4 5 0 systems: structure and regulation of expression I. Hanukoglu

859

Squalene epoxidase: the elusive flavoenzyme of sterol biosynthesis D. B. Jordan

865

Expression of membrane-bound flavoenzymes in yeast P. Urban, X. O. Breakefield, and D. Pompon

869

+

Binding of MPP and its analogs to the rotenone/piericidin site of NADH dehydrogenase (NADH-ubiquinone oxidoreductase) T. P. Singer, R. R. Ramsay, M. J. Krueger, and S. K. Youngster

873

Membrane-bound and cytosolic 45kDa proteins in human neutrophils and HL-60 cells Y. Nisimoto, and H. Otsuka-Murakami

877

Covalently-bound flavin in peroxisomal L-pipecolic acid oxidase from primates S. J. Mihalik, and M. C. McGuinness

881

Differences in properties of xanthine dehydrogenase and xanthine oxidase T. Nishino

885

Biochemical characterization of a mutant human medium-chain acyl-CoA dehydrogenase present in patients having deficient activity P. Brass, F. KrSutle, J. Stiemke, S. Ghisla, I. Rasched, N. Gregersen, B. S. Andresen, A. Strauss, and D. P. Kelly

895

Restriction analysis of the human medium-chain acyl CoA dehydrogenase (ACADM) genomic region A. I. F. Blakemore, D. Curtis, P. C. Engel, S. Ktyvraa, and N. Gregersen

901

NON REDOX-ACTIVE FLAVOPROTEINS Acetolactate synthase: a deviant flavoprotein J. V. Schloss, L. Ciskanik, E. F. Pai, and C. Thorpe

907

XXIII

LIST OF PARTICIPANTS

915

AUTHOR INDEX

921

SUBJECT INDEX

927

Introductory Lecture

FLAVINS AND FLAVOPROTEINS - PAST, PRESENT, AND FUTURE

Kunio Yagi Institute of Applied Biochemistry, Yagi Memorial Park, Mitake, Gifu 505-01, Japan

It is my great pleasure and honor to have been invited to deliver an introductory lecture in this symposium. Needless to say, a long long time has passed since the first appearance of the substance that has developed into the biological materials that we now call "Flavins and Flavoproteins". Compared with such long time, the time required for scientists to discover them and to elucidate their biological significance has been surprisingly short, namely, not more than 60 years. The first meeting of this symposium was held in 1965. Now, we are holding our tenth meeting, which means that we have been conducting these symposia for over 25 years, a period of time equal to nearly half of the total used for flavin research. This is also surprising. Riboflavin was found as a vitamin. If we define "vitamin" strictly, the term means nutrients essential to human growth and maintenance of health. The recommended allowance of such substances is only a few mg per day for a person. These substances have enormous physiological action in our body, cannot be synthesized in our body, and cannot be substituted by any other substance. Judging from these characteristics, riboflavin is most typical and probably one of the most important vitamins. So, I want to begin with the discovery of vitamins. Figure 1 shows a map of the South Polar Plateau. One geographic landmark is Takaki Promontory in the Antarctic Peninsula. This place was named by the UK-Antarctic Place-names Committee in 1959 with the explanation that Baron Kanehiro Takaki, Director-General of the Medical Dept. of the Imperial Japanese Navy, was the first man to prevent beriberi empirically

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

4

Fig. 1. A map of the South Polar Plateau. Courtesy of Dr. M. Matsuda.

by dietary additions, in 1882. Four other places surrounding Takaki Promontory were named Eijkman Point, Funk Glacier, Hopkins Glacier, and McCollum Peak. All five persons were pioneers in vitamin research. Figure 2 shows a portrait of Dr. Takaki. The naming of Takaki Promontory surprised Japanese people, because most Japanese had forgotten him as such a pioneer, even though some people still remembered him as the founder of the Jikei University School of Medicine, one of the biggest private medical schools in Japan. In 1883, over 100 years ago, 376 naval officers and sailors travelled in a warship called "Ryujo" to New-Zealand, South America and Honolulu. One hundred sixty-nine of them developed beriberi and 25 persons died. At that time, beriberi was a very serious paralytic disease in Asiatic countries. Dr. Takaki had already started his investigation in 1882 on the cause of beriberi and thought that it could be ascribed to the intake of food that was composed of only polished rice and some pickles. He changed the food from polished rice to European food that was composed of bread, meat, and so on. The following year, namely, 1884, naval officers and sailors of similar number travelled the same route by another warship called "Tsukuba", but no

5

Fig. 2. A portrait of Dr. Kanehiro Takaki. Courtesy of Dr. M. Matsuda.

beriberi occurred at that time. Later, in 1906, Dr. Takaki gave a lecture in England on his findings, and the whole text was published in Lancet. This paper had a strong impact on nutritionists. The results of Dr. Takaki were well confirmed in 1896 by the work of Dr. Eijkman, who demonstrated that chicken beriberi is due to polished rice and that rice bran contains a preventive factor which has a curing effect on beriberi. Then, in 1911, Dr. Funk isolated the beriberi-preventive factor from rice bran, and named it "vitamine". Such essential nutrients were investigated by pioneers such as Drs. Hopkins, McCollum and others by nutritional experiments; and among them Drs. Gyorgy, Kuhn, and Wagner-Jauregg found in 1933 a growth-promoting factor in the liver and heart, and then isolated a yellow pigment as such factor from egg white and named it "ovoflavin". Figure 3 shows a portrait of Dr. Paul Gyorgy. Independently, Drs. Ellinger and Koshara found "lyochrome" in milk, and showed it to be identical to ovoflavin and subsequently called it "lactoflavin". Then, Dr. Kuhn and his coworkers isolated them in crystalline form and showed them to be identical. In 1934, Dr. Karrer and his coworkers crystallized the same substance from the liver. Then, Kuhn's group and Karrer's group chemically synthesized them independently, and thus the structure was finally decided. From the chemical structure the term "riboflavin" was

6

Fig. 3. A portrait of Dr. Paul Gyorgy. Courtesy of Dr. K. Koike,

chosen. When I was a medical student in the University of Nagoya in 1939, these findings were already a part of the lecture content of our biochemistry course. However, in the lecture one of the most emphasized new findings was the discovery by Prof. Hugo Theorell of riboflavin phosphate as the prosthetic group of the "Gelbes Oxidations Ferment" of Prof. Otto Warburg. This enzyme was later called the "old yellow enzyme". It was emphasized that this discovery indicated a connection between vitamin and enzyme, or in wider meaning, between nutrition and biochemistry. I knew the name of Prof. Theorell at that time; and to my pleasure, later I was invited by him to study in the Nobel Medical Institute in 1957. Figure 4 shows a portrait of Prof. Hugo Theorell. I had many pleasant days with him there, but the most exciting days we shared were those during his stay in Japan for his plenary lecture in the General Assembly of the Japan Medical Association held in Nagoya in 1967. While relaxing in Japan, he told me personally an episode concerning his discovery of riboflavin phosphate by saying that "After I found that the pigment of the prosthetic group of the old yellow enzyme is riboflavin, I thought that riboflavin would be in its phosphoric acid ester. And I wanted to check this with the isolated prosthetic group. But it was already colourless. So I thought riboflavin was reduced or decomposed. By aeration, it could not be oxidized, and I realized that riboflavin was decomposed probably by some

7

Fig. 4. A portrait of Prof. Hugo Theorell. Courtesy of the Chunichi newspaper, Nagoya.

bacteria. But I thought phosphoric acid should remain. And I found an equimolar amount of phosphoric acid to riboflavin". Then, he educated me by saying that "You must not abandon any sample, even a minute amount, until the completion of your work." I want to return now to the time when I was a medical student. Because of my poor knowledge of chemistry, I asked the Professor of Biochemistry if I could study chemistry in his laboratory. Upon his approval, very often I worked there until late in the evening and sometimes even slept there. Fortunately and to my surprise, there was a beautiful large bed in the laboratory. I was told that the bed was the very one that had been used by Prof. Leonor Michaelis when he lived in Nagoya. I learned that Prof. Michaelis was the founder and the first professor and director of the Department of Biochemistry of the Aichi Medical College, now the Institute of Biochemistry, Faculty of Medicine, University of Nagoya. Figure 5 shows a portrait of Prof. Michaelis. He came to Nagoya in 1922 at the invitation of the Aichi Medical College, served there for 3 years and a half, and then left for the Rockefeller Institute in New York. When I was a medical student, many things he brought from Germany still remained in the laboratory. Somewhat influenced by Prof. Michaelis, I decided to study the enzyme-substrate complex predicted by him, when I started my own scientific work just after the Second World War in 1945. For this study, I

8

Fig. 5.

A portrait of Prof. Leonor Michaelis. Courtesy of Mrs. Ilse Michaelis Wollman and Mrs. Eva Michaelis Jacoby.

thought that flavoprotein would be a very good tool, since flavin coenzyme should serve as an indicator of an enzyme for its oxidoreduction state as well as for its complex formation with a substrate. At that time, as dissociable flavin enzymes, we knew only the old yellow enzyme and D-amino acid oxidase, and I selected the latter. After a longer time than I had expected, I obtained a crystalline complex of D-amino acid oxidase and D-alanine, in 1962. Figure 6 shows crystals of the so-called purple intermediate complex. This crystal is composed of equimolar amounts of the enzyme and the substrate. There is a strong charge transfer interaction between them, and they slowly dissociate into the semiquinoid form of the enzyme and the substrate

Fig. 6.

Crystals of the enzyme-substrate complex of D-amino acid oxidase. Courtesy of Elsevier Publishing Company.

9

Fig. 7. A portrait of Dr. R. K. Morton (left) with the author (right).

radical. This marked the first crystallization of an enzyme-substrate complex. By a series of papers concerning this finding published in Biochimica et Biophysica Acta, I became recognized by many specialists in enzymology. Among them, Dr. R. K. Morton paid special attention to me, and he visited me in Nagoya in 1962 on the way back to his home in Adelaide, Australia, from England. Although I knew his name by his work on crystallization of cytochrome b2, we met for the first time in Nagoya. Figure 7 shows a photograph of Dr. Morton and myself in my office. After fruitful scientific discussion, he told me that the international symposium on cytochrome he had organized in Canberra had been successful and that he wanted to organize a similar symposium on flavin enzymes to be held there as well. He knew that I had already organized a small international symposium on flavins and flavoenzymes in 1957 in Nagoya, and asked my opinion about this. I agreed with his idea. Unfortunately and sadly, he died on September 27th of the next year, following a laboratory accident. As for the symposium, however, "Flavins and Flavoproteins" was actually held. I got an invitation letter from Prof. E. C. Slater, and at that time I thought that the desire of Dr. Morton was to be realized by his good friend Prof. Slater following the passing of Dr. Morton. However, when I attended the symposium, which was held at the Trippenhuis, the home of the Royal Netherlands Academy of Sciences and Letters, between June 10th and June

10

15th of 1965, I realized that my supposition was not completely correct. From the opening address made by Prof. C. J. Gorter I learned that Dr. Morton had already visited Amsterdam before his visit to my laboratory and had discussed the matter of the symposium. Prof. Gorter said that "When the project to hold the symposium in Canberra turned out to be impracticable, Professor Morton proposed that it be held in Amsterdam. After Professor Slater obtained a favourable response from the leading workers in the field, he and Professor Veeger undertook the organization with the support of an Advisory Committee consisting of Professors Morton, Massey, Beinert, Chance, Handler, and Hemmerich. Both the International Union of Biochemistry and the Royal Netherlands Academy of Sciences and Letters agreed to sponsor the symposium, and sufficient financial support was obtained from various sources to ensure your presence today", and so on. Anyhow it is clear that the Amsterdam symposium was first proposed by Dr. Morton and realized by Prof. Slater and his colleagues. The meeting was exciting. I still vividly remember the days in Amsterdam, and I am happy to tell you that among the participants in that meeting, Drs. Robert Bray, Paolo Cerletti, Vincent Massey, Graham Palmer, Thomas P. Singer, and Ces Veeger are here with us this evening. Impressed by the Amsterdam meeting and realizing that the field was rapidly advancing, I wanted to organize a similar meeting. Taking advantage of the occasion of the 6th Congress of the International Union of Biochemistry held in Tokyo in 1967, I contemplated organizing a symposium in Nagoya. I asked Dr. Slater about his opinion, since I was afraid that the interval between two such meetings would be too short. Dr. Slater answered me in a favourable way, and even allowed me to call the symposium the "Second International Symposium on Flavins and Flavoproteins". Then, the Amsterdam symposium automatically became the 1st Symposium. The 2nd symposium was therefore held in Nagoya, and at that time, Dr. Henry Kamin volunteered to hold the 3rd meeting in the United States to continue the series. Unfortunately he passed away on September 15, 1988. I will never forget his warm and thoughtful character besides his excellent scientific contributions. Subsequent symposia have been held periodically ever since.

11 Table 1. International

Symposia

No.

Year

1

1965

Amsterdam

2

1967

Nagoya

3

1970

Durham

4

1972

5

6

Place

Flavins and Flavoproteins

Main Organizer (s)

Publisher

E. C. Slater C. Veeger

Elsevier Publishing Co.

K. Yagi

Univ. Tokyo Press Univ. Park Press

H. Kamin

Univ. Park Press Butterworth & Co.

Konstanz

P. Hemmerich

(Zeitschrift für Naturforschung)

1975

San Francisco

T. P. Singer

Elsevier Scientific Publishing Co.

1978

Kobe

K. Yagi T. Yamano

Japan Scientific Societies Press Univ. Park Press

1981

Ann Arbor

V. Massey C. H. Williams, Jr.

Elsevier/North-Holland

1984

Brighton

R. C. Bray P. C. Engel S. G. Mayhew

Walter de Gruyter

9

1987

Atlanta

D. E. Edmondson D. B. McCormick

Walter de Gruyter

10

1990

Como

B. Curti S. Ronchi G. Zanetti

Walter de Gruyter

Table 1 summarizes the number of symposia, main organizers, and the names of the publishers of the proceedings. Among the organizers listed in this table, another scientist, Prof. Peter Hemmerich, passed away on October 3rd, 1981. He was a serious and enthusiastic scientist. I will always remember his forceful discussions, not to mention his excellent scientific contributions. We learned much from him. A well-arranged sketch of his life and work was made by Dr. Massey and appeared in the first part of the proceedings of the 7th symposium. When I prepared the text for the present talk, I piled all the proceedings on top of one another, and found the height to exceed 30 cm. All the important findings related to flavins and flavoproteins made over the last 3 decades lay contained in the pile before me. Skipping further comment on these

12

|

Fig. 8. Appearance of a riboflavin-deficient rat.

important findings, I want to emphasize some medical implications of flavins deduced from my own research. Shortly after the Second World War, I started to work on flavoproteins, as I told you before; and at the same time I also studied riboflavin deficiency from a medical point of view. Figure 8 shows the appearance of a rat fed a riboflavin-deficient diet. Typical cataract, namely, the turbidity in the lens of the eye, is observable in this rat. Bleeding and some inflammation surrounding the eye and the mouth also occurred in this animal. These phenomena had already been found and described by earlier researchers, but the organs and tissues of such animals had not been examined in detail. When I dissected a typical riboflavin-deficient rat, I found changes in blood vessels, especially atherosclerosis-like change in the arteries. As riboflavindeficient animals look like aged animals, and since cataract and atherosclerosis are representatives of age-related diseases, I thought at that time that ariboflavinosis might be a model of age-related diseases and even of aging itself. Shortly later, Dr. Koichi Masuda reported that an endemic disease called "Shibi" disease, which occurred in Goshogawara, in the northern part of the main island of Japan, could be ascribed to riboflavin deficiency, and could be cured by the administration of riboflavin. Further, Prof. Takeshi Kimura reported a high incidence of atherosclerosis in this disease. Since the artery or the lens contains little or no riboflavin, such a change in

13 Table 2. Changes in serum lipid peroxide levels of rats. Serum lipid peroxide level (nmol/ml) Riboflavin in diet

Time on diet (week) 1

+

2.9 + 0 . 9 ( 1 0 ) 3.1 ± 1 . 0 (10)

2 3.1 ± 0 . 6 (10) 3.1 ± 0 . 8 (10)

3 2.8 ± 0 . 4 (11) 3.2 ± 0 . 4 * (13)

Serum lipid peroxide level was measured by the method of Yagi, and expressed in terms of malondialdehyde. Mean value ± SD is given. Numbers in parentheses indicate the number of animals tested. Significant difference from the corresponding value for rats fed a riboflavin-containing diet: *p12. C) N(5)-H which can CI) protonate or r C2) deprotonate at

reduced Flavin (Fl,.^)

low or high pH values. Scheme 1

pK>22

Ionisations of fully reduced flavin.

A) The importance of the N(l)-H ionisation has long been recognized. It strongly influences the reactivity of the reduced molecule, in particular towards oxygen. A shift of this pK also reflects changes in the redox state of the isoalloxazine

[1].

B) The ionisation of N(3)-H does not appear to have a major role in influencing the catalytic properties of the flavin. In some cases, e.g. with oxidized glycollate oxidase, the p K a of N(3)-H is lowered from -10 to ~6.7 due to the interaction with the

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

28 protein [2], Whether a similar interaction occurs between the reduced flavin and the protein is not known. 15N-NMR measurements at high pH (10-12) [3] give a pK a >12 for the reduced flavin N (3)—H, and with all proteins studied so far, this function appears not to be ionized [4]. C) N(5)-H does not have a pK in or near the physiologically relevant range. However, the N(5) position is directly involved in the uptake and release of redox equivalents. Its electronic configuration undergoes the largest changes (sp2 -> sp3) during the transition between the oxidized and the reduced state, i.e. during catalysis. In particular, it can serve as the acceptor of the hydrogen bound to the substrate a-carbon:

Flox

+

—C-*H OH

F'red N(5)-*H +

O

Since N(5)-H has been found to exchange with solvent [5], the mechanism(s) of this exchange is of biochemical importance. Dudley and coworkers [6] estimated the pK a for the protonation of N(5)-H as , authentic 6-HDNO. The transformation of the purified apo-6-HDNO (derived from the fusion protein) to holo-6-HDNO depends on FAD and on one of the known effector molecules (PEP, G3-P) (Fig. 3). The kinetic data essentially match those of the native enzyme, although the K m for FAD was found

0.5 instead of 3 /tM. The Ka value of the effectors are

0.5 mM for glyceraldehyde 3-phosphate and 1 mM for G-3-P or PEP. Such concentrations have been observed in cell lysates. The FAD attachment occurs best at a pH near 7, while it cannot be observed at pH 9.2 at which the 6-HDNO assay is run. The apoenzyme is much more susceptible to proteolytic degradation than holo-6-HDNO (5). The presence of 45% glycerol not only stabilizes the apoprotein but also appears to allow a conformation that facilitates the access of FAD to its binding site and the spontaneous covalent attachment. Still, G-3-P increases the rate of holoenzyme formation even in the presence of glycerol. The same observation was made in the presence of 20% saccharose.

104

FIG. 3 FAD- and effector-dependent holoenzyme formation from purified apo-6HDNO. 100 ni samples (1 ng protein) of purified 6-HDNO (from the fusion protein) in 20 mM Tris-HCl buffer, pH 7.0, 10 mM MgCl2, 5 mM mercaptoethanol and 45 % glycerol were incubated at 30°C for 30 min. Panel A: const. G-3-P (10 mM), variable FAD concentration (•). Panel B: const. FAD (7.5 2, b2), which also catalyze dehydrogenation of a-hydroxyacids and are believed to share mechanistic similarities. The three-dimensional crystal structures of these enzymes are now available [2,3]. Based on the similarity in protein structures around the flavin in these two enzymes, and strictly conserved residues in the primary structure of IX), we have used these data to interpret in molecular terms thereactionmechanism of IX), and interpret to die eflect of interaction of specific amino acidresidueswith the flavin on the physicochcmical properties of the flavoenzyme.

Structural Comparisons of Lactate Oxidase with Spinach Glycollate Oxidase and Yeast Flavocytochrome b2The subunit sequence of393 amino acids of LO shows considerable homology with the sequences of GO and b£ [ 1,4]. In the sequence comparison 69 residues show identity between till three enzymes. Included among these are residues which are involved in binding to the FMN side chain or isoalloxazine ring, or residues implicated in substrate binding and catalysis. Thus, Arg413 and Arg433 form salt linkages to the phosphate ofFMN, Asp409 is Il-bonded with the FMN ribityl-3'-

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

124 OH and Lys349 is H-bonded with the ribityl-2'-OH in b2- Exactly the same functions in the GO crystal structure are ascribed to the homologous residues Arg289, Aig309, Asp284 and Lys230 [3]. With LO die existence of the same homologous residues, Aig324, Arg344, Asp320 and Lys266, suggests strongly a veiy similar binding of FMN. Similar interactions are also seen between the protein and isoalloxazine ring, with 0(2) (hydrogen bond with Thrl55 in GO, Thf280 in b2 and presumably Thrl 78 in LO). Interaction of 0(4) with Gin 127 and Tyr 129 in GO is paralleled by the same interaction with Gln252 and Tyr 254 in t>2 and presumably by the equivalent Gin 150 and Tyr 152 in LO. Among the amino acid functions, which are assumed to be part of the catalytic machinery the conservation is impressive, as shown in figure 1.

Figure 1: Comparison of active center functional groups for lactate oxidase (LO), glycollate oxidase (GO) and flavocytochrome b2 (b2). Binding of the substrate in the active site appears to be similar in all three enzymes. From the crystal structures it is known that product or competitive inhibitor are bound on the Si-side of the flavin {2,3]; the same side was found with LO in stereospecificity studies employing 8-OH-5-deazaFMNH2 [5,6]. With b2 the caiboxylate of the substrate is held in a salt bridge with Arg 376 and by a hydrogen bond with the hydroxyl of Tyr 143 [2,7]. The same interactions are to be expected in the crystal structure of GO with Aig 257 and Tyr 24 and also presumably in LO where the same functions would be fulfilled by the strictly homologous residues Arg293 and Tyr44. All three proteins contain a strictly conserved histidine residue (His373 in b2, His 254 in GO and presumably

125 His290 in LO) which is believed to be the active site base responsible for the abstraction of the substrate a-hydrogen as a proton to form the carbanion species which then transfers its electrons to the flavin. In both b2 and GO there is an aspartate residue (Asp 282 in b2 and Asp 157 in GO) with its caiboxylate close to N( 1 ) of this histidine residue, and ascribed a typical change relay role. An homologous Asp 180 is found in LO. Finally a tyrosine residue loeated on the substrate binding side of the flavin (Tyr254 in hi, Tyrl 29 in GO and Tyr 152 in LO) appears to be strictly conserved, and has been thought possibly to play an important role in catalysis, since in both t>2 and GO, the oxygen atom of the tyrosine hydroxyl is in hydrogen bond distance to the substrate a-OH, and thus may serve to facilitate the removal of the proton from the substrate hydroxyl to form the final keto acid product. However, in the Tyr254Phe mutant of t>2 there is 2% residual catalytic activity [8] and the GO Tyrl 29Phe mutant has approximately 10% the catalytic activity of the wild type enzyme [9]. Catalysis a n d role of functional groups Although a three dimensional structure does not yet exist for LO the homologies discussed above, and the similarities of the chemical reactions catalzyed (see discussions in [4,7) allow the deduction that the three enzymes (LO, GO and b2) work by the same basic chemical mechanism. The mode of substrate binding can be derived from the three dimensional structures of the active center published for GO and b2. In figure 2 we have simply placed the substrate lactate in the active center cavity such as to yield an optimal interaction. The same procedure was carried through in a three dimensional system using the conventional stick-frame display and space filling models (not shown) from which it can be confirmed that the two dimensional picture of figure 2 is reasonable. In the model of figure 2 catalysis will be initiated by abstraction of the a-hydrogen as a proton. The base involved has been proposed to be a histidine [2] (His 290 in LO, see figure I), which is linked to an aspartic acid (Asp 280), in a charge relay system. Tyr 44 interacts with the carbonyl or the carboxylate and this might contribute to the acidi fication of the a-hydrogen. Tyrl 52 is shown to interact with the substrate aOH group and might help in keeping it in the proper orientation. A question which is often brought up in the context of the reaction mechanism, refers to the mode of stabilization of the carbanion formed, i.e. of the transition state. We have proposed oxalate to be a transition state analog [ 10,11J binding in a bidentate manner to a basic group serving in the fixation of substrate (Atg293 in LO), and to the protonated base, which abstracts the a-hydrogen, in this case His290 [4]. Oxalate (dianionic form) can be fitted nicely into the active center of models of LO where it interacts with the two positively charged amino acids Arg293, His290, and the two tyrosines 44 and 152. (Full) délocalisation of the negative charge of the substrate carbanion to the carbonyl of the C(l)-caiboxy1ate would form a planar molecule as shown in figure 3, in which the C a negative charge is stabilized by the interaction with Tyr 44. This could go as far as to reach a complete transfer in which Tyr44 is in its anionic form, as shown. Clearly in a transition state such an extreme (mesomeric) form is not required to exist, and transfer of chargé might be only partial. In any case the interaction described appears appropriate for such a transition state stabilization interaction. Important mechanistic

126

Figure 2: Possible orientation of lactate, isoalloxazine and functional groups involved in binding and catalysis in the Michaelis complex of lactate oxidase.

Y152 Figure 3 Possible mode of stabilization of (a mesomeric form of) the carbanion generated by abstraction of the substrate a-hydrogen as a proton by His 290. The protonated form of Tyr44 is envisaged to form a hydrogen bond to the carbonyl of substrate information on lactate oxidase was obtained from the study of its reaction with glycollate [ 12-14]. With the latter, in contrast to the case of L-lactate, both hydrogens (Re and Si) were shown to

127 undergo abstraction, although at vastly different rates [ 13]. The salient difference between the reaction with L-lactate and glycollate, however, is the occurrence with the latter of two different intermediates arising from abstraction of the a-hydrogen, and before (or concomitant with) formation of die complex of reduced enzyme and glyoxylate. These two intermediates were shown 113,14] to be derived from abstraction, respectively of the Re- and Si hydrogen of glycollate, and thus most probably have enantiomeric structures. Most importantly the one derived from rupture of the Sihydrogen bond isrelativelystable. Its structure was shown to be that of an N(5) glycollyl flavin adduct as shown in figure 4A. By analogy and by comparison of the corresponding properties the second intermediate was proposed to be that derived from abstraction of the Re-hydrogen (which corresponds stereochemically to the a-H in L-Lactate) [13,14]. The two sets of reactions leading to the two intermediates are shown infigures4A and 4B. The difference in stability between the two adducts can be nicelyrationalizedby comparing the three dimensional orientation of the asubstituents and their interactions with Tyr 152. In the case of the labile adduct (figure 4A) the interaction is shown to facilitate the fragmentation of the adduct, while it is absent in the stable adduct (figure 4B). This proposal requires an active role ofTyrl52 in catalysis. In fact, in a Tyr254Phe mutant of b2 and in a similar one (Tyrl 39Phe) of GO the activity is reduced to 2 and 10 % that of the native enzymes [8,9]. Theresultsobtained with glycollate and described earlier clearly show that N(5) covalent adducts are viable intermediates in the catalytic dehydrogenation of glycollate. that this deduction can be extrapolated to the mechanism of dehydrogenation of L-lactate is not certain in all its details, but we think itreasonableto assume, that the same basic mechanism will be operative within this family of enzymes and substrates. As discussed in detail elsewhere [4,12], the covalent N(5) adduct might be a true intermediate only in the case of glycollate, where the stericrequirementsare lower than for lactate. With the latter the adduct might be a transition state, i.e. formation of the N(5)Ca bond might be incomplete and be concerted with fragmentation. In fact, the crystal structures of b2 and GO suggest the possibility of steric overcrowding in the substrate binding site. With both enzymes the peptide chain comes close to theflavinRe face, with Ala 198 in \>2 and Ala 79 in GO being the nearest residue to the flavin N(5) position. In LO the homologous residue is Gly99. The distance between N(5) and the CH3 group of Ala79 in GO is 4. 6 A . If the arrangement of the protein around the flavin is similar in LO to that in b j and GO, there should be no overcrowding between Gly99 and the glycollyl cflH, permitting the formation of the observed flavin N(5)-glycollyl adducts. However, with lactate as substrate steric crowding introduced by the methyl residue may prevent the formation of a stable lactoyl adduct Also, in a putative lactoyl-N(5) adduct in the active center of either GO or l>2, steric overcrowding seems probable, and would result in destabilization of the adduct. It is conceivable that evolution has created conditions in which the transition statelntermediate is not stabilized too much in order to promote catalysis. Furthermore, work with chemical models has shown that overcrowding plays a crucial role in the stability of N(5) adducts. Thus, the adduct of formaldehyde toflavinN(5) is quite stable,while the corresponding one with acetaldehyde cannot be observed [15]. Similarly N(5) alkylated flavinium cations cannot be obtained when the substituent is

128

isopropyl, while they are stable in the case of ethyl, and methyl [16].

^OOC o

HrT^ib-^ QsC-

-H, OH

N. ^N®.O

H

(Bo)

O

"OjC-jp-Hs, IB"" O

Figure 4 (A), top panel: Reaction of glycollate involving abstraction of Hr c and formation of a labile covalent adduct which can decay to reduced enzyme and glyoxylate. Note that in this case the interaction of the glycollate a-OH of the adduct with Tyrl 52. promotes the decay.

129

1

D

- —

/

r o

tl «a; M wc H(

b rvu "OjC—_y

a,c—I—OH

"He

Hne

t ^ C - j - OH

H(S|) B—

0

Hne I IB"

fY152l

K266

Figure 4: (B), top panel: mode of interaction of glycollate with the active center of LO. Note that the glycollate Hg;reactsand leads to formation of a covalent adduct, which does not decay toreducedenzyme and glyoxylate. Lower panel: proposed structure of the stable covalent intermediate and interaction with active center functional groups. Note the absence of interaction of the glycollate a-OH with Tyrl 52.

130 Acknowledgements: We acknowledge many interesting discussions 'with Dr. F. Lederer on mechanistic topics, and financial support from the Deutsche Forschungsgemeinschaft to SG and from the U.S. Public Health Service (GM 11106:) to VM.

References: 1. Giegel, D.A., Williams, C.H.Jr., and Massey, V. 1990. J.Biol.Chem. 2656626-6632 2. Lederer, F. and Mathews, F.S. 1987. in Flavins and Flavoproteins (Edmondson, D.E. and McCormick, D.B. eds) pp 133-142, de Giuyter, Berlin, 3. Lindqvist, Y., and Branden, C.I. 1989. J.Biol.Chem. 264,3624-3628 4. Ghisla, S., and Massey, V., 1991. in "Chemistry and Biochemistry of Flavoproteins" (Müller, F., ed), CRC press, Inc, in press 5. Manstein, DJ., Massey, V., Ghisla, S. and Pai, E.F. 1988. Biochemistry 27,2300-2305 6. Manstein, D.J., Pai, E.F., Schopfer, L.M. and Massey, V. 1986. Biochemistry 25,6807-6816 7. Lederer, F., 1991. in "Chemistry and Biochemistry of Flavoproteins" (Müller, F., ed), CRC press, Inc, in press 8. Reid, G.A., White, S., Black, M.T., Lederer, F., Mathews, F.S. and Chapman, S.K. 1988. Eur. J. Biochem. 178,329-333 9. Macheroux P., Massey, V., Thiele, DJ., Söderlind, E., and Lindqvist, Y., this volume 10. Ghisla, S., and Massey, V., 1975. /. Biol. Chem. 250,577-584 11. Ghisla, S., and Massey, V. 1977. J. Biol. chem. 252,6729-6735 12. Ghisla, S., and Massey, V.1989. Eur. J. Biochem. 181,1-17 13. Massey, V., Ghisla, S., and Kieschke, K., 1980. J. Biol. Chem. 255,2796-2806 14. Ghisla, S., and Massey, V. 1980. J. Biol. Chem. 255,5688-5696 15. Blankenhorn, G., Ghisla, S., and Hemmerich, P. 1972. Z. Naturforechung 27B, 1038-1040 16. Ghisla, S., Hartmann, U., Hemmerich, P., and Müller, F., 1973. Liebig"s Ann. Chem. OSSMIS

LACTATE OXIDASE: MUTAGENESIS AND EXPRESSION OF THE MYCOBACTERIAL GENE

Ute Müh, David A. Giegel, Vincent Massey, and Charles H. Williams, Jr. Department of Biological Chemistry, and Department of Veterans Affairs, Ann Arbor, MI. 48109, USA

Introduction Lactate oxidase from Mycobacterium smegmatis is an FMN-dependent monooxygenase, the mechanism of which has been extensively studied (1). The enzyme catalyzes the oxidation of L-lactate to acetate, carbon dioxide and H 2 0 , using molecular oxygen as oxidant. The gene for the protein fromM smegmatis has recently been cloned and sequenced (2). The enzyme crystallizes in bright yellow plates, but the structure so far has escaped determination. The investigation of the active site is therefore aided by studies of homologous proteins. Lactate oxidase (LO) is mechanistically related to flavocytochrome b 2 (b 2 ) and glycolate oxidase (GO). The structure and sequence of both enzymes have been determined (3,4). Sequence alignment shows an identity of 33 % (GO) and 25 % (b 2 ) with an overall homology of 61 % and 44 % respectively. All residues constituting the active sites of GO and b 2 can be found as identical residues in homologous positions in lactate oxidase. Homology Comparison and Mutagenesis Lactate oxidase, glycolate oxidase and flavocytochrome b 2 catalyze the oxidation of an a-hydroxycarboxylic acid to an a-ketoacid. The enzymes are comparable in the reductive half-reaction. They differ, however, in the ensuing oxidation of the flavin. In b 2 the electrons are passed on to protoheme IX, located in a separate domain of the protein. In both GO and lactate oxidase, molecular oxygen enters the active site and is reduced to H 2 0 2 . In GO the oxidized product and H 2 0 2 are then released. In lactate oxidase, however, both reaction products are retained sufficiently long

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

132

for the peroxide to decarboxylate the pyruvate to acetate. For all amino acid residues that had been proposed in earlier investigations (1) to be responsible for catalysis and substrate binding in lactate oxidase, residues could be identified in the protein sequence, aided by the strong homologies to the active site residues in G O and b 2 . A lysine close to the N ( l ) had been proposed to stabilize the anionic form of the reduced chromophore. A histidine on the si side of the pyrimidine-ring of the flavin is a probable candidate for the active site base. By modification studies an arginine had been proposed to bind the carboxylic moiety of the substrate (5). We have selected these residues as targets of site-directed mutagenesis. Additionally we have changed the two cysteines that were inactivated with FDNB (C104, C 203) to alanines (6). A further target is cysteine 287, which introduces a charge unique to the active site of lactate oxidase (Val in GO and Leu in b 2 ), table 1. TABLE 1: Targets of Mutagenesis (* Completed) Indicated Residue FDNB modification

Negatively charged

Mutation

Indicated Residue

Mutation

*

C 104 A

Active site

H 290 Q

*

C203A

base

*

C287A

Substrate

Y 44 F

binding

Y 152 F

residue in LO

R 293 K Positively charged

*

K 266 M

residue close to N ( l )

*

K 266 R

Stabilization

D 180 A

of base

Expression of the Gene in E. coli Lactate oxidase shows only low levels of expression in E. coli under control of either the tac (pKK 223.3 from Pharmacia) or the phage T7 promoters (pSKM13+ from Stratagene). Current data indicate that the problems encountered are not due

133

to the expression of insoluble protein (inclusion bodies). We are working with the hypothesis that translation is reduced, probably due to unfavorable signals for the E. coli protein synthesis complex. Improving the codon usage and the sequence for the ribosome binding site have both led to an increase in lactate oxidase activity. Looman et al. (7) describe the influence of the codon following the initiation methionine on the level of transcription. The described efficiences were assigned values ranging from 0.1 % to 1.5 % with an average of 0.4 %. We have performed a silent mutation on the serine following the initiation codon, thereby changing its efficiency value from 0.2 % to 0.8 %. The mutation was designated SIS. The D N A sequence from Mycobacterium shows a region 5' to the gene with a strong homology to the Shine-Dalgarno consensus sequence (8). However, the spacing from the initiation A U G is only 4 basepairs long and contains predominantly GC - nucleotides. By polymerase chain reaction we have reconstructed the upstream region of the gene. A 45mer oligonucleotide was constructed, which contained the T7 glO-L ribosome binding site as described by Olins (9). We are cloning the gene for lactate oxidase into the expression vector pET-3a from Studier et al. (10). This vector is pBR 322 - derived and contains the T7 promoter as well as the translation signals for gene 10 protein. The ribosome binding sequence is followed by an Nde I site which allows the insertion of the target gene with optimal spacing to the initiation AUG. Fig. 1 shows the levels of expression achieved with the various systems. Conclusions Traditional biochemical techniques have been used extensively to investigate the reaction mechanism of lactate oxidase (4,11). The architecture of the active site was further confirmed when the protein was sequenced and homology comparisons could be made with mechanistically related enzymes. To improve the current understanding of the mechanism, it is of interest to specifically exchange the indicated amino acids. We have constructed the required mutagenesis and expression system and will now proceed to analyze the enzymatic activities of the site-directed mutants. As a further goal we hope that the purification of recombinant protein may lead to crystals that facilitate the determination of the three-dimensional structure.

134

Fig. 1:

EXPRESSION SYSTEMS, specific activity ( M mol 0 2 / min x mg protein) measured in the crude extract of the varied expression systems in E. coli

Construct pGM 3 pGM 5 pGM 9 pGM 15

Promoter T 7 tac T 7 T 7

Plasmid pSKM13+ pKK 223.3 pSKM13+ pSKM13+

Improvement SIS SIS, per modified ribosome binding site

Acknowledgements This work was supported by the Health Services and Research Administration of the Department of Veterans Affairs (C.H.W.) and by grants GM21444 (C.H.W.) and GM11106 (V.M.) from the National Institute of General Medical Sciences.

References 1. 2. 3.

Ghisla, S. and Massey, V. (1980) J. Biol. Chem. 255, 5688-5696 Giegel, D.A., Williams, C.H., Jr., and Massey, V. (1990) J. Biol. Chem. 265, 6626-6632 Lederer, F., and Mathews, F.S. (1987) in Flavins and Flavoproteins (Edmondson, D.E., and McCormick, D.B., eds) pp. 133-142, Walter de Gruyter, Berlin 4. Lindqvist, Y„ and Bränden, C.-I. (1989) Biol. Chem. 264, 3624-3628 5. Peters, R.G., Jones, W.C., and Cromartie, T.H. (1981) Biochemistry 20, 2564-2571 6. Giegel, D.A., Massey, V. and Williams, C.H., Jr. (1987) J. Biol. Chem. 262, 5705-5710 7. Looman, A.C., Bodlaender, J., Comstock, L.J., Eaton, D., Jhurani, P., de Boer, H.A., and van Knippenberg, P.H., (1987), EMBO, 6, 2489-2492. 8. Stormo, G.D., Schneider, T.D., Gold, L.M., (1982), Nuc. Ac. Res., 9, 2971-2996. 9. Olins, P.O., Devine, C.S., Rangwala, S.H., Kavka, K.S., (1988), Gene, 73, 227-235. 10. Studier, F.W., Rosenberg, A.H., Dunn, J.J., and Dubendorff, J.W., Methods in Enzymology 185, in press 11. Massey, V., and Ghisla, S., (1990) in Flavoproteins (Muller, F., ed.) CRC, submitted

MOLECULAR BIOLOGICAL STUDIES ON STRUCTURE-FUNCTION RELATIONSHIP OF D-AMINO ACID OXIDASE

Yoshihiro Miyake, Kiyoshi Fukui, Kyoko Momoi, Fusao Watanabe, Masazumi Tada, Motoshige Miyano, Saori Takahashi Department of Biochemistry, National Cardiovascular Center Research Institute, Suita, Osaka 565, Japan

Introduction In order to investigate the structure and function of D-amino acid oxidase (DAO), we isolated cDNA clones encoding the entire protein sequence of DAO from pig, human, rabbit, and mouse kidney cDNA libraries and determined the nucleotide sequences (1-4). We also constructed seven point-mutated pig DAO cDNAs (5, 6) and expressed the mutant DAOs by an in vitro expression system (7). Moreover, we developed a large scale expression system and purification procedure for recombinant pig DAO (8) . In this report, gene expression of the DAOs and the structure and function of recombinant wild type and mutant DAOs are compared with each other.

Results. Tissue DAO activities in animal kidneys DAO activities of tissue extracts from pig, human, rabbit, and mouse kidneys were measured by a spectrophotometric method (7). Pig kidney showed a high activity (0.205 unit/mg protein). However, the activities of human and mouse kidneys were 0.013 and 0.001 unit/mg protein, respectively, and it was undetectable in rabbit kidney with 20 mM D-alanine and at 25 °C.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

136 5'-UT

3'-UT

ORF

198 nt

1041 nt M

347a.a.

200 nt ^G(-3) M

Pig (3211 n t ) 392 nt

1041 nt

Human(l633 nt)

347a.a.

jC(-3)

701 nt

1972 nt

M

1041 nt

351 nt

Rabbit (2093 nt)

347a.a.

68nt jG(-3) M \

/ 25

544 nt

1035nt 345 a . a . L

Mouse(1647 n t )

173

Fig. 1. Outline of nucleotide sequences of pig, human, rabbit, and mouse DAO cDNAs. Solid boxes, open reading frame (ORF); solid circles, polyadenylation signals; open circle in pig cDNA, the signal that has no corresponding mRNA; UT, untraslated regions; nt, nucleotide; a.a., amino acid; A, G, and C, nucleotides at the position -3; M and L, Met and Leu; and open circles in mouse cDNA, missing residues. Comparison of animal DAO cDNAs The outline of nulcleotide sequences of pig, human, rabbit, and mouse DAO cDNAs is shown in Fig. 1. The open reading frames of pig, human, and rabbit DAO cDNA consisted of 1041 nucleotides encoding

347

amino

acids.

On

the

other

hand,

the

protein

coding region of mouse DAO cDNA consisted of 1035 nucleotides encoding 345 amino acids. The cDNAs contained polyadenylation signals

in

the

3'-untranslated

regions.

By

Northern

blot

analysis, three sizes of DAO mRNA were detectable in pig and rabbit

kidneys,

and

only

a

single

DAO mRNA

of

about

2-kb

existed in human and mouse DAO mRNA. DAO mRNA corresponding to the

second

circle) was

polyadenylation

signal

in

pig

not found. The 5'-untranslated

DAO

cDNA

(open

region of rabbit

DAO cDNA was more than 3 times longer than that of pig DAO cDNA, and in addition, the nucleotide at the position -3 was cytosine

nucleotide,

whereas

it

was

adenine

or

guanine

137

nucleotide in the others. The long 51-untranslated region and the deviation from Kozak's rule (9) in rabbit DAO cDNA was shown to be related with translational regulation from the fact that the removal of the 51-untranslated region up to 10 bases by Nael digestion and replacement of the cytosine nucleotide with guanine nucleotide induced the synthesis of rabbit DAO in an in vitro expression system, although the enzyme was not synthesized without such processing and mutation (3) . Comparison of primary structure of animal DAOs Amino acid sequences of the four animal DAOs predicted from the nucleotide sequences of the DAO cDNAs were compared with each other. The result is shown in Fig. 2. The highest sequence homology was obtained between pig and mouse DAOs, when gaps were made in the amino acid sequence of mouse DAO at the position 25 and 172. The sequences of human, rabbit, and mouse DAOs were highly homologous to that of pig DAO, and the identity was 84%, 80%, and 77%, respectively. Five cyteine residues were all conserved, and 18 proline residues out of 21-23 were also conserved. The sequence at the positions 7-12, which is predicted as FAD binding site (10) and the carboxylterminal sequence of Ser-His-Leu, which is considered to be a signal for translocation into peroxisomes (11), were conserved among the four animal DAOs. The initiation codons were all methionine, and the first 17 amino acids from the aminoterminus were almost identical and hydrophobic, indicating that the animal DAOs are synthesized as the mature form without processing. Prediction of secondary strucuture of animal DAOs The secondary structure of the four animal DAOs were analysed by the method of Chou and Fasman (12). The predicted secondary structures were also highly homologous. Moreover, the random coil regions were mainly localized in the carboxyl-terminal half of the sequence and scarcely existed at the positions

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O 3 3 3 3 H >H>H >. CJ CJCJ CJ cj u CJ CJ X X X X

fe,fe, fe.

o CJ CJ CJCJ O CJ CJCJ CN J •4 k4 J Eh Eh Eh Eh > > > > < Eh < EH CU O X se J Eh > CO CJ (J U CJ CU eu ex, eu o M M M M ro M M hH M H >• >< >h CU eu eu & co CO CO CO z Z X z >H >H >H >H M M M M cj u cj CJ ce ce CO •4

O CJ CJ CJ CJ >H >H ro z z z z X X X X M HM M > > > > ed w 6] fei Eh EhEh < z z X co co co co co

o u u u co CN j eu Ch cu (N Q Q a Q X X X X Eh EH EH Eh M 1-4 (-H J M M M t-H lu Z X X X ss se se ss

o ce ex ce ex cr> > M M > CN o o a o CU CUcu cu ex ex ce oc, > > > > CU CUcu cu ce ce ce ce

EH M Cl] CQ co CQ D < O ce E

EH z M [2] < CQ CO CJ E CQ 3 M D < O CU X ex e

O co cu cu co O CJ CJCJ CJ m fe, Eh ce ce co ce «-4 J O O Oí 3 u u w w ce ce ce ce w w CJ w j j •4 J

< fe.

(tì

XI -a -p e •H M (0 43 aj 43 T3 (d c — M .-i ». fi .—. H O »1 e -h o — g 0 .e a; w - +J -H Di-M X h in o a m oí a) 4-1 c a O H V in c a) -H U X! c a) a d < cted 0) in 13 m aj o -a -p •HOC Ü -H o d 13 -H a) n ra -a a cj H o

fe, fe, fe. fe fe, fe, fe z < CJ E M D Ou X

CJ CJCJ CJ

CN • tji •H

^ -P -H a) lo Oth i l Çh O -H e

139 amino acid sequence N 1 u.

(P)

100

I

III

(H)

i

l u l l

(R)

i

•

(M)

i

•

(P) (H) (R) (M)

AA_

200

I I I• I i i

C 347

300

•

I I•

i • m i

I I i I

I i

m • i I I

i •

I

I

• i

218-264 region ( V(228)

, 3612.

1987.

2. Miyake, Y., K. Fukui, K. Momoi, F. Watanabe, T. Shibata. 1987. In: Flavins and Flavoproteins (D.E. E d m o n d s o n and D.B. McCormick, eds.). Walter de Gruyter, Berlin-New York, p. 501. 3. Momoi, K., K. Fukui, F. Watanabe, Y. Miyake. Lett. 238, 180.

1988.

FEBS

EXPRESSION O F MOUSE KIDNEY D-AMINO ACID OXIDASE IN ESCHERICHIA COLI: PURIFICATION AND CHARACTERIZATION O F THE RECOMBINANT PROTEIN

Masazumi Tada, Kiyoshi Fukui, Motoshige Miyano and Yoshihiro Miyake Department of Biochemistry, National Cardiovascular Center Research Institute, Fujishiro-dai, Suita, Osaka 565, JAPAN

Introduction Significant

variations

in

the

activities

of

D-amino

oxidase

(EC 1.4.3.3, DAO) in kidney and liver among

animal

species

biochemical

have

been

activities are unknown. the

reported

events which produce

question,

we

(1).

However,

the differences

In this paper,

expressed

an

in

in order to

active

acid

various

mouse

the

tissue address

enzyme

in

Escherichia coli cells w i t h the use of an isolated cDNA clone for DAO from a mouse kidney cDNA library

(2).

Purification

of the synthesized mouse enzyme was performed to characterize its enzymatic properties.

Results and Discussion For efficient synthesis of mouse kidney DAO in E^ coli, expression

system,

in

which

the

strong

tac

combined with the high copy plasmid, pUC19, w i t h some modification. introduced

into

ribosome-binding synthetic

introducing

Initially,

EcoRI

sequence

site of

oligonucleotide

PUCES223-3S. modified

the

Then,

to a

the

remove SphI

site

at

SphI

downstream

pKK223-3

by

using

(5'-AATTCGCATGCG-3 1 ),

mouse the

promoter

(3) was

a unique

just

DAO

cDNA,

pMDA015,

5 ' -untranslated the

initiation

the

site

was

from

the

a

12

bp

yielding was

also

sequence

codon

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

is

employed

using

by in

148

vitro mutagenesis (2). Finally, the Sphl-EcoRV site of the modified cDNA was inserted into the Sphl-Smal site of pUCES223-3S. The resultant plasmid was designated as pESMDl (Fig.l). Western blotting analysis with the use of an anti-porcine DAO antiserum (Fig.2A) revealed that a single polypeptide band was observed for HB101 cells harboring pESMDl, but not for those harboring a plasmid without the cDNA insert. The mobility of the immunoreactive recombinant mouse protein was identical to that of the band detected in the mouse kidney extract, demonstrating synthesis of the appropriately sized protein product. For the purification of mouse DAO synthesized in coli, a cell pellet (6 g) from 1 L of culture was extracted, and then fractionated with ammonium sulfate at 35% saturation. Following dialysis against 50 mM sodium phosphate, pH 6.3, containing 40 pM FAD, 1 mM sodium benzoate and 5 uM leupeptin (buffer A), the solution was applied to a hydroxylapatite column that had been equilibrated with buffer A excluding FAD. The adsorbed protein was eluted with 200 mM sodium phosphate, pH 6.3, containing sodium benzoate and leupeptin. The eluate was dialyzed against 20 mM Tris-HCl, pH 8.0, containing 40 viM FAD, 1 mM sodium benzoate and 5 uM leupeptin and then loaded onto a DEAE sepharose column that had been equilibrated with 20 mM Tris-HCl, pH 8.0, containing 5 uM leupeptin (buffer B). The adsorbed protein was eluted with a linear salt gradient of KC1 (0-0.2 M) in Buffer B. The fractions were assayed by both immunoreactivity of DAO with the anti-porcine DAO antiserum and DAO activity, and the peak fractions were collected to use as the purified mouse DAO. In a typical experiment, 0.5 mg of mouse DAO was obtained from 1 L of culture. SDS-polyacrylamide gel electrophoresis of the purified preparation revealed a single protein band with an estimated molecular weight of 36 kD, which is slightly smaller than that of the porcine kidney enzyme (38 kD)(Fig. 2B) . Amino

Figure 1. Schematic diagram of an expression plasmid pESMDl for mouse DAO. The closed and open arrows represent the tac promoter and rrnB terminators, respectively. The closed and open boxes correspond to the coding and 3'-untranslated sequences of mouse DAO cDNA, respectively. Regions a, b and c were derived from pMDA015 (mouse DAO cDNA), pKK223-3 and pUC19, respectively.

A

1 2

3 4

B

1 2 94 -

mm

75-

6 7 -

•

50-

43

-

30

-

20.1

-

27-

,-3 Mr x 10'

3

Mr x 10.-3

Figure 2. Expression and purification of mouse DAO synthesized in E^ coli. (A) Western blot analysis. A cell-free extract was prepared from E^ coli HB101 harboring PUCES223-3S (lane 2) or pESMDl (lane 3). Aliquots of cell-free extract (2.5 y.1) and a mouse kidney extract (125 ug)(lane 4) were subjected to 10% SDS-PAGE. DAO purified from porcine kidney (80 ng)(lane 1) was used as a standard. (B) SDS-PAGE. Two ug each of porcine kidney DAO (lane 2) and the purified mouse DAO (lane 3) were electrophoresed on 10% SDS-polyacrylamide gel and stained with Coomassie Blue. The molecular weight standards (lane 1).

150

acid sequence analysis was performed on a preparation of the purified

protein

sequenator.

using

an

Applied

Biosystems

470A

protein

The amino acid sequence of the amino-terminal 15

residues was completely identical to that predicted from the mouse DAO cDNA (2) . In order to investigate enzymatic properties of the purified mouse

enzyme,

kinetic

spectrophotometric value

of

the

analysis

assay

mouse

method

enzyme

for

was as

performed

described

D-alanine

was

sodium

benzoate

competitively,

benzoate was 180 uM. the

mouse

protein) These

enzyme

(9.7

ymol

was comparable

results

substrates

is

show

and

the

On the other hand, D-alanine

to that of

that

the

significantly

lower

Km

Ki

mM.

value

oxidized/min of

In

inhibited for

the Vmax value

the porcine

affinity

a

The

38.6

parallel, the enzymatic activity of mouse DAO was by

using

(4).

mouse

than that of

per

enzyme

mg (3).

enzyme the

of

for

porcine

enzyme, in spite of having a high sequence homology (2) and a very similar molecular properties pattern which

of

of

mouse

size with each other.

DAO

might

tissue-specific

exists

as

a

single

expressed as a single mRNA

be

expression copy

The

related with in

of mouse

mouse

enzymatic

the DAO

genome

unique gene, and

species in kidney and brain

not in liver (2).

References 1. Birkofer, L., R. Wetzel. 1940. Z.Physiol.Chem. 264, 31. 2. Tada, M., K. Fukui, K. Momoi, Y. Miyake. 1990. Gene press) 3. Watanabe, F., K. Fukui, K. Momoi, Y. Miyake. 1989. Biochem.Biophys.Res.Commun. 165, 1422. 4. Fukui, K., K. Momoi, F. Watanabe, Y. Miyake. 1988. Biochemistry 27, 6693.

(in

is but

PURIFICATION O F RECOMBINANT MUTANT PORCINE D-AMINO ACID OXIDASES

Motoshige Miyano, Kiyoshi Fukui, Fusao Watanabe, Tada, Saori Takahashi and Yoshihiro Miyake

Masazumi

Department of Biochemistry, National Cardiovascular Center, Research Institute, Fujishiro-dai, Suita, Osaka 565, Japan

Introduction Significant data on the reaction mechanism of D-amino oxidase

(DAO)

have

unsettled the

been

role

of

catalytic reaction. we

constructed

contain

a

amino

it

However,

acid

residues

is

in the

In order to investigate this problem,

expression

cDNA

accumulated.

acid

fragment

plasmids encoding

for

mutant

mutant

Phe(228) and Leu(307), respectively.

DAOs

DAOs

which

possessing

The mutant DAOs were

then synthesized in E ^ coli cells and purified from the cell extracts.

Moreover,

the

enzymatic

properties

were

analyzed.

Results and Discussion The cDNAs and the expression plasmids for the mutant Phe-228

and

Leu-307

mutants,

procedures as described coli HB101.

were

constructed

by

harvested

The mutant DAOs were purified from a

hydroxylapatite

column chromatography.

and

a

the

DEAE-Sepharose

The Phe-228 and Leu-307 mutants were

purified as the holoenzyme (Figure 1) ;

the

(1,2) and were used to transform E.

culture medium, and about 80 g of cell paste was extract

by

The transformed E^ coli was grown in 12 L of

by centrifugation. cell

DAOs,

and the apoenzyme,

respectively

about 10 mg of the mutant enzyme was

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

152

obtained from 80 g of each cell paste. gel

electrophoresis

corresponding

to

showed

the

a

single

molecular

Analysis of the amino-terminal sequences of the first terminus

of

the

size

SDS-polyacrylamide

band of

at

pig

sequences

che DAO

position (38

revealed that

17 and 13 residues from the

Phe(228)

and

Leu(307)

identical with that of the native DAO.

kD).

were

the

amino-

completely

The enzyme reactions

of the Phe-228 and Leu-307 mutants were analyzed kinetically using D-alanine as a substrate.

The Km values of the Phe-228

and Leu-307 mutants for D-alanine were 156 and 22.2 mM,

and

the Vmax values were about 66 % and 58 % that of the native DAO,

respectively.

Both

mutant

DAOs

were

inhibited

by

benzoate competitively, and the inhibitory effect of benzoate for the Phe-228 mutant was remarkably less than that for the native DAO, and it was moderate for the Leu-307 mutant. Ki values

of

the Phe-228

and Leu-307 mutants

were 10.7 and 0.16 mM, respectively.

for

The

benzoate

As the Leu-307

mutant

was purified as the apoenzyme, the affinity of FAD for two mutants was investigated kinetically.

the

The K d value

the Leu-307 mutant for FAD was 10 pM, whereas

of

the value of

the Phe-228 mutant was comparable to that of the native DAO. The

reactions

of

the

Phe-228

D-propargylglycine(D-PG)

were

and

Leu-307

analyzed

mutants

whether

or

not

with the

mutant DAOs are inactivated by this suicide substrate.

The

wild type DAO was rapidly inactivated as reported with

the

native DAO(4,5). the

Phe-228

However, no inactivation was observed w i t h

mutant.

Moreover,

the

Leu-307

mutant

inactivated with a slower rate than the wild type DAO.

was At 50

mM D-PG, half time for inactivation at 25°C was estimated to be

1.6

Leu-307

min

with

mutant.

the The

wild

type

absorption

DAO

and

6.4

spectrum

min

of

the

mutant differed from that of the w i l d type DAO Namely,

the

two peaks

are blue-shifted

peaks of the wild type DAO.

from

with

the

Phe-228

(Figure

1).

corresponding

The absorption spectrum of the

Phe(228) mutant was also measured in the presence of several

153 Figure 1: Absorption spectra of the purified recombinant DAOs in 50 mM pyrophosphate, pH 8.3 at 15°C. 1; wild type DAO, 2; Phe-228, 3;Leu-307.

300

400

500

600

Wavelength (nm)

Figure 2: Absorption spectra of oxidized and reduced forms of the wild type and Phe-228 DAOs a, wild type DAO; b, Phe-228 mutant; 1 and 3, the oxidized forms; 2 and 4, the reduced forms. The enzymes were reduced with 20 mM D-alanine.

Wavelength (nm)

154

concentrations mutant

of

tended

benzoate.

to

convert

The

to that

spectrum of

of

the

Phe-228

the benzoate

complex

form on increasing the concentration of benzoate. the dissociation

constant of

benzoate was

estimated to

2.9 mM, which is 967-fold greater than that of DA0(5).

the

Irrespective of the great changes in the

properties, converted

to

the

absorption

that

of

the

spectrum fully

of

Phe-228

reduced

However,

form

be

native

enzymatic

mutant with

20

was mM

D-alanine, and the spectrum of the reduced form was the same as that of the reduced form of the wild type DAO (Figure 2).

Acknowledgement This research was

supported in part by a Grant-in-Aid

for

Scientific Research (63570137 and 02857040) from the ministry of Education, Science and Culture of Japan.

References 1. Watanabe, F., K. Fukui, K. Momoi, Y. Miyake 1989. J.Biochem. 105, 1024. 2. Watanabe, F., K. Fukui, K. Momoi, Y. Miyake 1989. Biochem.Biophys.Res.Commun. 165, 1422. 3. Horiike, K., Y. Nishina, Y. Miyake, T. Yamano 1975. J.Biochem. 78, 57. 4. Marcotte, R., C.T. Walsh 1978. Biochemistry 17, 2864. 5. Massey, V., H. Ganther 1965. Biochemistry 4, 1161.

LIMITED

PROTEOLYSIS

STUDIES

ON THE APO-

, HOLO-,

BENZOATE FORMS OF PIG KIDNEY D-AMINO ACID

HOLOENZYME-

OXIDASE.

G i o v a n n a T o r r i T a r e l l i , M a r i a A. V a n o n i , a n d B r u n o

Curti

Dipartimento di Fisiologia e Biochimica Generali, d e g l i S t u d i di M i l a n o , M i l a n o , Italy

Università'

Armando

Negri

Istituto di Fisiologia Veterinaria e d e g l i S t u d i di M i l a n o , M i l a n o , Italy

Biochimica,

Università

Introduction The

kinetic

DAAO)

properties

from

primary

pig

structure

structural localization catalysis. enzyme

of

the

have

of

several

been

protein

been

no

available,

D-amino

have

gene

However,

is

of

kidney

yet

acid

oxidase

and

the

acid

along

residues

three-dimensional (1) .

In

order

the

three

enzyme

forms,

complex,

holoenzyme-benzoate

i.e.

limited

the

of

its

with

the

implicated

structure to

information on the different conformational with

1.4.3.3,

studied;

sequence

determined

amino

(EC

extensively

gain

associated

holo-,

proteolysis

have been carried out on the three enzyme species

the

structural

states

apo-,

of

in

and

experiments (2).

Results The apo-, D-amino

holo-,

acid

and holoenzyme-benzoate

oxidase

(1

mg/ml)

were

forms

of p i g

incubated

with

t r y p s i n o r c h y m o t r y p s i n for d i f f e r e n t t i m e s in 50 m M buffer,

pH

8.0,

at

u n s e n s i t i v e t o 10 %

25°C.

(w/w)

The

enzyme

was

chymotrypsin, while

in

kidney either

Hepes/K+

all

incubation

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

cases with

156

TT

1.2 Iflfe

ft

& | — S — I

*

*

—i

0.8

0.4 0.0

60

Time (min)

180

120

Figure 1. Limited proteolysis of D-amino acid oxidase. Apo( O,0) , holo- ( • , • ), and holoenzyme-benzoate complex (A , • ) of D-amino acid oxidase were incubated with 10% trypsin (w/w), at 25°C. At different times aliquots were withdrawn for activity measurements and analysis of the densitometric pattern after SDS-PAGE, and Coomassie Blue staining. The fraction of residual activity (open symbols) and the fraction of the density corresponding to the 39-kDa and 37-kDa protein species over the total protein present (closed symbols) are reported as a function of the incubation time. 10 % (w/w) trypsin, under non-denaturating conditions, caused enzyme inactivation which were different for the three enzyme forms of DAAO

(Figure 1). The proteolysis patterns

shown) were also different when apo-, holo-, zyme-benzoate

complex

Particularly,

tryptic

form

of

D-amino

activity

despite

yield

a stable

of

digestion

acid the

DAAO were

oxidase native

or the

incubated

of

led

the to

39-kDa

37-kDa polypeptide.

no

(data not

with

holoentrypsin.

holoenzyme-benzoate loss

protein

of

was

Incubation

catalytic cleaved

of the

to

apoen-

zyme with trypsin led to monophasic activity loss, while the native

polypeptide

species, and

which

13.4-kDa

inactivation presence 80%

was

of

of the

was

first

further

polypeptides. and

two

the

In

degradation

distinct

total

cleaved

degraded

protein

the

37-kDa

active

unstable

25-kDa

holoenzyme, patterns

populations present

to

to the

of

was

both

indicated

protein rapidly

the the

molecules: inactivated

157 1 M—K—Y

100 M—K

m m fnm

e r-

1 M

200 K—HYY

ae ui

a o

1 M

25kDa

1 M

20kDa

r»»co n n

328 K

37kOa

300

r» e

329 I

221 R

222 G

347 L

2kDa

347 L

14.5kDa

222 G

1

347 SHL

H

13.4kDa---|

Figure 2. Products of limited proteolysis of D-amino acid oxidase. The N-terminal sequences of the products obtained by limited tryptic digestion of apo-, holo-, and holoenzymebenzoate complex of D-amino acid oxidase are shown in the figure. The upper line is the schematic representation of the native 39-kDa polypeptide, where amino acid residues that have been chemically modified are indicated. For clarity, the relative sizes of the proteolytic fragments have not been taken into account. concomitant

to

its

degradation

14.5-kDa polypeptides,

the

to

the

latter being

unstable

25-kDa

further

degraded

and to

the 13.4-kDa species. The residual 20% of the protein molecules were major

slowly

degraded

proteolysis

to the active

products

obtained

37-kDa

upon

species.

incubation

three forms of D-amino acid oxidase with trypsin were ted by either SDS-electrophoresis or gel filtration

The

of

the

isola-

cromato-

graphy, and further analyzed. Their N-terminal sequences were determined, thus allowing us to identify the sites of tryptic cleavage

(Figure 2). The stable 37.0-kDa polypeptide resulted

from a cleavage of the native protein at Lys-328, which also yielded

a

C-terminal

2.0-kDa

fragment.

The

unstable

25-kDa

and 14.5-kDa or 13.4-kDa polypeptides obtained directly

from

the holoenzyme or from the cleavage of the 37-kDa polypeptide derived

from

the

apoenzyme,

depend

on tryptic

digestion

at

Arg-221. These

results

experiments changes,

confirm

indicating

which

occur

previous the

upon

spectroscopic

existence

binding

of

of

and

kinetic

conformational

coenzyme

to

the

apo-

158 protein,

and

the primary

of

benzoate

site

of

to

tryptic

holoenzyme cleavage

(l) .

Interestingly,

in holoenzyme

is

Arg-

221. This residue is located within a cluster of amino (residues 204-228) which have been implicated in the active site. Furthermore, binding

of

benzoate,

stabilizes

a

presumably

the

a

protein

it is worthwhile to point out pseudo-substrate, conformation

surrounding

region,

to

the

where

Lys-328

of

the

Arg-221,

is no more

polypeptide

apo- and holoenzyme-benzoate retained

full

catalytic

and

accessible of

with

properties

with respect to the corresponding native enzyme forms. results the

are

FAD

enzyme, terminal

in agreement with the tentative

binding

domain

in

the

N-terminal

D-

scale.

activity

unaltered kinetic parameters, and coenzyme binding

to

cleavage

forms

amino acid oxidase was isolated on a semi-preparative This

that

holoenzyme

trypsin. The stable 37-kDa polypeptide derived from at

acids

enzyme's

These

localization region

of

of the

and with the hypothesis that the function of the

C-

region of D-amino acid oxidase could be related

to

the import of this enzyme into the peroxisomes, by Gould et al.

as

suggested

(3).

Acknowledgement This work was supported by MURST and CNR, Roma, Italy.

References

1. Curti, B., S. Ronchi, and M. Pilone Simonetta. 1991. In: Chemistry and Biochemistry of Flavoenzymes. Vol 2 (F. Muller, ed.) CRC Press, Ine, Boca Raton , in press 2. Torri Tarelli, G., M. A. Vanoni, A. Negri, and B. 1990. J. Biol. Chem. in press 3. Gould, S. , J. , G. A. Cell. Biol. 107. 897-905

Keller,

and

S.

Subramani.

Curti.

1988.

J.

REINVESTIGATION OF THE ROLE OF LYSINE RESIDUES IN D-AMINO ACID OXIDASE

T. Simonie, L. Mannucci, A. Negri, G. Tedeschi and S. Ronchi Istituto di Fisiologia Veterinaria e Biochimica Università' di Milano, Milano, Italy

Introduction D-amino acid oxidase

(DAAO) from pig kidney

is considered

a

model protein for the oxidase class of flavoenzymes. However, the lack of a known tertiary structure has hampered a detailed elucidation of its active site topology. Furthermore, pancies

in the conclusions

drawn

from

chemical

descre-

modification

and site-directed mutagenesis studies have been reported. particular chemical modification of DAAO by

In

fluorodinitroben-

zene (1) suggested that Lys 204 could be involved in the substrate binding. In contrast mutation of Lys 204 to Glu produces a fully active enzyme (2). In the light of these contrasting results we decided to reinvestigate the role of lysine residues

in DAAO using

a lysine-specific

reagent, pyridoxal-5'-phosphate

chemical

modifying

(PLP), which has been shown to

inhibit apo-DAAO in a previous study by Miyake and Yamano (3). In this report we extend the investigation of the effect of PLP on the holo- and the holo-benzoate complex forms.

Results The kinetic of inhibition of apo-DAAO modified

by

different

concentrations of PLP reflects the one previously reported by Miyake et al.(3). The inhibition of holo-DAAO follows the same pattern, although the inhibition rate is slower. On the contrary the holo-benzoate complex is not significantly inhibited

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

160 b y P L P . D e t e r m i n a t i o n of t h e k i n e t i c p a r a m e t e r s

for the

f o r m s of e n z y m e m o d i f i e d w i t h 8.75 m M P L P for 30 m i n , by

reduction

with

phenylglycine but

a

and

decrease

NaBH4, Ki

for

in t h e

shows the

no

changes

competitive

turnover

in

three

followed

Km

inhibitor

for

D-

benzoate

number with respect

to

those

f o r t h e n a t i v e e n z y m e . T h e d a t a s u g g e s t a n "all o r n o n e "

type

of

PLP-

inactivation.

modified found Kd

for

The

apo-DAAO,

to be the

5.2xl0

-6

dissociation

determined ,

the

binding

the

coenzyme

DAAO site are

of

still

the

binds

for t h e not

of

FAD

from

was

o n e o r d e r of m a g n i t u d e h i g h e r t h a n

dissociation

The modified

costant

spectrophotometrically, coenzyme

benzoate,

anionic

moyety

significantly

from

native

suggesting of

affected

that

benzoate by

the

enzyme. both

and

modification

w i t h P L P . I n o r d e r t o identify t h e l y s i n e r e s i d u e s m o d i f i e d P L P in t h e p r i m a r y s t r u c t u r e of t h e e n z y m e w e p e r f o r m e d r e n t i a l H P L C m a p p i n g of t r y p t i c p e p t i d e s o b t a i n e d b y apo-,

holo- and holo-benzoate

complex

previously

for by

diffe-

digesting

modified

30 m i n i n c u b a t i o n w i t h 8.75 m M P L P f o l l o w e d by r e d u c t i o n

by

with

T a b l e 1: S u m m a r y of t h e r e l a t i o s h i p b e t w e e n p e r c e n t r e c o v e r y of the various peptides containing £ - p y r i d o x y l - l y s i n e (see F i g . 1) a n d t h e d e g r e e of i n a c t i v a t i o n of t h e v a r i o u s f o r m s of DAAO after incubation under the same conditons used for the H P L C t r y p t i c d i g e s t m a p p i n g s of Fig. 1. T h e p e r c e n t o f r e c o v e ry f o r t h e l a b e l l e d p e p t i d e s w a s c a l c u l a t e d b y a m i n o a c i d a n a l y s i s of t h e v a r i o u s f r a c t i o n s a f t e r H P L C c h r o m a t o g r a p h y a n d of t h e m a t e r i a l b e f o r e d i g e s t i o n (taken a s 100%). Peptide

% recovery

%

inhibition

APOENZYME 59 59 59 59

36 50 50 69 HOLOENZYME A B C D

L Y S 116 L Y S 163 LYS 33 L Y S 204

44 44 44 44

36 14 50 38

HOLOENZYME-BENZOATE 36 19 50 11

COMPLEX 5 5 5 5

161

Fig. 1 RP-HPLC chromatography o f t r y p t i c d i crests of• 71) PLP'-modif i e d a p o DAAO, ( 2 ) P L P - m o d i f i e d h olp-DAAO , ( 3 ) PLPB CD lodified holo-DAAObenzoate complex. Modif i c a t i o n by PLP o f t h e v a r i o u s DAAO f o r m s was J L ^ J l L performed as d e s c r i b e d in the tejct. Digestions were c a r r i e d out on t h e carboxymethylated mater i a l s using t r y p s i n at 1 : 5 0 r a t i o f o r 4h a t 37* C. HPLC chromatographies B CD A B w e r e p e r f o r m e d on a Bondapack C , R (Waters c 3 , 9 x 3 0 0 mm) -"cblumn, u m sing a l i n e a r gradient ^f> J from 0 t o 50% CHoCN i n 0.1% t r i f l u o r o a c e t i c a c i d in 64 min and monitored at 325 nm o ( w a v e l e n g t h o f maximum c a b s o r b a n c e of (0 £-py r i doxy 1 - l y s i n e ) . Peaks A, B, C and D c o r n responded t o p e p t i d e s c o n t a i n i n g Lys 1.15, Lys o 163, Lys "33 and Lys 204 w labelled by PLP, SI A B CD r e s p e c t i v e l y . A l l peaks n n o t markea by l e t t e r s did not c o n t a i n p r o t e i c m a t e r i a l a s judgea by a mino a c i d a n a l y s i s . The peak marked by an a s t e r i s k contained r e s i d u a l ' ' ' ' I ' ' ' ' I ' I I I I I I I I I I I I I I I I 1 FAD. 10 20 30 40 50 - C (3) « r o u p .

a substrate

site

they

position,

suggest at

imine,

energy

accelerating

act

the

which has have

that

the C ( 3 ) ,

by

only

a

large

a low

af-

atoms

correspond

repulsively

induces

The

steric

the h y d r o g e n which

plane

in the

to

active

s t r a i n or d i s t o r t i o n

the c a t a l y t i c

of

reaction.

References 1. M a s s e y ,

V. , G a n t h e r ,

2. M a s s e y ,

V. , G h i s l a ,

H.

1965.

Biochemistry

S.

1974.

N.Y.Acad.Sci.

3. Y a g i , K . , O k a m u r a , K . , J. B i o c h e m . 66, 581

Naoi,

M.,

Takai,

R.,

6. N i s h i n a , 7. Q u a y .

Miyake,

Y. , S a t o ,

S. , M a s s e y .

Y.

1987,

J.

K. , S h i g a , 1977.

Biochem. K.

1990.

B i o c h e m i s t r y Jjj,

446.

Kotaki.

Y.,

101. J.

1161

227,

A.,

4. N i s h i n a , Y . , M i u r a , R . , T o j o , H., M i y a k e , S h i g a , K. 1986. J. B i o c h e m . 99, 3 2 9 5. M i u r a ,

4,

Watari,

1969. H.,

581

Biochem. 3348

A.

107,

726

D-AMINO ACID OXIDASE EXPRESSED UNDER INDUCTION CONDITIONS IS ENZYMATICALLY ACTIVE IN MICROPEROXISOMES OF Rhodotorula

gracilis

Mirella Pilone Simonetta, Maria-Elisa Perotti and Loredano Pollegioni Department of General Physiology and Biochemistry, Milano, 26 via Celoria, 20133 Milano, Italy

University

of

Introduction D-Amino acid oxidase (EC 1.4.3.3;DAAO) is very widely distributed in various animal tissues, where it is considered a marker enzyme for peroxisomes (1); it is also present in many other organisms. Recently DAAO has been obtained in a homogeneous and stable form from the yeast Rhodotorula:

this

DAAO, the first to be obtained as a pure flavoprotein from a microorganism source (2), is expressed to a high level under conditions of metabolite induction (0.3% total cell protein)(3). Rhodotorula

is an aerobe lipid-storing yeast which shows a

number of peculiar metabolic characteristics; the presence of peroxisomes in this interesting organism has been either ruled out or not unequivocally proved (4).

Results and Discussion We have performed a study on the presence of DAAO in Rhodotorula

cells combining biochemical, cytochemical and

morphometric approaches. Specific affinity-purified antibodies against pure DAAO were prepared: in blot transfer analysis of both the homogeneous enzyme and the crude yeast extract after SDS-PAGE only the 39k band corresponding to the protein monomer specifically reacted, indicating that the yeast enzyme is

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

168 Fig. 1. Immunoblotting of Rhodotorula crude extract and purified DAAO. Total proteins were stained with Amido Black (lanes AC) and DAAO was detected using streptavidin-biotinylated peroxidase complex and DAB as substrate (lanes D-G). Lanes D and E were treated with Accell-purifled IgG diluted 1:100 in blocking solution, lanes F and G were treated with the corresponding preimmune IgG. (A, D and F) 2.5 fig of pure DAAO; (B, E and G) 100 fig of Rhodotorula crude extract; (C) 3 fig/each of Pharmacia standard weight proteins: albumin (67k), catalase (60k), lactate dehydrogenase (36k) and ferritin (18.5k).

A

B

C

D E

F G

synthesized as the mature form (Fig.l). For DAAO localization a method of postembedding immunolabeling with the indirect immunogold technique was worked out on whole cells (the unusual composition of Rhodotorula

cell

wall prevented the obtainement of intact spheroplasts): under enzyme induction conditions, DAAO appears to be localized very specifically in the matrix of peroxisome-like structures, while in non-inducing conditions immunoreactivity was always absent and peroxisomes were seldomly observed (Figs.2 and 3). On the basis of these data, DAAO cannot be regarded as an universal marker for peroxisomes, as generally accepted. The results are in complete agreement with the fact that in Rhodotorula

DAAO is

synthesized only when D-alanine is present as inducer (and highly expressed when D-alanine is the nitrogen source and glucose is the carbon one)(3); in the same growth conditions only a 3-fold increase in specific catalase activity is observed in cultures containing D-alanine compared to these containing ammonium sulfate. By ultrastructural enzyme cytochemistry in these structures catalase activity is also detected and by the same technique the presence of DAAO is confirmed. On the basis of the biochemical and cytochemical data and of their size, the DAAO containing organelles of Rhodotorula

can be regarded as microperoxisomes.

169

Figs. 2 and 3. Rhodotorula grown on inducing medium (2) and on standard medium (3) after 0.5% glutaraldehyde fixation, embedding in LR White and inununolabeling for OAAO with the indirect immunogold technique. Cells grown on inducing medium are positive to (DAAO) immunolabelling, which is confined to microperoxisomes (arrows), as indicated by the distribution of gold granules. Cells grown on standard medium have very few and small microperoxisomes (arrow), which do not contain any DAAO immunoreactivity. Microperoxisomes are also more developed than in cells grown on standard medium. Bars: 0.5 fim.

Morphometric measurements on Rhodotorula

cells grown in

induction conditions revealed that the extra load of peroxisomal protein (DAAO) is not only associated with a moderate increase in organelle size (by 31% with respect to non-induced cells) but also with the biogenesis of new peroxisomes, since a 241% increase of peroxisome volume density was found (Table 1). In yeasts the development and metabolic function of microbodies (peroxisomes, glyoxysomes) depends very much on the growth environment; by changing the composition of the growth medium it is generally possible to induce both proliferation of organelles and the subsequent transport into them of newly synthesized enzymes for the new metabolic roles. It appears that DAAO in Rhodotorula

is largely employed to

utilize D-alanine as nitrogen and/or carbon source in peculiar growth environments; its metabolic functions in this organism

170 Table I. Morphometric analysis of Rhodotoruta cells grown on standard medium and DAAO inducing medium.

Cells on standard mediun

Cells on inducing mediun

Peroxisome V v *

0.3U0.047 (SEM)

1.06±0.110 (SEM)*

Peroxisome size (Mm) **

0.1610.007 (SEM)

0.21+0.008 (SEM)*

Mitochondria V v

11.65+0.060 (SEM)

12.76+0.650 (SEM)

Lipid, glycogen, vacuoles V v ****

13.94+0.910 (SEM)

31.01+1.590 (SEM)*

*

V y = volune fraction occupied by peroxisomes in 100 (un3 of cytoplasm

** ***

size expressed as the diameter of a circle of equivalent area V « volune fraction occupied by mitochondria in 100 jun3 of cytoplasm Increase is significant (P

0.001)

c o u l d p e r h a p s be simpler t h a n those p l a y e d by p e r o x i s o m a l o x i d a s e s in a n i m a l m e t a b o l i s m

(5). DAAO a p p e a r s from our

studies

t o b e s y n t h e s i z e d as the m a t u r e p o l y p e p t i d e n o t u n d e r g o i n g p r o t e o l y t i c a l p r o c e s s i n g after t r a n s l a t i o n in t h e c y t o s o l or d u r i n g its i m p o r t a t i o n into the peroxisomes, as in f a c t o c c u r s for m o s t o t h e r p e r o x i s o m a l proteins. I d e n t i f i c a t i o n of a S K L r e l a t e d p e r o x i s o m a l targeting signal at t h e c a r b o x y t e r m i n u s t h e Rhodotorula

e n z y m e w a s attempted, but t h i s t e r m i n a l

of

portion

of t h e p r o t e i n d i d n o t appear susceptible to d i g e s t i o n by c a r b o x y p e p t i d a s e in our experimental

conditions.

References 1. De Duve,C., P.Baudhuin. 1966. Physiol.Rev. 46, 323. 2. C u r t i , B . , S.Ronchi, M.PiloneSimonetta. 1991. In: C h e m i s t r y a n d B i o c h e m i s t r y of Flavoenzymes, vol.11 (F. Miiller, ed.). C R C P r e s s in press. 3. P i l o n e Simonetta,M., R.Verga, A.Fretta, G . H a n o z e t . J . G e n . M i c r o b i o l . 180. 593.

1989.

4. M a r e s , D . 1982. M y c o p a t h o l o g i a 80, 179. 5. H a m i l t o n , G . A . , M.M.Afeefy, Al-Arab, E.J.Brusch, D . J . B u c k t h a 1 , C . L . B u r n s , R . K . H a r r i s , D.A.Ibrahim, S.G.Kiselica, W . A . L a w , R . P . R y a l l , S.S.Skorczynski, P.P.Venkatesan. 1987. In: P e r o x i s o m e s in Biology and M e d i c i n e (H.D.Fahimi, H . S i e s , eds.) S p r i n g e r - V e r l a g , Berlin, p.223.

ON THE RECOMBINATION PROCESS OF APO-D-AMINO ACID OXIDASE FROM Rhodotorula

gracilis

Paola Casalin, Loredano Pilone Simonetta

Pollegioni,

Bruno

Curti

and

Department of General Physiology and Biochemistry, of Milano, 26 via Celoria, 20133 Milano

Mirella

University

Introduction The preparation of an apoprotein has long been recognized as a tool

to

study

the

interactions

between

protein

and

coenzyme

and to gain further insight on the enzyme catalysis. Recently, in

the

case

extended

to a

apo-f lavoproteins,

the use

investigate devised

of the

of

FAD

analogues

enzyme active

method

for

the

these

site.

as

In

preparation

studies

probes

to we

a

reconstitutable

apoprotein of D-amino acid oxidase from the yeast gracilis.

Rhodotorula

In the present work, we report some of the physico-

chemical properties of this apoprotein along with studies

been

this perspective, of

specific

have

between

reconstituted

holoenzyme

and

comparative

native

D-amino

acid oxidase.

Results and Discussion The apoenzyme of D-amino acid oxidase Rhodotorula

gracilis

(EC 1.4.3.3, DAAO) from

was obtained by dialyzing the holoenzyme

against 2 M KBr in potassium phosphate 0.25 M, pH 7.5, 0.3 mM EDTA,

5

recovery

mM was

2-mercaptoethanol about

and

20%

80%; the apoprotein

glycerol.

The

regained

over

protein 90%

of

the initial specific activity when assayed in the presence of excess FAD.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

172

Table 1: Spectral Properties of Yeast D-Amino Acid Oxidase

A

Native holoenzyme

0.1%

Fluorescence maxima (nm)

Absorbance maxima (nm)

335,530

274,376,455

335,530

274,380,458

335

278

274 nm:2.78

Reconstituted holoenzyme

n.d.

Apoenzyme

278 nm:2.14

The

of

presence

phosphate

concentration was essential

ions

and

glycerol

in order to recover

at

high

a stable

and

reconstitutable apoprotein. The

reconstituted

holoenzyme

showed

an

identical

spectrum in comparison to that of native protein reports

the

spectral

parameters

for

both

the

absorbance

[1]. Table 1 apo-

and

the

holo- forms of yeast DAAO. Interestingly, eluted

as

a

on

gel

single

filtration symmetrical

experiments peak,

the

apoenzyme

corresponding

to

a

molecular weight of 39k, which indicates that the protein was quantitatively present as a monomer; under the same conditions the

reconstituted

holoenzyme

eluted

as

a

dimer

of

79k,

analogously to what observed for the native holoenzyme [2].

40 FAD added

(ul)

Fig. 1 Determination by flavin fluorescence of the dissociation constant for the FADapoprotein complex. 0.25 jiM apoprotein was titrated with 7.95 nK FAD at 18°C, in 250 mM potassium phosphate buffer, 2 mM EDTA, 5 mM 2-ME, 10% glycerol, pH 7.5. Inset: a = ratio of reconstituted holoenzyme to total protein, L p = concentration of added flavin.

173

Titration of apoprotein with increasing amounts of FAD resulted in a complex with a 1:1 stoichiometry and in a full recovery of catalytic activity. The equilibrium binding of FAD to apoprotein was measured by the quenching of FAD fluorescence (Fig.l) [3]: a K d of 2.2x10" 8 M was calculated, a value in good agreement with the K d of 1.8X10-8M determined by differential spectroscopy. Such a low value is in agreement with the observation that the native enzyme can withstand prolonged dialysis without loss of FAD and that in the assay mixture the addition of FAD is not required for maximal activity. The data are also consistent with a 1:1 molar ratio of binding as already observed in the activity titration experiments. The kinetics of the formation of the apoprotein-FAD complex was studied either by following the quenching of protein and flavin fluorescence, or by differential spectroscopy as well as by following the appearance of the catalytic activity. Both absorbance and fluorometric experiments revealed the presence of a two-stage process with a large, rapid quenching in the first phase (koj3S«3.0 min-^), followed by a slow secondary change (k obs «0.10 min -1 ) which represents only 4-6% of the total fluorescence quenching (Fig.2). Kinetic studies on the recovery of catalytic activity following the addition of FAD

u U7 OJ

0

5

10 Time

(min)

15

20

Fig. 2 Quenching of apoprotein fluorescence upon addition of FAD. Changes in protein fluorescence after addition of 1.50 pM purified FAD to 0.03 fiM apoprotein in 250 mM potassium phosphate buffer, 0.3 mM EDTA, 5 mM 2-ME, 20% glycerol, pH 7.5, at 16°C. Excitation wavelength: 285nm; emission wavelength: 335nm.

174

to

the

apoprotein

under

different

experimental

conditions

showed, w i t h i n the dead time of the instrument, no lag in the appearance of maximal activity. Thus,

even

if

for

binding

of

FAD

to

both

process

[4], in R.gracilis

the

yeast

and

apoprotein

pig

kidney

appears

to

enzymes

be

a

the

two-stage

DAAO all the major changes

occurred

in the initial rapid phase. No intermediate species appear be

present

in

the

binding

of

FAD

to

the

yeast

DAAO:

recombination process apparently involves a sequential from monomer+FAD to monomer-FAD and to dimer-FAD 2 ,

but

to the

process in our

conditions a transient monomer-FAD species was not detected.

Acknowledgements This

research

was

supported

by

grants

from

MURST

and

Italy.

References 1. Pilone Simonetta,M., L. Pollegioni, P. Casalin, B.Curti & S. Ronchi.1989. Eur.J.Biochem. 180.199-204 2. Pilone Simonetta,M., M.A. Vanoni & P. Casalin 1987. Biochim.Biophys.Acta 914/136-142 3. Stinson,R.A. & J.J. Holbrook.1973. Biochem.J. 719-728 4. Massey,V. and B. Curti.1966. J.Biol.Chem. 3423

131.

241.3417-

CNR,

FAD ANALOGUES AS ACTIVE SITE PROBES OF Rhodotorula

gracilis

D-AMINO ACID OXIDASE

Loredano Pollegioni, Mirella Pilone Simonetta Department of General Physiology and Biochemistry, University of Milano, 20133 Milano, Italy Sandro Ghisla Fakultät für Biologie, Universität Konstanz, D-7747 Konstanz, FRG

Introduction The wide range of chemical reactivity exhibited by flavoprotein enzymes is determined fundamentally by the nature of the protein moiety. Studies on the interactions of the isoalloxazine nucleus of flavin with the enzyme are therefore important in understanding this reactivity; flavin analogues are extremely useful probes for elucidating structure-function relationships (1, 2) . We have used a number of modified flavins and explored their interactions with the active center of D-amino acid oxidase from the yeast Rhodotorula

gracilis.

The enzyme contains one FAD per protein monomer of 39k and a E274/E455 ratio of 8.2 characterises pure holoenzyme (3). A stable and reconstitutable apoprotein has been recently obtained: the binding constant of FAD was 2.0xl0 - 8 M

(4), one

order of magnitude lower than the corresponding value for pig kidney DAAO.

Results and Discussion 8-SH-FAD was bound by the apoprotein of Rhodotorula

DAAO

and the paraquinoid flavin form was stabilized (Fig.l); K>k2, giving an intrinsic value of pKi of 8.7. The measured value with [a-2H]-alanine is 8.5±0.13, so the intrinsic value of pKi must be at least 8.5. This pKa value of 8.5-8.7 is probably not significantly different from the pKa value of 8.2 found for serine. Therefore, the observed pH dependence for both substrates is due to the requirement that a group on the enzyme with a pKa value of about 8.4±0.2 be deprotonated for activity. Since the pKa value of the amino group of the substrate is not seen, the protonated form must bind to the enzyme.

Figure 3. pH Dependence of the V/K value for D-amino acid oxidase with (I) D-alanine and (n) D-serine as substrates, 25

Tryptophan monooxygenase

Figure 4. pH Dependence of the V/K value for (I) phenylalanine and (n) methionine and (s) the Ki value for indoleacetamide for tryptophan monooxygenase at 25 °C.

The pH dependence of the V/K value

for phenylalanine shows pKa values of 5.6±0.04 and 9.7±0.04, while

that

for methionine

shows only the acidic value

of

196

5.4±0.06. The pKi value for indoleacetamide, a competitive inhibitor versus phenylalanine, shows pKa values of 5.7±0.05 and 10.4±0.05 (Figure 4). The D V/K p h e value is small (1.29±0.21) and pH independent. This establishes that phenylalanine is not sticky, while methionine may be. The measured DV/KPhe value is too small to be the intrinsic value and reduction is irreversible, so there must be a significant internal commitment. An enzymic residue with a pKa value of 5.7 must be deprotonated for reduction by this enzyme.

Acknowledgement s This research was supported in part by funds from the Texas Agricultural Experiment Station and by NIH Biomedical Research Support Grant RR-07090. J.J.E. was supported by the USDA Agricultural Graduate Studies Program.

References 1. Ghisla, S. 1982. In: Flavins and Flavoproteins (V. Massey and C.H. Williams, Jr., eds.). Elsevier, New York. pp. 133-142. 2. Sherry, B., and R. H. Abeles. 1985. Biochemistry 24. 25942605. 3. Hutcheson, S. W., and T. Kosuge. 1985. J. Biol. Chem. 260r 6281-6287. 4. Bright, H. J., and D. J. T. Porter. 1975. In: The Enzymes, 3rd ed., Vol. XII (P. Boyer, ed.) Academic Press, New York, pp. 421-505.

STRUCTURAL AND KINETIC ANALYSIS OF FLAVINE ADENINE DINUCLEOTIDE MODIFICATION IN ALCOHOL OXIDASE FROM METHYLOTROPHIC YEASTS.

Leonid V. Bystrykh Institute of Biochemistry and Physiology of Microorganisms, USSR Acad. Sei., Pushchino, 142292, USSR Richard M. Kellogg, Wim Kruizinga Department of Organic Chemistry, University of Groningen, 9747 AG, Groningen, The Netherlands Lubbert Dijkhuizen, Wim Harder Department of Microbiology, University of Groningen, 9751 NN, Haren, The Netherlands Jacques Vervoort, Willem J.H. van Berkel Department of Biochemistry, Agricultural University, Wageningen, 6703 HA, The Netherlands

Introduction Alcohol oxidase (MOX), a major peroxisomal protein of methanolutilizing yeasts, has been shown to possess two different forms of flavine adenine dinucleotide, natural FAD and so called modified FAD (mFAD). A comparison of homogeneous preparations of MOX purified from different yeast strains revealed significant differences in the ratios of FAD and mFAD assembled into the apoenzyme (1-3). The mechanism of FAD modification and its structure remained to be elucidated. Here we present results on the structure determination of mFAD by and 13C NMR, HPLC and enzymatic analysis.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

198

FMN

Fig.1. Decoupled

n

CNMR

mFMN

spectra of FMN (left) and mFMN (right).

Results *H and 13C NMR analysis of mFAD revealed that the isoalloxazine ring as well as the adenosine part of the molecule were intact. mFAD showed the same number of carbon atoms as FAD and an identical distribution of methyl, secondary, tertiary, and quaternary carbon atoms. Mild shifts were observed in the isoalloxazine and adenine moieties. However, significant differences were observed in the sugar chains. Two dimensional proton-proton spectra (COSY, NOESY, TOCSY) gave complicated pictures both for FAD and mFAD, especially in the sugar region where the overlap of absorptions (7 protons from ribitol and 5 protons from ribose) is severe. On the basis of literature partial assignments (4-6), it was possible to trace and assign both the ribose and ribitol chains in FAD entirely. In mFAD the ribose chain was also followed and assigned. However, the ribitol chain overlapped severely at H2' with the ribose absorptions; moreover, coupling between H2'and H3' virtually disappeared. In order to simplify the structural analysis of mFAD, it was cleaved by phosphodiesterase to mFMN and AMP. The latter compound was proven to be authentic both by enzymatic (with myokinase, pyruvate kinase and lactate dehydrogenase) and HPLC analysis. The 13C spectra of FMN and mFMN (Fig.l) show small but important differences in the sugar regions. The connectivities are identical in both cases. Two interpretations

199

F1 p p m Fig.2. COSY spectra of FMN (left) and mFMN (right). The diastereotopic protons Hla', Hlb' are at lowest field in both spectra. The ordering of absorptions is as given in the text.

seemed plausible at this point: a) the phosphate has migrated on the ribitol course,

chain

also

in

(7) or b) the sugar moiety mFAD)

is

not

ribitol

but

in mFMN

a

(and of

diastereoisomer

thereof. Possibility a) was eliminated on the basis of the NMR

decoupled

spectrum

of

mFMN,

clearly

indicating

phosphate group is bound at the 5' position with a shift almost identical to natural FMN. HPLC that the dephosphorylation product obtained

of

found

that

significantly

the

COSY

(Fig.2).

In

spectra both

of FMN

chemical shifts remained the same: Hlb'

(diastereotopic

protons)

(diastereotopic protons)
v0H H^l r*0H c H^l ^OH

FAD

base

h

V

h2c'I

I

A

base-H

J

y0H

°

ch2o®o(p)o-{ H^I^OH H^ls*0H

mFAD

base -H*

base

References 1. Sherry, B.,

R.H. Abeles. 1985. Biochemistry, 24, 2594-2605.

2. Bystrykh, L.V., V.P. Romanov, J. Steczko, Y.A. Trotsenko. 1989. Biotechnol. Appl. Biochem. 11, 184. 3. van der Klei, I.J., L.V. Bystrykh, W. Harder. 1990. In: Methods in Enzymology, Academic Press Inc., 188, p. 420. 4. Ulrich, E.L., W.M. Westler, J.L. Markley. Lett. 24, 473.

1983.

Tetrahedron

5. Breitmaier, E., W. Voelter. 1972. Eur. J. Biochem. _31' 234. 6. Kainosho, M., Y. Kyogoku. 1972. Biochemistry 11_, 741. 7. Nielsen, P., P. Rauschenbach, A. Bacher. 1984. In: Flavins and Flavoproteins (R.C. Bray, P.C. Engel and S.G. Mayhew eds.). Walter de Gruyter, Berlin, p. 71.

STUDIES ON YEAST-DERIVED AND MUTANT GLUCOSE OXIDASES

Sumita Chakraborty, Vincent Massey Department of Biological Chemistry, University of Michigan Medical School, Ann Arbor, Michigan 48109-0606, U.S.A.

Jennifer Stratton-Thomas, Steven Rosenberg Protos Corporation, Emeryville, California 94608, U.S.A.

Introduction Recently, the gene for glucose oxidase of A. niger has been cloned and expressed in yeast (1). The yeast-derived enzyme has almost the same

specific activity and the same steady

state kinetics as the native enzyme but it appears to be more stable than the native protein. It contains a higher percentage of carbohydrate compared to the native one (60% vs. 16%) and could be deglycosylated upto 60% using Endo-H with little loss of activity. The enzyme contains three cysteine residues; the one at 521 has been changed to serine by site directed mutagenesis. We will report here on some properties of the mutant and the use of 6-azido flavin as an active-site probe of the native enzyme.

Results and Discussion Properties

of 6-Azido-FAD bound Glucose

Oxidase:

Glucose oxidase in which the native flavin is replaced by 6-N3-FAD

(2) has an absorption maximum at 42 6 nm and is very

stable in the absence of light. On irradiating with visible light, there is a color change from yellow to green, with a

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

202

shift in ^- max from 42 6 nm to 438 nm and development of a long wavelength band centered around 610 nm ( Fig.l). The spectral pattern of the photoirradiated product is similar to that of enzyme bound 6-NH2-FAD. The light-irradiated enzyme, however, is completely inactive, whereas 6-NH2-FAD enzyme has ~2% the activity of normal enzyme. Precipitation of the protein with 15% TCA after lightirradiation, showed that ~40% of the total flavin is bound covalently to the protein, probably through a protein residue containing a free amino function. The 6-azido-FAD enzyme can be converted to the 6-amino-FAD form on reduction with glucose. But the elimination of N2 from the reduced enzyme is very slow ( t^/2 ~3hr ) compared to that from the 6-N3-FMN bound to the apoprotein of Old Yellow Enzyme or to that from 6-N3-FAD.D-amino acid oxidase (2). Properties

of the mutant

Glucose Oxidase :

The only cysteine at the C-terminal region of the C O "I yeast-derived glucose oxidase, serine by site-directed mutagenesis

, has been changed to (3). The absorption

spectrum of this mutant has the same characteristic bands at 278, 382 and 452 nm as those of wild type and max yeast-derived enzyme. It has a high catalytic activity

(see

below), and like the wild type enzyme, forms a flavin N(5) adduct with sulfite Kinetic Analysis:

(5).

In a stopped flow enzyme-monitored turnover

experiment, an 8 H.M solution of the mutant glucose oxidase in 0.13 M phosphate, pH 5.6 at 3° C, was reacted with various concentrations of glucose saturated with oxygen, and the change in optical density at 450 nm was recorded with time.

203 0.4 — Before Irradiation -After Irradiation

r^—r 300

400

500

600

700

800

Wavelength ( n m ) Fig.l Effect of light irradiation on 6-azido-FAD bound glucose oxidase 0 . 2 4 -.«-Oxidised Enzyme

0.02 0.16

-

0.08

-

Glucose

O m
2 un(^er anaerobic conditions (data were emitted), if H^O^ acted as an activator, it should be a nonessential activator. Likewise, if H^O^ acted as an electron acceptor, it would show a sequential mechanism. Under the restricted conditions, such that there is no appropriate electron acceptor other than this substance, this might act as an electron acceptor. PMP/PNP oxidase is classified as one of oxidoreductases. However, in the present experiment, we have found that the enzyme can act as peroxidase depending on the conditions used. Recently, some wood rot fungi have been found to secrete the enzyme which is able to decompose lignin in conjunction with H^O^ (9). These enzymes contain a hone prosthetic group as in a peroxidase or catalase. However, it has been established that PMP/PNP oxidase is a simple flavoprotein (3). The enzyme is distinct in that it acts as a peroxidase, in spite of the simple flavoprotein. REFERENCES 1. McCormick, D.B. & A. H. Merrill, Jr. 1980. In: Vitamin B-6 Metabolism and Role in Growth (Tryfiates, G.P. ed). FN Press, Westport, CT, p.1-26. 2. Merrill, A.H., Jr., M. N. Kazarinoff, H. Tsuge, K. Horiike, & D. B. McCormick. 1979. Methods Enzymol. 62, 568-574. 3. Tsuge, H., K. Ozeki, K. Sen-maru & K. Ohashi. 1979. Agric. Biol. Chem. 43, 1801-1807. 4. Tsuge, H., K. Ozeki & K. Ohashi. 1980. Agric. Biol. Chem. 44, 2329-2335. 5. Tsuge, H., K. Itoh, F. Akatsuka, T. Okada & K. Ohashi. 1984. Biochem. Int. 6, 743-749. 6. Tsuge, H., T. Okada, I. Nakane, S. Uchida, R. Sugiyama & K. Ohashi. 1984. In: Flavin and Flavoproteins (R. C. Bray, P. C. Engel & S. G. Mayhew, eds). Walter de Gruyter & Co., Berlin, p.585-588. 7. Suzuki, S. & K. Ueno. 1981. In: Illustrated Laboratory Techniques Series Vol. 1, Anaerobic Bacteria (N. Kosakai. ed). Vol. 1, Igaku-shoin, Tokyo, Japan, p. 17-27. 8. Hildebrandt, A.G., I. Roots, M. Tjoe & G. Heinemeyer. 1978. Methods Enzymol. 52, 342-350. 9. Tien, M. & T. K. Kirk. 1984. Proc. Natl. Acad. Sci., USA. 81, 2280-2284.

Monooxygenases

CHEMICAL FUNCTIONS OF AMINO ACID RESIDUES IN THE ACTIVE SITE OF PARAHYDROXYBENZOATE HYDROXYLASE AS STUDIED BY SITE DIRECTED MUTAGENESIS AND BIOPHYSICAL TECHNIQUES

Barrie Entsch Department of Biochemistry, Microbiology, and Nutrition, University of New England, Annidale, N.S.W. 2351, Australia Bruce A. Palfey, David P. Ballou, and Vincent Massey Department of Biological Chemistry, University of Michigan, Ann Arbor, MI, 48109, U.S.A.

Introduction Parahydroxybenzoate hydroxylase (EC 1.14.13.2) (POBASE) is the prototype for the study of the flavoprotein oxygenases. A large body of mechanistic (1-3) and detailed crystallographic information (4 and its references) has been published for this protein. Although several chemical intermediates in catalysis, involving both the flavin and the substrate, have been characterised, there is still not enough evidence to chemically model the aromatic hydroxylase mechanism. It is clear that the protein is vital to the reaction pathway, but the X-ray structure provides only partial insight into the functions of the protein (15). Some fundamental and puzzling features of the aromatic hydroxylases remain. For example, how does the binding of substrate to the enzyme initiate the exergonic reactions with NADPH and with oxygen? How does the reaction with oxygen lead exclusively to the transiently stabilized flavin C(4a)-hydroperoxide, which then transfers the distal oxygen atom to the substrate to form product? How does the protein retain the substrate in the active site once the flavin has been reduced? The availability of a high-resolution structure (1.9 A ) of the enzyme-substrate complex (4) has provided the opportunity to rationally use site-directed mutagenesis to perturb the active site in order to address these types of questions.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

220 In this report we provide some results with selected mutants of POBASE (made from the gene, pobA from Pseudomonas aeruginosa PAOIC (5)) and discuss the significance of the mechanistic changes caused by the mutations.

Methods A segment of pobA containing the protein code for the enzyme (one polypeptide of 394 amino acids) plus the termination signal for transcription, was incorporated into an expression vector for Escherichia coli containing the lac promoter. A single-stranded form of the gene was also produced in the phage, Ml3. The phage was then used to generate specific mutants of pobA by the method of Kunkel et al. (6). The mutant genes were then incorporated into the vector and expressed in E. coli (which has no natural function for the gene), and the protein was purified by the methods described by Entsch et al.(2,l). The specific mutants studied here are listed in Table 1. Crystals of the enzyme suitable for X-ray analysis were generated by the method of van der Laan (8). Other methods are referred to in Table and Figure legends.

Results and Discussion Crystals of POBASE from P. aeruginosa are isomorphous with the crystal form of enzyme from P. fluorescens. The crystal space group is C222\ and the cell dimensions are a=71.79, b=146.57, and c=88.06 (M.L. Ludwig, pers. commun.). Thus, the published structural coordinates for POBASE from P. fluorescein apply to this isolate of enzyme from P. aeruginosa. There are only two conservative sequence differences between the isolates, and they are at the surface of the enzyme (5). Residues in the active site Figure 1 summarizes some information concerning the protein residues that surround the flavin and substrate in the active site of the enzyme. A number of these residues contain functional groups interacting with the substrate. Arg 214 provides an essential electrostatic attraction for the carboxyl group of the substrate, and ser 212 helps to orient the substrate through its carboxyl group. As they are on the si side of the flavin, neither of these residues

221

Protein residues near the isoalloxazine ring 44-47; Arg-Ala-Gly-Val (si face and under ring) 293-300; Pro-Thr-Gly-Ala-Lys-Gly-Leu-Asn (re face) Protein residues near the substrate 44-47; Arg-Ala-Gly-Val Trp 185 Tyr201 210-214; Leu-Cys-Ser-Gln-Arg (around carboxyl group) Tyr 222 293-296; Pro-Thr-Gly-Ala (around phenolic group) Tyr 385 Figure 1: Above: A space-filling representation of the enzyme-product (3, 4 dihydroxybenzoic acid; 3,4 diOHB) complex taken from the three-dimensional coordinates available from the Brookhaven Data Bank (1PHH), showing the four amino acid residues mutated in this work. The enzyme-substrate complex has a structure similar to this, with the substrate ring rotated by 14". Below: A listing of residues in the immediate environment of flavin and substrate.

222

are close to the vicinity of the hydroxylation processes. The most interesting other residues are three tyrosines. Tyr 222 appears to hydrogen-bond to the carboxyl group of the substrate, and at the same time is almost in van der Waals contact with the benzene ring of the flavin, perhaps helping to close off the reaction site from solvent. Tyr 201 hydrogen-bonds to the phenolic oxygen of the substrate, and with the oxygen of Tyr 385, forms a network of phenolic groups with the substrate. These tyrosines are on the re side of the flavin ring, where all the chemistry occurs, and have direct links to the loop of protein that appears to be involved in positioning NADPH in the active site. Early results with substrate analogues had shown that the 4-hydroxyl group of the substrate was important to the control of the overall reaction (9). Hence, we made conservative, single- residue changes to Tyr 201, 222, and 385. Their functional groups were removed with the smallest possible structural change - by replacement with phenylalanine. The pyrimidine ring of the flavin is enclosed in protein, and has a number of hydrogen-bonds to the backbone chain of the protein. There are no obvious residues with functional groups involved in these interactions, with the exception of Asn 300 (see Fig 1). The side-chain amide group of this residue hydrogen-bonds to the C(2)0 of the flavin ring, and is at the base of a large helix that may also influence the electronic environment of the flavin. These features are expected to affect the reactivity of the flavin in catalysis. We changed Asn 300 to aspartate, to introduce a negative charge next to the flavin. Provided there is no structural rearrangement, this charge may also modify the electronic form of the reduced flavin, which has been assessed as the anion at N-l in the wild type enzyme (10). Physical properties of mutants Wild type and all four mutants listed in Table 1 were expressed in E. coli, purified successfully, and found to be stable. The wild type was found to be indistinguishable from the enzyme isolated from its natural source. The columns of relative activity and percent hydroxylation (Table 1) show that all four mutations have a major effect on the natural function of the enzyme, but do not totally eliminate catalysis. The effects of each mutant are clearly different, and each has the potential to illuminate aspects of the mechanism. It is noteworthy that the least active mutant is the only one that forms product with every turnover of the enzyme. The crystal structure shows potential hydrogen-bonds between the tyrosines and the substrate. Thus, it is surprising that none of the tyrosine mutations significantly change the

223

00 o co o sc c« o c/1 9 u t-l o s

>n ft. 0

< z

03 K O •a •5 es NO

1/-1 vo

s

f»i NO

Ov NO

00

00

o X o & C

Ö

(S

O

Os

"Al »c se c ts ON

r-^ SC Cl

u cd H

£ m

ON

ITI

•o c

Ci

oo oo

o

o

Ci

o (N >o 00

»-H o

o r-

«ri ON

oo OÑ

ON

>o r-

(S

NO

o

1/1

IT) o « r-, I S"- 3

m

>

J=

rr> u

Sft s ofl c •o 98% from the hydroperoxide at a rate of only 1.8

the hydroperoxide must have a potential

half-life of at least 20 s. Surprisingly, the Tyr385Phe will hydroxylate the product of the normal reaction (3,4-dihydroxybenzoate) at a rate similar to that for pOHB. This produced

228 gallic acid, a substance toxic to the cell. The wild type enzyme avoids hydroxylation of the product. Presumably, the Tyr 385 phenolic group prevents the product from binding in a conformation conducive to hydroxylation. The Tyr222Phe mutant contrasts starkly with the other tyrosine mutants. From its position in the active site, this tyrosine could influence solvent access to the flavin N(5) position and the orientation of the substrate ring to the flavin ring. By removing the hydrogen-bond to the substrate in the mutant, a greater freedom of movement of the benzene rings of Phe 222 and substrate could occur. This hypothesis seems to be reflected in the kinetic analysis. The hydroxylation rate is about 15-fold slower and the hydroperoxide decay rate about 20-fold faster than wild type (see Fig. 2). The overall result is to make this mutant a poor hydroxylase, but a very active oxidase. The last mutant of this group, Asn300Asp, can influence hydroxylation only indirectly through the flavin and the protein. This influence is indeed profound. The hydroxylation rate is slowed by nearly 50-fold from that of wild type, but the flavin hydroperoxide completely hydroxylates the substrate. This implies a similar 50-fold slower rate of decomposition of the hydroperoxide. Thus, we conclude that the flavin hydroperoxide is less electrophilic, which could mean that the aspartate negates the dipole effect of helix 10 on the flavin that has been proposed to assist hydroxylation (15). Perhaps at the same time, the aspartate is raising the pKa of the proton on N-5 of the flavin, making the peroxide more stable. The end result is a good hydroxylase, but a poor catalyst! Our knowledge of the oxygen reactions of POBASE, and subsequently, other hydroxylases, has been helped by including azide anion in reaction mixtures (7). It slows down all catalytic reactions following the formation of the flavin hydroperoxide, and incidentally, also strongly inhibits flavin reduction by NADPH. In contrast to all other hydroxylases, the reaction rates of the Asn300Asp mutant are almost unchanged by a high concentration of azide. It is clear that the azide effect on the wild type enzyme is analogous in many respects to the effect of the aspartate group in the active site of the mutant. Azide even lowers the redox potential of the wild type enzyme as does the conversion of Asn 300 to Asp (Table 1). In Fig. 2, an intermediate labelled II is shown in the hydroxylation pathway. This refers to a species of the enzyme observed in catalysis when selected substrates such as 2,4dihydroxybenzoate (2,4-DOHB) are hydroxylated (7). Recent studies of chemical models have suggested that II may be a complex of oxygenated flavin and product radicals formed

229 from intermediate I via k'5 (Fig. 2) (16). Hydrogen atom transfer between these radicals leads to intermediate III via kg. In our earlier studies of this enzyme with the natural substrate (7,17), we never detected species II in the reaction, and had assumed that it must decay too quickly to detect. None of the mutants studied here show any sign of species II in the reaction with pOHB, but do when the analogue, 2,4-DOHB is bound to the enzyme (data not shown). Thus, it seems that the occurrence of II is a function of the substrate structure in the active site. This conclusion has led us to propose that these hydroxylases can function by two alternate mechanisms. With the substrate, pOHB, heterolytic fission of the flavin hydroperoxide occurs (via k5), but with 2,4-DOHB, homolytic fission occurs (via k' 5 , k'g). Multiple pathways of reaction are often observed with hydroperoxides, and in particular, flavin hydroperoxides can react as either electrophilic or nucleophilic reagents (1).

Acknowledgment This research was supported by grants from the U.S. Public Health Service (GM 20877 to D.P.B., GM 11106 to V.M., GM 08270 to B.A.P.), the University of Michigan (Rackham Graduate School), and the Australian Research Council (B.E.).

References 1

Ballou, D.P. 1984. In: Flavins and Flavoproteins (R.C. Bray, P.C. Engel, and S.G. Mayhew, eds.) Walter de Gruyter, Berlin, pp. 605-618.

2.

Entsch, B. 1990. In: Methods in Enzymology, Vol. 188 (M.E. Lidstrom, ed.) Academic Press, Orlando, pp. 138-147.

3.

Massey, V., L.M. Schopfer, and R.F. Anderson. 1988. In: Oxidases and Related Redox Systems (T.E. King, H.S. Mason, and M. Morrison, eds.) Alan R. Liss, New York, pp. 147-166.

4.

Schreuder, H.A., P.A.J. Prick, R.K. Wieringa, G. Vriend, K.S. Wilson, W.G.J. Hoi, and J. Drenth. 1989. J. Mol. Biol. 208, 679-696.

5.

Entsch, B„ Yang Nan, K. Weaich, and K.F. Scott. 1988. Gene 21, 279-291.

6.

Kunkel, T.A., J.D. Roberts, and R.A. Zakour.1987. In: Methods in Enzymology, Vol. 154, Academic Press, Orlando, pp. 367-382.

7.

Entsch, B„ D.P. Ballou, and V. Massey. 1976. J. Biol. Chem. 251. 2550-2563.

230 8.

Van der Laan, J.M. 1986. Doctoral Dissertation, University of Groningen, Netherlands.

9.

Entsch, B„ D.P. Ballou, M. Husain, and V. Massey. 1976. J. Biol. Chem. 251, 73677379.

10. Vervoort, J. 1986. Doctoral Dissertation, Agricultural University of Wageningen, The Netherlands. 11. Massey, V. 1990. This volume. 12. Williamson, G., D.E. Edmondson, and F. Müller. 1988. Biochim. Biophys. Acta 953. 258-262. 13. Schopfer, L.M. and V. Massey. 1980. J. Biol. Chem. 255, 5355- 5363. 14. Van Berkel, WJ.H. and F. Müller. 1989. Eur. J. Biochem. 122, 307-314. 15. Schreuder, H.A., W.G.J. Hoi, and J. Drenth. 1990. Biochemistry 22, 3101-3108. 16. Anderson, R.F., K.B. Patel and M.R.L. Stratford. 1987. J. Biol. Chem. 262, 1747517479. 17 Entsch,B. and D. P. Ballou 1989. Biochim. Biophys. Acta 222, 313-322.

SITE-DIRECTED MUTAGENESIS OF PARA-HYDROXYBENZOATE HYDROXYLASE FROM PSEUDOMONAS FLUORESCENS.

A.H. Westphal, K. Eschrich, W.M.A.M. van Dongen, J.A.E. Benen, A. de Kok, W.J.H. van Berkel. Department of Biochemistry, Agricultural University, Wageningen, The Netherlands.

Introduction p-Hydroxybenzoate hydroxylase from Pseudomonas fluoresceins is the model enzyme among the FAD-containing aromatic hydroxylases. The reaction pathway has been established in detail (1) and X-ray diffraction studies have resulted in high resolution three-dimensional models of the enzymesubstrate and enzyme-product complexes (2). The exact mechanism of hydroxylation however is still unclear (3,4) . We report here on the cloning and sequencing of the gene for p-hydroxybenzoate hydroxylase from P.fluorescens. Furthermore, the construction is described of mutant enzymes that might be helpful in further elucidation of the catalytic mechanism. A detailed description of the cloning and sequence determination will be published elsewhere (5). Results P.fluorescens chromosomal DNA was partially digested with Sau3A and DNA fragments of 4-6 kbp were ligated into the BamH.1 site of pUC9. After transformation of E.coli TG2 cells with the recombinant plasmids colonies were screened with antiserum against p-hydroxybenzoate hydroxylase for the production of the enzyme. From one positive clone a 7.1-kbp plasmid (pAW44) was isolated. Digestion of the 4.6-

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

232

kbp insert by £coRl and Cla1 and subcloning yielded a plasmid (pAW45) with an 1.65-kbp insert containing the complete gene encoding p-hydroxybenzoate hydroxylase. After ligation of this insert into M13mpl8 the DNA sequence was obtained by the dideoxy chain-terminating method of Sanger et ai.(6).The derived primary structure agreed completely with the chemically determined amino acid sequence (7). The homology of the nucleotide sequence with the DNA sequence of the gene encoding p-hydroxybenzoate hydroxylase from P.aeruginosa (8) is very high. In the coding region 14 nucleotides are different resulting in only two amino acid substitutions at positions 228 and 24 9. The expression of p-hydroxybenzoate hydroxylase in E.coli TG2 cells under influence of the vector encoded lacZ promoter is five times higher (about 10% of cellular protein) as found for the expression in E.coli of the gene from P.aeruginosa (8). The enzyme, fully saturated with FAD, was purified by a simple three-step procedure (Table 1).

Table 1.

Step

Purification of p-hydroxybenzoate hydroxylase from E.coli TG2 (pAW45) (30 g wet cells). Protein

Activity

Specific Activity

Yield

mg

U

U/mg

%

Cell-free 2836 extract Protamine 1001 sulfate Cibacron-blue 250 agarose 178 DEAE-Sepharose

13314

4.7

100

10712

10.7

81

9511

38.0

71

8920

50 .1

67

233 M13mpl8 DNA containing the 1.65 kbp Pstl-Clal fragment described above was used for oligonucleotide directed in vitro mutagenesis according to the method of Kunkel (9). Two types of mutated enzymes have been constructed until now. In the first type, cys-116 was replaced by a serine residue. Cys-116, located near the enzyme surface, is slowly oxidized in air, resulting in a variety of oxidation products (10). It has been postulated that these oxidation products hamper the crystallization of the enzyme (11). In contrast to wild-type enzyme (10), C116S was resistant to oxidation by hydrogen peroxide. The second type of mutations concerns amino acid residues possibly directly involved in activation of the substrate. Tyr-201 and Tyr-385, both located close to the hydroxyl moiety of the substrate (2), were replaced by phenylalanine residues. Preliminary results indicate that the mutations strongly decrease the overall reaction rate without dramatically affecting the binding of ligands to the oxidized enzyme (Table 2). Table 2.

Some properties of p-hydroxybenzoate hydroxylase wild-type and mutant enzymes.

Enzyme

Specific activity

Kd substrate

Kd product

Kd NADPH

pH 8.0

pH 7.0

pH 7.0

pH 6.4

U/mg

jlM

¡Im

¡IM

wt

50

33

286

232

C116S

50

32

290

237

Y201F

1

34

341

248

Y385F

1

12

178

136

234

Acknowledgement This research was supported in part by the Netherlands Foundation of Chemical Research (S.O.N.) with financial aid from the Netherlands Foundation for Scientific Research (N.W.O.). K.Eschrich was supported by a grant of the Program Bureau for Biotechnology. References 1.

Husain, M. and V. Massey. 1979. J.Biol.Chem. 254, 6657-6666.

2.

Schreuder, H.A., P. Prick, R.K. Wierenga, G. Vriend, K. Wilson, W.G.J. Hol, J. Drenth. 1989. J.Mol.Biol. 2ÜR, 679-696.

3.

Van Berkel, W.J.H. and F. Müller. 1989. Eur.J.Biochem. 179, 307-314.

4.

Entsch, B. and D.P.Ballou. 1989. Biochim.Biophys.Acta 999 r 313-322.

5.

Westphal, A.H., J.A.E. Benen, W.J.H. van Berkel, W.M.A.M. van Dongen, A. de Kok. 1990. Eur.J.Biochem., submitted.

6.

Sanger, F., S. Nicklen, A.R. Coulson. 1977. Proc. Natl. Acad. Sei. USA JA, 5463-5467.

7.

Wijnands, R.A., W.J. Weijer, F. Müller, P.A. Jekel, W.J.H. van Berkel, J.J. Beintema. 1986. Biochemistry 4211-4218.

8.

Entsch, B., N. Yang, K. Weaich, K.F. Scott. 1988. Gene 71, 279-291.

9.

Kunkel, T.A., J.D. Roberts, R.A. Zakour. 1987. In: Methods in Enzymology, Vol. I M , pp. 367-382.

10.

Van Berkel, W.J.H. and F. Müller. 1987. Eur.J.Biochem. 1£2, 35-46.

11.

Van der Laan, J.M., M.B.A. Swarte, H. Groendijk, W.G.J. Hol, J. Drenth. 1989. Eur.J.Biochem. 179, 715724 .

PARTIAL

SEQUENCES

OF

PHENOL

HYDROXYLASE. AND

SIMILARITIES

WITH OTHER FLAVOENZYMES Halina Y. Neujahr and Torsten Sejlitz Dept. of Biochemistry and S-10044 Stockholm, Sweden

Biotechnology,

Royal

Institute

of

Technology,

Introduction The

flavoprotein phenol

oxygen-dependent group

of

hydroxylate

substrate

in

(EC

ortho-hydroxylation

"external

hydroxylases",(1,2). can

hydroxylase flavin

Phenol

of

(3,4)

catalyzes

phenol(s),

which

monooxygenases",

hydroxylase is the only

unsubstituted

common

1.14.13.7)

phenol, with

its

the

main

most

the

NADPH-

places

called

it

also

in

studied

It

aromatic

has

the

"aromatic

flavin monooxygenase,

substrate.

and

no

which

phenolic

hydroxylases

(2).

Phenol hydroxylase has, sofar, only been found in strains of the lower eukaryote Trichosporon properties respects

cutaneum.

with

related

a strictly

enzymes

soil

phenol

yeast.

While

sharing

hydroxylase

differs

a single

N-terminal

general

in

several

(3,5-8).

N- and C-terminal

Sequences

Edman

of the

which

aerobic

(1,2),

degradation

whole protein

supports our earlier assumption

enzyme digestion

(5). with

Overlapping

fragments

Staphylococcus

yielded

of two identical from

sequence,

subunits in the

hydroxylamine

cleavage

and

dimeric from

V8 protease formed a continuous stretch of

80

residues from the N-terminal (Fig 1). The progressive release of amino acids by carboxypeptidase A suggested the sequence

-Leu-Ser-Thr-Ala at the C-terminal.

Sequence homologies with other flavoenzymes are shown in Fig 2, including the sequence "fingerprint", common to binding sites for NAD, NADP and FAD, derived by Wierenga et al. and defined as a the "ADP-binding pap-fold" (10). These authors showed that a sequence which matches the "fingerprint" or deviates from it at a single position can be used to predict the occurrence of such a binding fold with

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

236 great

confidence.

The

sequence

from

Asp-9

to

Asp-42

of

phenol

matches the "fingerprint" except at Asp-9 and thus qualifies as an

hydroxylase "ADP-binding

pap-fold". In each of the two flavoenzymes with known 3-D structures, PHB and GR, such an ADP-binding-fold is located near the N-terminal. (cf Fig 2 and Ref 10) and so it does in phenol hydroxylase. 1

10

TKYSESYCDV L

20

LIVGAGPAGL HI

30

MAARVLSEYV Ml-

40 50 60 RQKPDLKVRI I O K R S T K V Y N G Q A D G L Q CRT HI 11 VI 70 LESLKNLRIA

80 DK1XSEXNDM

-H2-

-vH Fig 1. The N-terminal amino acid sequence of phenol hydroxylase. Hydroxylamine (H) and S t a p h y l o c o c c u s V8 protease (V) fragments are indicated below the sequence. Solid lines indicate the part of the peptides that were successfully sequenced by Edman degradation. The sequence of residues 1-26 was also determined by Edman degradation of the whole protein. I f) 1 | « 1 | loop—| ( » 1 D V I I V G A G P A G l M A A R V l S E Y V R t J K P D l K V R l ID QYAI I G A G P S O l l L G Q l L I I K A - - 01 - - - O N V t l t DYLV I G G G S G G L A S A R R A A t l . - - O A - HAAVVE 0A!VIGGGFG0lYAVKKlR0t--tEl--K»qAF0 DMVVGGGSTOCClAORlANl-DDqNl-TVALIt KIVIVGGGAGGLEMATQlGIIKlGRKKKAKtTLVD * * . Fingerprint (10) o o o G O O o t > o o ft PHY 9-42 PIIB 4-32 OR 22-50 CIIH 7-36 A0X 8-39 HUM 6-39

Fig 2. ADP-binding sequence motifs in FAD-dependent enzymes. The sequence from Asp-9 to Asp-42 of phenol hydroxylase is aligned with the sequences of phydroxybenzoate hydroxylase from Pseudomonas fluorescens (PHB, EC 1.14.13.2) [13], human glutathione reductase (GR, EC 1.6.4.2) [14], cyclohexanone monooxygenase from Acinetobacter sp. (CHM, EC 1.14.13.22) [15], alcohol oxidase from Hansenula polvmorpha (AOX, EC 1.13.13) [16], NADH-dehydrogenase from E.coli (NDH. EC 1.6.99.3) (17) and with the "fingerprint" sequence of Wierenga et al [10]. The symbols in the "fingerprint" denote: glycine (G), small and hydrophobic residues (O; A,I,L,V,M or C), a basic or hydrophilic residue ( ; K,R,H,S,T,Q or N) and an acidic residue (A;D or E). The secondary structure elements of the proposed pap-fold are indicated.

237 Sequences from NADPH-binding In

free

solution,

the

Site

inhibitor

of

phenol

hydroxylase,

pyridoxal-5'-phosphate

(PxyP), acts competitively with NADPH, while it does not displace bound FAD nor does it perturb the

spectrum of the free enzyme (7). The aldehyde group of PxyP

reacts with E - N H 2

to form a Shiff's base. The phosphate moiety may bind to

phosphate binding sites at a distance of ca 1.2 nm, an estimated maximum distance between Shiff's

the a - c a r b o n base

with

and the phosphate

marker was introduced the binding

of

group

sodium[3 H]borohydride,

a

into phenol hydroxylase

NADPH

as

distinct

in

PxyP-Lys.

By

covalently

attached

to locate

sequences

from those

affecting the

reducing

the

radioactive involved

binding

of

in

FAD.

Digestion with S t a p h y l o c o c c u s V8 protease of enzyme labelled in this way yielded three peptides (PV1, PV2, PV3) with sequences as in Fig 3(9).

PV1:

KGGRVDRTKFTPE

PV2:

YVRQKPDLKVRIIDKRSTKVYNGQADGLQCRTLE

PV3 :

R V F I AGDACHTHSPjCAGQGMNTSMMD

Fig 3. Sequences around the phosphopyridoxylated lysyl residues (underlined), determined in peptides obtained by digestion with S t a p h y l o c o c c u s V8 protease. PV1:

KGGRVDRTKFTPE ~ I I I II AIGRVPNTKDLSL

G R ( 2 8 8 - 3 0 0 ) :

PV3: PHB(2 80-305)

:

RVFIAGDACHTHSPKAGQGMNTSMMD 1 1 1111 1 r 1 1 RLFLAGDAAHIVP PTGAKGLNLAAS D

Fig 4. Alignment of PV1 with residues 288-300 of human GR and of PV3 with residues 280-305 of PHB. The labelled lysyl residues are underlined. The

sequence

of

PV1

has

38%

homology

with

residues

288-300

of

human

glutathione reductase (Fig 4), a sequence located at the junction of the NADPH domain

and the central domain (11). The sequence of PV3 has 42%

homology

with a sequence in the active site of p-hydroxybenzoate hydroxylase (Fig 4), (12). The sequence of PV2 is located near the N-terminal (Tyr-29 to Glu-62, cf Fig 1)

238 with the labelled residue (Lys-43) at the C-terminal of a predicted ß a ß

structure,

likely to be involved in the binding of the ADP part of FAD (10). Thus only PV1 and PV3, but not PV2, may form part of the NADPH-binding site. References 1.

Massey, V., A. Claiborne, K. Detmer, L.M. Schopfer. 1982. In: Oxygenases and Oxygen Metabolism (M. Nozaki, S. Yamamoto, Y. Ishimura, M.J. Coon, L. Ernster.eds) AP, p. 185,

2.

Müller, F. 1985. In: Biochem. Soc. Trans. H , p. 443.

3.

Neujahr, H.Y., K.G. Kjellén, 1978. J. Biol. Chem. 2 5 1 . 8835.

4.

Neujahr, H.Y., A.

5.

Sejlitz, T.,

6.

Neujahr, H.Y., A. Gaal, 1975.

7.

Neujahr, H.Y.

8.

Neujahr, H.Y. 1988. Biochemistry 2 2 , 3770.

9.

Sejlitz, T., C. Wemstedt, Â. Engström, H.Y. Neujahr. 1989. 225.

Gaal,

1973.

Eur. J. Biochem. H , 386.

H.Y. Neujahr, 1987.

Eur. J. Biochem. H Q , 343.

Eur. J. Biochem. S i , 351.

K.G. Kjellén, 1980.

Biochemistry J j L 4967.

10. Wierenga, R.K., P. Teipstra, W.G.J. Hoi. 1986.

Eur. J. Biochem. 187.

J. Mol. Biol. M l , 101.

11. Pai, E.F., P.A. Karplus, G.E. Schultz. 1988. Biochemistry, 2 1 . 4465. 12. Weijer.W.J., J. Hofsteenge, Eur. J. Biochem. 133.109.

J.J.

Beintema,

R.K.

Wierenga,

13. Hofsteenge, J., W.J. Weijer, P.A. Jekel, J.J. Beintema. 133. 91. 14. Krauth-Siegel.R.L., R. Blattespiel, M. Saleh, R. Untucht-Grau. 1982. J. Biochem. 121. 259. 15. Chen, Y.C.J., O.P. Peoples,

E.

J.

1983. Schiltz,

Drenth.

1983.

Eur. J. Biochem R.H.

Schirmer,

C.T. Walsh. 1988. J. Bacteriol. 170.781.

16. Ledeboer, A.M., L. Edens, J. Maat, C, Visser, J.W. Bos, C.T. Verrips. 1985, Acid R e s . I i _ 3063.

Nucl.

17. Young, I.G., B.L. Rogers, H.D. Campbell, A. Jaworowski, D.C. Eur. J. Biochem. 116. 165.

1981.

Shaw.

2-AMINOBENZOYL-COA MONOOXYGENASE/REDUCTASE, A NOVEL TYPE OF FLAVOPROTEIN HYDROXYLASE

B. Langkau, S. Ghisla Dept. of Biology, University of Konstanz, 7750 Konstanz, FRG V. Massey Dept. of Biol. Chem., University of Michigan, Ann Arbor, MI-48109, USA G. Fuchs Dept. of Appi. Microbiol., University of Ulm, 7900 Ulm, FRG

Introduction Recently we reported on the unusual course of a reaction catalyzed by 2-aminobenzoyl-CoA monooxygenase/reductase from a Pseudomonas sp.(l). This enzyme catalyzes the insertion of one atom of oxygen into the aromatic moiety of the CoA substrate. This enzyme is uncommon in several aspects: a) It is the first flavin dependent hydroxylase to act on a CoA thioester as a substrate, b) it inserts a hydroxyl function into the para position of the aromatic amine and c) it catalyzes the reduction of an intermediate to yield a hydrogenated and hydroxylated product. In order to better understand the chemistry of this peculiar reaction course, and possibly to gain information on the general mechanism of flavin-dependent hydroxylation, we have studied some of the basic properties of this enzyme in comparison with the properties of some typical hydroxylases.

Results and Discussion Spectral properties of the enzyme 2-Aminobenzoyl-CoA monooxygenase/reductase shares some common properties with the dehydrogenase-monooxygenase class of flavoproteins (2,3,4). The enzyme does not exhibit any semiquinone formation during the course of either photoreduction (EDTA/light) or reduction

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter&Co., Berlin • New York - Printed in Germany

240 with NADH and yields a fully reduced flavin spectrum. In analogy to all hydroxylases no formation of a sulfite-N(5)-adduct was observed upon incubating the enzyme with up to 20 mM sulfite for 17h. The addition of 2-aminobenzoyl-CoA causes significant alterations in the visible absorption of 2-aminobenzoyl-CoA monooxygenase/reductase. The absolute and difference absorbance spectra are shown in Figure 1A and B. A plot of % ES-complex formed versus the equivalents of substrate added indicates half-site reactivity (Figure 2). 2-Aminobenzoyl-CoA primarily binds to one subunit of the dimeric enzyme tightly (Kd=l|iM) and to the second subunit only weakly, and with only small spectral perturbations, which make it difficult to determine accurately the Kd value. This correlates with the observation of substrate inhibition in turnover experiments at [2-aminobenzoyl-CoA] >150 (J.M (5).

Wavelength (nm)

Wavelength (nm)

Figure 1. Spectrophotometric titration of 2-aminobenzoyl-CoA monooxygenase/reductase with 2-aminobenzoyl-CoA. Conditions: 52.6 |J.M enzyme bound flavin, 50 mM sodium phosphate, pH 7.4, 0.1 mM EDTA; temperature 4°C. All spectra were recorded versus the corresponding concentration of substrate in the reference cuvette and corrected for dilution. A. Spectrum of 2-aminobenzoyl-CoA monooxygenase/reductase before( ) and on ( ) addition of 62.7 |j.M substrate. B.Calculated difference spectra obtained on addition of 2.3 (iM, 8.9 |iM, 17.1 |iM, 25.4 (iM, and 62.7 (J.M 2-aminobenzoyl-CoA (substrate concentrations above 62.7 (iM resulted in a decrease of A absorbance at most wavelength).

241

Figure 2. Plot of %ES-complex (determined from A absorbance at 443 nm) versus equivalents substrate added to the enzyme. Conditions as in Figure 1. The stoichiometry of 2aminobenzoyl-CoA/enzyme bound flavin=0.5 : 1 was estimated from the point of intersection of initial slope of ES-complex and maximal absorbance changes. Equivalents=[S]o/[E-FI]o

Kinetics of enzyme reduction and reoxidation The course of reduction with NADH is also clearly biphasic as shown in Figure 3, one half of the enzyme reacting very fast (complete reaction within < 1 min), the second half being reduced with a t lfl 12.5 min. Reoxidation of reduced enzyme with either molecular oxygen (Figure 4) or NEM as electron acceptor again exhibits strongly biphasic reaction courses.

Figure 3. Course of reduction of 2-aminobenzoyl-CoA monooxygenase/reductase with NADH under anaerobic conditions. Conditions: 15.1 pM enzyme, 50 mM sodium phosphate, pH 7.4, 0.2 mM NADH; temperature 4°C. Half-reduction within the mixing time (< 1 min). The second phase occurs with a t lfi = 12.5 min (pseudo-first order approximation).

Figure 4. Course of reoxidation of photoreduced 2-aminobenzoyl-CoA monooxygenase/ reductase by 0 2 . Conditions: 8.9 pM enzyme, 50 mMNaPi, pH 7.4,12mMEDTA;4°C.Upon admission of air into the anaerobic cuvette halfreoxidation is observed within the mixing time of (< 0.5 min). The second phase of the reaction yields >98% reoxidized enzyme within 40 min.

242 Catalytic reaction with N-ethylmaleimide (NEM) NEM is an artificial substrate for the enzyme, being converted to NES in the presence or absence of oxygen. This finding suggests a strict separation between the two reaction parts and provides a possibility for uncoupling substrate reduction from hydroxylation steps. Steady state analysis of 2-aminobenzoyl-CoA monooxygenase/reductase, varying both NEM and NADH concentrations yields a series of parallel Lineweaver-Burk plots (not shown), suggesting a ping pong mechanism (Scheme 1). The kinetic constants determined from these plots were Km (NADH) = 26 jiM, K m (NEM) = 0.47 mM and V mii = 5650 min"1 O

O

Scheme 1. Mechanism for the reaction of 2-aminobenzoyl-CoA monooxygenase/reductase with NEM.

Conclusions Half-site reactivities observed in binding, reduction andreoxidation experiments suggest that 2aminobenzoyl-CoA monooxygenase/reductase is an unusual flavin-dependent hydroxylase, although some properties common to this class of enzymes were found.However, it cannot be concluded yet, whether the two subunits of the dimeric enzyme catalyze independently one part of the reaction (hydroxylation or reduction) or are involved both in all steps of catalysis.

References 1.

Langkau, B., S. Ghisla, R. Buder, K. Ziegler, G. Fuchs: Eur. J. Biochem.(in press)

2.

Strickland, S., V. Massey. 1973. J. Biol. Chem. 248, 2944.

3.

Spector, T„ V. Massey. 1972. J. Biol. Chem. 247, 4679.

4.

Massey, V., F. Müller, R. Feldberg, M. Schuman, P.A. Sullivan, L.G. Howell, Mayhew, R.G. Matthews, G.P. Foust. 1969. J. Biol. Chem. 244, 3999.

5.

Buder, R„ G. Fuchs. 1989. Eur. J. Biochem. I M , 629.

S.G.

STRUCTURAL STUDIES ON THE PORCINE LIVER MULTISUBSTRATE FLAVIN-CONTAINING MONOOXYGENASE

Katy K. Korsmeyer, Lawrence L. Poulsen, and D.M. Ziegler Clayton Foundation Biochemical Institute, Department of Chemistry and Biochemistry, University of Texas at Austin, Austin, TX 78712

Introduction

Porcine liver flavin-containing monooxygenase

(FMO) (EC

1.14.13.8) catalyzes the oxygenation of a wide range of structurally diverse soft nucleophiles.

Studies on mechanism

(cf Ref 1. for recent review) have shown that the oxygenatable substrate is not required for flavin reduction by NADPH nor for oxygenation of dihydroflavin by molecular oxygen.

The latter

step yields the enzyme-bound 4a-hydroperoxyflavin which is stabilized by the microenvironment at the active site. Previous reports have shown that NADP + (2,3) and phosphatidylserine

(4) contribute to stabilization of the

oxygenating intermediate, but, potential contributions of other non-protein components are largely unknown.

Further insights

into the molecular basis for this unusual feature of FMO will require detailed structural information including, initially, a complete description of the composition of FMO. The amino acid sequence deduced from cDNA clones (5) suggests that FMO contains two potential N-glycosylation sites located at Asn-120 and Asn-314 (Figure 1).

The studies

described in this report demonstrate that porcine liver FMO contains covalently bound carbohydrates.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York-Printed in Germany

244

1 M A K R V A I

V (G A *

G V S G) L A S I K C *

...

*

111 C S V T K H E D F N V F D A V M Y C T G

T T G Q W D V V T L F L T N P Y L P L D

181 I F K D K S V L V V

(G M G N S G)T D I A

...

268 N Y G L I P E D R I

O L R E P V L

...

301 P S

V V F N S S P E

*

I K E V K E

NS

*

N D E

C E G K Q E

S A

*

EE

Figure 1. Partial Primary Sequence of Porcine Liver FMO Derived from cDNA Nucleotide Sequence. Peptides obtained from automated Edman sequencing of V-8 protease digests after transferring to polyvinylidine membranes (6,7) are underlined. N-linked oligosaccharide binding sites with the consensus sequence of Asn-X-Ser/Thr are identified by asterisks. Flavin binding site of Gly-X-Gly-X-X-Gly (residues 9-14) and NADP+ binding site of Gly-X-Gly-X-X-Gly/Ala (residues 191-196) are in parenthesis. A

I

2

3

4

5

6

7

8

9

B

MW

1

2

3

4

5

6

7

B

9

66.000-

mm 21,500-

Figure 2. Western Blots of FMO Samples. A.After transfer, blot was immunostained with rabbit anti-hog liver FMO antibody. Bands were detected using anti-rabbit IgG antibody conjugated with horse-radish peroxidase. Color developed with 4-chloro-lnapthol as substrate. B.After transfer, blot was incubated with Concanavalin A followed by incubation with rabbit antiConcanavalin A antibody. Bands were visualized as stated above for A. Lane 1.0.2^g of 99% pure FMO. Lane 2.0.4Hg of 39% pure FMO. Lane 3.0.4\ig of 56% pure FMO. Lane 4.0.4|ig of 41% pure FMO. Lane 5.0.4|lg of 46% pure FMO. Lane 6-9.0.2\lg of single bands isolated by agarose gel electrophoresis. MW.Molecular weight standards BSA, OVA, trypsin inhibitor, and lysozyme with only OVA and lysozyme showing positive staining for carbohydrates.

245 I

2

3

43,000

Figure 3. SDS-PAGE of partially purified FMO, denatured, and treated with Endo H. Lane 1. 5|ig control sample without Endo H incubated at 37°C for 18 hrs. Lane 2.Treated with lOmU Endo H at 37°C for 18 hrs. Lane 3.Treated with 25mU Endo H at 37°C for 18 hrs.

21,500

Results FMO apparently fully resolved from contaminants on SDS-PAGE mini-gels stained positive for carbohydrates by the PAS (periodic acid/Schiff's base) and the Thymol-sulfate methods (8). These results strongly indicate that FMO bears covalently attached carbohydrates. Samples of FMO at different stages of purity, subjected to SDS-PAGE, were transferred to two nitrocellulose sheets. One was treated with IgG to pig liver FMO and the other with the plant lectin Concanavalin A. Concanavalin A reacts with a-D-glucose and a-D-mannose in oligosaccharides. Binding visualized as described in Fig.2 shows that the bands binding anti-FMO IgG correspond exactly to the bands binding Concanavalin A,confirming the presence of carbohydrates on FMO. Oligosaccharides linked to asparagine residues belong either to the high mannose or the complex types. The high mannose type results from addition of the oligosaccharide to the Asn-X-Ser/Thr of the growing peptide chain on the luminal side of the endoplasmic reticulum. Processing of the high mannose chain as the protein is

246

transported through the Golgi apparatus results in the complex type. The two can be distinguished by treatment with Endo-Nacetylglucosaminidase H (Endo H) which hydrolyzes the chitobiose sites of N-linked high mannose type oligosaccharides. As shown in Fig.3, FMO treated with Endo H showed a decrease in molecular weight of approximately 5kDa indicating the presence of high-mannose type oligosaccharides. Studies to characterize the binding site of the carbohydrates, the composition of the oligosaccharides, and the effect its absence has upon enzyme activities are in progress.

Acknowledgement This research was supported by the Foundation for Research.

References 1. Ziegler, D.M. 1988. Drug Metab. Rev. H , 1-32. 2. Jones, K.C. and Ballou, D.P. 1986. J. Biol. Chem. ¿£1, 25532559. 3. Ziegler, D.M., Poulsen, L.L., and Duffel, M.W. 1980. In: Microsomes, Drug Oxidations and Chemical Carcinogenesis, Vol.2 (M.J. Coon and et al, eds.) Academic Press, pp.637-645. 4. Poulsen, L.L., Nagata, T., and Taylor, K.L. 1987. In: Flavins and Flavoproteins (D.E. Edmondson and D.B. McCormick, eds.) Walter de Gruyter, Berlin, New York. pp.577580. 5. Gasser, R., Tynes, R.E., Lawton, M.P., Korsmeyer, K.K., Ziegler, D.M., and Philpot, R.M. 1990. Biochem. 22., 119-124. 6. Cleveland, D.W., Fischer, S.G., Kirschner, M.W., and Laemmli, U.K. 1977. J. Biol. Chem. 2£2, 1102-1106. 7. Matsudaira, P. 1987. J. Biol. Chem. 232., 10035-10038. 8. Gander, J.E. 1984. In: Methods Enzymol., Vol.104 (W.B. Jakoby, eds.) Academic Press, pp.447-451.

THE COFACTOR DEPENDENT INTERACTION OF MOLECULAR OXYGEN WITH PHENYLALANINE HYDROXYLASE

S.W. Bailey, J.P. Crow, J.E. Ayling Department of Pharmacology, College of Medicine, University of South Alabama, Mobile, AL 36688, USA

The interactions of tetrahydropterins and dihydroflavins with molecular oxygen share many common features. Our recent studies show that these two classes of enzymes may differ in the rate determining step of the reaction, and that this appears to be associated with other distinctions between the overall pathways. With the flavin dependent hydroxylases the step involving formation of the C4a-hydroperoxide intermediate by interaction of oxygen with reduced enzyme is not rate limiting (1,2). However, with the tetrahydrobiopterin dependent phenylalanine hydroxylase from rat liver there is a strong correlation between the maximum velocity (at saturation with all three substrates) and the nonenzymatic rate of one-electron transfer between cofactor analog and 0 2 (3) . The rate of one-electron transfer from analogs of (6R)tetrahydrobiopterin ((6R)-BH4) to 0 2 to form the cofactor radical and superoxide was determined by suppressing the major sources of chain propagation, a condition difficult to achieve with unblocked dihydroflavins. The decrease of tetrahydropterin autooxidation caused by superoxide dismutase (SOD) and/or catalase, like that with dihydroflavin (4) , has long been recognized. However, propagation by comproportionation between the oxidized guinoid dihydro species and reduced cofactor, PH4 + gPH2 = 2(PH3"), has been overlooked. This property is well understood with the F1H2/F1 couple which is in rapid equilibrium with moderate concentrations of the semiquinone (5,6). Pterin radicals are not so stable, but are produced fast enough to circumvent the initiation reaction. This route was eliminated

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

248

by inclusion of a high activity of dihydropteridine reductase (DHPR). In the presence of DHPR, SOD, and catalase the rate of NADH disappearance is linear with time, and first order in both tetrahydropterin and 02. Cofactor analogs were synthesized which span a range of kobs of over two orders of magnitude (3) . Even the fastest observed (0.2 M"1 sec"1) is between 102 and 103 times slower than the rates of l-electron transfer from 5-alkyldihydroflavins to 0 2 (7,8). The measurement of the maximum velocity with phenylalanine hydroxylase required determining the Km's for all three substrates as a function of cofactor structure. Establishing steady state kinetic parameters of 0 2 for oxygen affinities below 250 ;xM was facilitated by development of a method utilizing progress curve analysis (Figure 1)(3). Oxygen 300

500

Figure 1 250

(6B)-PropyWH4

200

18//M

© 150

O) S? °

X

100

6,7-(CH,VH4 K,- 210/iM

50

0

1

50

I

100

1

150

h

200

Time (seconds)

250

300

0

20

40

60

80

100 120 140

K m for0 2 (%)

F i g u r e 1. Progress curves for oxygen consumption with (6R)propyl-tetrahydropterin (PH4) or 6, 7-(CH3) 2-PH4 as cofactor with determined from the integrated rate equation. The amount of enzyme was adjusted so that reactions were completed in 4-5 min. Figure 2. Relationship between kcat and oxygen K„, for 11 cofactor analogs in which Ri is H or CH3, and R 2 is CH3, phenyl, propyl or dihydroxypropyl; R3 is H, CH3 or benzyl.

249 consumption was coupled to NADH oxidation with DHPR, and absorbance changes digitized over the four minute time course of a typical reaction. The integrated rate equation was fitted directly (Figure 1) through the use of the implicit variable capability of MINSQ (Micromath, Salt Lake City, Utah). A linear relationship was found between the maximum rate of hydroxylase dependent cofactor oxidation and the rate of nonenzymatic one electron transfer with the majority of compounds investigated. These showed a catalytic effectiveness of about 10 Molar. However, a small group of analogs, including the natural cofactor, (6R)-BH4, was observed to have up to a 10-fold higher effectiveness. Specifically, in the case of (6R)—BH4 this increase was dependent on the hydroxylase being pre-activated by phenylalanine (3). An inverse relationship was observed between K,^ for O2 and V^x, with a variety of cofactor analogs (Figure 2). In more typical enzyme reaction mechanisms, if affinity for a substrate changes with kcat, K,,, increases with increasing turnover. A possible explanation of results presented in Figure 2 is that molecular oxygen adds immediately after a rate limiting but reversible step. However, as discussed above, the rate limiting step appears to be associated with one-electron transfer from cofactor to already bound 02. This seeming contradiction could still be met by the following Scheme:

It is generally believed that formation of the observed C4ahydroperoxyl flavin intermediate of the monooxygenases proceeds through formation of the {Flsem, 02~} pair which is close in free energy to the main transition state (7,8), although recently an

250

alternate interpretation has been presented (9). In the case of tetrahydropterin cofactors, the collapse of the {PH4+", 02~} pair to the hydroperoxide, which is yet to be directly observed, may not be as facile as with dihydroflavin. The binding of the second 0 2 could be used to drive an unfavorable equilibrium in the forward direction. Such a process could either be an example of alternating sites (phenylalanine hydroxylase is a dimer or tetramer of identical subunits) or the second oxygen may simply displace the first from its initial binding site (iron?). In either case there is no free enzyme present in steady-state turnover. The above Scheme allows an inverse for relationship between Vmax (determined primarily by kp) and 0 2 if k.p is faster than both k p and the slowest subsequent step.

Supported by NIH grants GM-30368 and NS-26662.

1. Entsch, B., D.P. Ballou, V. Massey. 1976. J. Biol. Chem. 251, 2550-2563. 2. Entsch, B., M. Husain, D.P. Ballou, V. Massey, C. Walsh. 1980. J. Biol. Chem. 255, 1420-1429. 3. Bailey, S.W., S.B. Dillard, J.P. Crow, J.E. Ayling (in preparation). 4. Massey, V. , G. Palmer, D.P. Ballou. 1973. In: Oxidases and Related Redox Systems (T.E. King, H.S. Mason, and M. Morrison, eds.) Univ. Park Press, Baltimore, Md. pp 25-49. 5. Gibson, Q.H., J.W. Hastings. 1962. Biochem. J. 83, 368-377. 6. Muller, F., P. Hemmerich, A. Ehrenberg. 1971. In: Flavins and Flavoproteins (H. Kamin, ed.) University Park Press, Baltimore, Md. pp. 107-122. 7. Eberlein, G., T.C. Bruice. 1983. J. Am. Chem. Soc. 105, 6685-6697. 8. Bruice, T.C. 1984. In: Flavins and Flavoproteins (R.C. Bray, P.C. Engel, and S.G. Mayhew, eds.) Walter de Gruyter, Berlin, New York. pp. 45-55. 9. Massey,V.,L.M. Schopfer and R.F. Anderson. 1988.In: Oxidases and Related Redox Systems (T.E. King, H.S. Mason and M. Morrison, eds.). Alan R. Liss, Inc., New York. pp. 147-166

Flavin Dependent Bioluminescence

MECHANISMS OF BACTERIAL LUCIFERASE A N D AROMATIC HYDROXYLASES

Shiao-Chun Tu, Humphrey I. X. Mager, Ruxin Shao, Ki Woong Cho, and Lei Xi Department of Biochemical and Biophysical Sciences, University of Houston Houston, Texas 77204-5500

Bacterial Luciferase Hastings and colleagues (1-3) have proposed a scheme that has become a foundation for our mechanistic understanding of bacterial luciferase. FMNH2 reacts with O2 to form the 4ahydroperoxyFMNH intermediate II. I I either decays to form F M N and H2O2 with little or no light emission or reacts with an aldehyde to form the assumed 4a-peroxyhemiacetalF M N H (III). The breakdown of I I I is proposed to follow a Baeyer-Villiger mechanism forming directly a fatty acid and the primary excited species 4a-hydroxyFMNH* (IV*). In the presence of a lumazine protein (LP) (4, 5), the bioluminescence kinetics are changed, quantum yields increased, and the emissions blue shifted (e.g. from 490 to 476 nm). This endogonic shift, however, can not be attributed to a Forster energy transfer from IV* to LP. An FMN-containing yellow fluorescent protein (YFP) was also found to affect the emission kinetics, intensity, and color (6, 7). Contrary to a claim (6), energy transfer from IV* to YFP alone can not account for the observed phenomena (7). On account of these findings, the fundamental questions regarding the identity of the primary excited species and the mechanism for its formation, with or without LP or YFP, should be re-addressed. An Electron (Charge) Transfer Mechanism for Luciferase: Based on the original formulation by Mager and Addink (8), a chemically initiated electron exchange luminescence (CIEEL) mechanism for luciferase (9-12) is shown in Scheme 1. II reacts with aldehyde R

H

R'

A

B

C

0 Scheme 1

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

E

254 (stage A) to form a caged flavin radical and alkylhydroxyl oxy radical pair (stage C) via A—>C or A—>B—>C. Following a conversion of the initial oxidizing oxy radical to a reducing carbon-centered radical (stage D), a 1-e transfer occurs leading to the formation of either HF4a-OH* (Ea) or R-COOH* (Eb) as the primary excited species. In the latter case, an energy transfer from R-COOH* to HF-4a-OH will generate the secondarily excited emitter HF-4aOH* (Ec). Energy transfer from R-COOH* to an alternative fluorophore X (e.g. LP or YFP), if present, could also occur to form X* as an emitter (Ed). (I) CI EEL Mechanism

in General:

Catalani and Wilson (13) have shown that some

CIEEL reactions are rather inefficient (quantum yields

Therefore, a general applica-

bility of CIEEL mechanisms should be viewed with caution. However, in bioluminescence, the binding of the emitter by a luciferase could significantly enhance the emission efficiency. Moreover, bioluminescence, involving bound reactants and intermediates, should enjoy a marked entropic advantage over the nonenzymatic chemiluminescence. (II) Electron versus Charge Transfer:

It has been pointed out that the electron transfer

CIEEL mechanism can be modified to involve charge transfer complexes without violating the basic principle (13, 14). We concur with this consideration, and the caged radicals shown in stages C and D of Scheme 1 could be replaced by charge transfer complexes. (III) Intermolecular

versus Intramolecular

Transfer:

The radical pair (C) could be

formed following either intermolecular or intramolecular electron transfers (12). The A—>C process would involve an assisted homolysis of the peroxy bond to form (HF-4a-0") + ' and HO 1 . Intermolecularly, the HO- adds to aldehyde to give, along with (HF-4a-0~) + ', the radical pair (C). For A—>B—>C, an intramolecular 1-e transfer from the flavin N5 to the peroxy bond of HF-4a-peroxyhemiacetal would form the same radical pair. The reaction of II with aldehyde to form an aliphatic dioxirane intermediate was proposed by Cho and Lee (15). The same dioxirane intermediate and & CIEEL mechanism were recently postulated by Raushel and Baldwin (16) without referring to earlier reports (e.g. 8-10, 15). An intermolecular electron transfer from HF-4a-OH to the dioxirane could also form the radical pair (C) (16). As a model, dimethyldioxirane was reacted with 5-ethyl-4a-hydroxy-3-methyl4a,5-dihydrolumiflavin (5-EtF-4a-OH). Only trace chemiluminescence was detected and the addition of 3-methyllumiflavin as a fluorophore had no positive effect. Therefore, these limited results so far do not suggest dioxirane as an intermediate for luciferase. All electron transfer processes described above can be modified to depict corresponding charge transfers. (IV) Primary Excited Species:

By reduction of parinaric acid (17), we have obtained

cis-parinaric aldehyde that retains the absorption and fluorescence ( E m m a x 420 nm) of the parent acid and exhibits a significant bioluminescence activity with luciferase. Assuming that

255 R - C O O H * is the primary excited species and the energy transfer to HF-4a-OH is x Me3Lm s o x /Me3Lm o ;)X (VI) LP- and YFP-induced Bioluminescence:

Bioluminescence shifts induced by L P and

YFP are due to emissions from LP-lumazine* and YFP-FMN*. The emission enhancement by L P most likely results from a higher emission quantum yield of 0.54-0.59 for L P (20) than the 0.18 for HFMN-4a-OH (18). The fluorescence quantum yield of Y F P - F M N has not been determined but possibly could also be >0.18. The altered emission kinetics could be a secondary effect of luciferase conformational change induced by LP and Y F P binding. Therefore, the possibility remains that LP and YFP are simply energy acceptors (Scheme 1, Ebd). Alternatively, LP and YFP could also directly participate in the chemical reaction. This latter possibility, reasonably supported by experimental findings, has significant mechanistic implications and should be rigorously verified. In this regard, we now propose the following mechanism (Scheme 2) involving LP and YFP (X) as a reactant. F + H20

R-COOH + X H-C-OH I R

R-C

Scheme 2 A 1-e transfer from X to 4a-peroxyhemiacetalFMNH (III) peroxy bond generates X + \ HF-4a-0~, and an alkylhydroxy-oxy radical (A). Alternatively, the N5 of I I I donates one electron to the peroxy bond to form the (HF-4a-0~) + - and the alkylhydroxy-oxy radical. X subsequently transfers one electron to the flavin radical to generate HF-4a-0" and X + > (B). A similar scheme can start with I I and aldehyde without involving I I I as discussed for Scheme 1. The alkylhydroxy-oxy radical transforms into a carbon-center radical and a

257 subsequent 1-e transfer to X + - generates the X*. The HF-4a-0~ decays to form oxidized flavin and water. A similar scheme has been proposed by Hastings (21) but with one critical difference, namely a reductive formation of a crucial (HF-OH) - - anion is hypothesized. However, using 5-EtF-OH, we could not detect any electrochemical formation of such a 4ahydroxyflavin radical anion at as low as -1.5 V. Scheme 2 also depicts a 1-e oxidation (designated "superoxidation") of the so called "fully" oxidized flavin and lumazine. Using 3-methyllumiflavin (3-MeF o x ), tetraacetylriboflavin (Ac4Rf o x ) 3-methyl-5-ethyllumiflavinium cation (5-EtF ox + ), 6,7,8-trimethyllumazine (Me3Lm 0X ), and 3,6,7,8-tetramethyIlumazine (Me4Lm o x ) in acetonitrile, we have indeed observed 1-e oxidations in all cases (Table 1; subscript "sox" = superoxidized) (10, 12). Moreover, reduction of the superoxidized 3-MeF sox + - leads to chemiluminescence attributable primarily to excited 3-MeF ox (12).

Mechanism of Reduced Flavin Oxidation The oxidation of reduced flavoenzymes by O2 follows (E:FH2 + O2 —> E:F + H2O2) for oxidases or (E:FH2 + O2 + SH

E:HF-4a-OOH:SH -> E:F + SOH + H 2 0 ) for mono-

oxygenases. With certain effectors, monooxygenases partition between a hydroxylase and a pseudooxidase activity (E:FH 2 + O2 + SH —> E:HF-4a-OOH:SH -> E:F + SH + H 2 0 2 ) . Although oxidase and pseudooxidase activities follow the same overall reaction, the latter activity involves a HF-4a-OOH intermediate that has ever been detected for oxidases. Site-directed mutagenesis was carried out for Vibrio harveyi luciferase to convert the highly reactive a C y s 1 0 6 to an alanine (ocC106A) or a valine (aC106V) (22). Using decanal and dodecanal, the aC106A enzyme is 40-60% active and the aC106V enzyme is 2-10% active. Similar results have also been reported independently (23). Therefore, the a C y s 1 0 6 is not essential to luciferase chemical catalysis. Most interestingly, the aC106V enzyme is active in catalyzing the oxidation of FMNH2 (Fig. 2). The autooxidation of FMNH2 can be 0

Fig. 2. Stopped-flow time course of autooxidation and luciferase-catalyzed oxidations of FMNH 2 . Oxygen (0.25 mM) in 0.05 M Pi, pH 7.0, was reacted at 23°C with an equal volume of solution containing 7.9 hM FMNH2 ( - E), 8.7 |iM FMNH2 and 12 nM wild-type luciferase (+ W-T), or 11 nM FMNH 2 and 15 nM of ocC106V luciferase (+ aC106V). The single time scale at top is for the autooxidation and the double time scale is for the two enzymatic oxidations.

, <
89. 4. Schmidt, T. M., K. Kopecky, K. H. Nealson. 1989. Applied and Environmental Microbiology, 55, 2607. 5. Colepicolo, P., K.-W. Cho, G. O. Poinar, J. W. Hastings. 1989. Applied and Environmental Microbiology 2601. 6. Meighen, E. A., J. W. Hastings. 1971. J. Biol. Chem. 246, 7666. 7. Tu, S.-C., J. W. Hastings. 1975. Biochemistry 14, 4310. 8. Baldwin, T. O., M. M. Zigler, D. A. Powers. 1979. Proc. Natl. Acad. Sei. USA Z6, 4887. 9. Baldwin, T. O., L. H. Chen, L. J. Chlumsky, J. H. Devine, M. M. Ziegler. 1989. J. Biolumin. Chemilumin. 4, 40. 10. Ziegler, M„ L. Chen, L. Chlumsky, J. Devine, T. Baldwin. 1988. J. Biolumines. Chemilumines. 2, 279.

ON THE FLAVINS

MECHANISM AS MODEL

OF BACTERIAL COMPOUNDS FOR

LUCIFERASE.4a,5-DIHYDROREACTION INTERMEDIATES.

Jens W. Eckstein and Sandro Ghisla Dept. of Biology, University of Konstanz, 7750 Konstanz, FRG

Introduction The monooxygenation of long-chain aldehydes to form the corresponding carboxylic acids catalyzed by the FMN-dependent bacterial luciferase produces light as one of the reaction products (1): FMNH2 + O2 + RCHO -» FMN + H2O + RCOOH + hV The reaction producing excited states involves a 4a,5dihydroflavin intermediate (3). The breakdown of a 4aperoxyhemiacetal intermediate is the key step and has been formulated as an intramolecular CIEEL (chemically initiated Electron exchange luminescence ) mechanism (2,5,6). Based on the assumption that charge or electron transfer from an activator to the peroxide is connected with the rate-limiting step, a linear free energy xelationship (LFER) should be found between the one-electron oxidation potential of the electrondonating activator (the reduced 4a,5-dihydroflavin) and the rate of light emission decay. This relationship was tested by us some years ago using 8-substituted FMN derivatives as cofactors of the luciferase. Indeed we observed the expected dependence on the redox potentials of such FMN analogues in a linear free energy relationship (4). For this correlation the 2e _ -redox potentials of the F1Q^/1, 5—Fl^-^^ couples were used since the one-electron oxidation potentials of 4a,5dihydroflavins were not available; however, we reasoned that the two sets of potentials would exhibit the same dependence on substitution. To verify experimentally this fundamental premise, we have synthesized several 4a,5-dihydroflavins with varying residues at the 8-position and measured their oneelectron oxidation potentials by cyclic voltammetry (for detailed experimental conditions see (7)).

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

270

Results and Discussion One-electron oxidation of the 4a,5-dihydroflavin model compounds is reversible under the conditions used. The oneelectron oxidation potentials were tested on a Hammettrelationship concerning substituent effects according to the Figure la 800

800

E (measured, mV) Figure lb ->

>

900 -

-o CD

700

-r -

E

Chloro

Methoxy/*

500

E LU

-T

Methyl

=5 W

co CD

•

Amino

300 -400

jf

i -300

.

i -200

-100

Eo' (mV)

method described by Swain et.al.(8). Using optimized resonance and field constants an excellent correlation was obtained

271

between calculated and measured one-electron oxidation potentials (figure la). Comparing the effect of substitution at the 8-position on the one-electron oxidation potentials of 4a,5-dihydroflavins and on the 2e~-redox potentials of FMN derivatives we have found the very good correlation shown in figure lb. This confirms that the assumptions mentioned in the introduction are correct. A plot of the log of the rate of light emission decay (k) in the luciferase reaction against the one-electron oxidation potentials yields a value for a being about one third (=0.31) of the value found in a correlation with the 2e~-redox potentials of FMN derivatives (figure 2). Altogether we are now confident that the electron/charge donating properties of the reduced flavin moiety of the luciferase-bound 4a,5-dihydroflavin intermediate determines the rate and effectiveness of excited state production. The dependence shown in figure 2 is consistent with an intramolecular CIEEL mechanism in which partial charge transfer in the transition state can be envisaged as well(10).

1:8-methoxy 2: 8-methyl 3: 6,7-dimethyl 4: 8-methylthio 5: 8-nor 6: 8-fluoro 7: 8-chloro 8: 8-methylsulfonyl

500

1000

E(mV)

1500

Figure 2 Correlation of one-electron oxidation potentials with the rate of light emission decay in an assay with V.harveyi luciferase. Rate constants were taken from (4). One-electron oxidation potentials were measured by cyclic voltammetry or calculated by the method of Swain et.al (8) .

1

For discussion of typical a-values of =0.3 refer to (9)

272 Acknowledgement We would like to thank Dipl.-Chem. S. Kroner for her assistance with the cyclic voltammetry measurements. J.W.E. was supported by the LGFG of Baden-Württemberg.

References 1.

Hastings, J. W. 1978. Meth. Enzym. ¿7,

2.

Koo, J. Y. & Schuster, G. B. 1977. J. Am. Chem. Soc. 21, (18), 6107-6109.

3.

Kurfürst, M., Macheroux, P., Ghisla, S. & Hastings, J. W. 1987. Biochim. Biophys. Acta 924. 104-110.

4.

Macheou::, P., 3ck3t-=ir., J. & Ghisla, S. 1987. In: Flavins and Flavoproteins

125-135.

(D. E. Edmondson and D. B. McCormick,

eds.) Walter de Gruyter, Berlin, pp. 613-619. 5.

Macheroux, P. & Ghisla, S. 1984. In: Flavins and Flavoproteins

(R. C. Bray, P. C. Engel and S. G. Mayhew,

eds.) Walter de Gruyter, Berlin, pp.

669-672.

6.

Mager, H. I. X., Sazou, D., Liu, Y. H., Tu, S.-C. & Kadish, K. M. 1988. J. Am. Chem. Soc. 110. 3759-3762.

7.

Eckstein, J.W. & Ghisla, S. 1990.

8.

Swain, C.G., Unger, S.H., Rosenquist, N.R. & Swain, M.S. 1983. J. Am. Chem. Soc. 105, 492-502.

9.

Scandola, F., Balzani, V. & Schuster, G.B. 1981. J. Am.

(in preparation)

Chem. Soc. 1Ü1, 2519. 10.

Wilson, T. 1985. In: Singlet O2, Vol. II (A.A. Frimer, ed.) CRC Press, Boca Raton. 37-65.

KINETIC AND MECHANISTIC INVESTIGATION OF THE BACTERIAL LUCIFERASE REACTION

Frank M. Raushel, Husam M. Abu-Soud, Leisha S. Mullins, Wilson A. Francisco, and Thomas O. Baldwin Department of Chemistry and Biochemistry & Biophysics, Texas A&M University, College Station, TX 77843 USA

Introduction

Bacterial luciferase from Vibrio harveyi catalyzes the production of visible light from reduced FMN, molecular oxygen, and a long-chain aliphatic aldehyde. The other products are oxidized FMN, water, and the corresponding carboxylic acid (1). The chemical mechanism of this reaction has been previously shown to involve 4a-hydroperoxyflavin (2) and perhaps a dioxirane intermediate (3). Stopped-flow kinetic techniques have now been utilized to obtain microscopic rate constants for the interconversion of the various intermediates and enzyme complexes.

Results and Discussion

Stopped-flow techniques have been used to study the kinetics of the luciferase reaction in the absence and presence of decanal. All studies have been carried out in the dark under nitrogen atmosphere in Bis-Tris buffer, pH 7.0 at 25°C. Changes in the flavin absorption at 380 and 445 nm, in addition to monitoring the production of visible light with time, have been used in the construction of a kinetic model for the bacterial luciferase reaction. These optical probes occur on time scales ranging from + H2O2 -> HO- + H2O + O2) further react with VI and/or m to give II and IV. During these processes, greenish light is produced peaking at -500 nm in MeCN. This luminescence is believed to arise from Fl ox * (= III*; pathway B). However, due to a low spectral resolution at the present, a contribution of CL peaking -480 nm, such as from excited species derived from the reduction of II (pathway C), can not be excluded. We have found that the HO* produced in Fenton's systems reacts with Fl ox to give similar CL. We believe that this long known (8) but mechanistically poorly understood CL could also involve flavin species equivalent to II and/or IV as intermediates.

In conclusion, evidence has been obtained for the formation of unusual superoxidized lumazines and flavins and derived 4a-substituted flavin radicals which, upon back transfer of

280 an electron, could give rise to excited species. This has provided a chemical basis to study the possible occurrence of similar species in the blue and yellow fluorescent protein-induced bioluminescence. As a first approximation, these processes are proposed to follow the Eqs. 5a, b [Y ox = Lumox or Fl ox ; HFl-4a-Xi = FMNH-4a-peroxide plus aldehyde or FMNH-4aperoxyhemiacetal; D - = the reducing carbon centered radical R-C(OH)2 which has already been formulated (1) to arise in an intramolecular 1,2-hydrogen shift from the primary oxygen 0 1 centered radical R-CHOH derived from HFl-4a-Xi; D = RCOOH + H+], HFl-4a-Xi + Yox -> HFl-4a-X 2 + Y s o x + - + D -

(5a)

YSOx+- + D " -» Yox* + D

(5b)

The question as to whether charge transfer complexes play a direct role in luminescent pathways, as recently suggested (9, 10), will be considered in further refining Eqs. 5a, b.

Acknowledgements:

This work was supported by grants GM 25953 from NIH and E-1030

from the Robert A. Welch Foundation.

References 1. Mager, H. I. X., R. Addink. 1984. In: Flavins and Flavoproteins (R. C. Bray, P. C. Engel, and S. G. Mayhew, eds.) de Gruyter, Berlin, p. 37. 2. Mager, H. I. X., D. Sazou, Y. H. Liu, S.-C. Tu, K. M. Kadish. 1988. J. Am. Chem. Soc. 110, 3759; relevant references therein. 3. Kaaret, T. W., T. C. Bruice. Photochem. Photobiol. (in press) 4. Mager, H. I. X., S.-C. Tu, Y.-H. Liu, Y. Deng, K. M. Kadish. Photochem. Photobiol. (in press) 5. Getoff, N„ S. Solar, D. B. McCormick. 1978. Science 201., 616. 6. Ballard, S. G„ D. C. Mauzerall, G. Tollin. 1976. J. Phys. Chem. 80, 341. 7. Heelis, P. F„ B. J. Parsons, G. O. Phillips, A. J. Swallow. 1989. J. Phys. Chem. 22., 4017; relevant references therein. 8. Strehler, B. L„ C. S. Shoup. 1953. Arch. Biochem. Biophys. 47, 8. 9. Catalani, L. H„ T. Wilson. 1989. J. Am. Chem. Soc. 1 U , 2633. 10. McCapra, F. 1990. J. Photochem. Photobiol. 51, 21.

ON THE MECHANISM BIOLUMINESCENCE

OF D I T H I 0 N I T E / H 2 0 2 - I N D U C E D

BACTERIAL

Shiao-Chun Tu and Ki Woong Cho Department of Biochemical and Biophysical Sciences, University of Houston Houston, Texas 77204-5500

Introduction Bacterial luciferase catalyzes a bioluminescent reaction (quantum yield -0.1) utilizing FMNH2, O2, and a long-chain aldehyde as substrates. When the in vitro reaction is carried out in the presence of H2O2, an additional emission is observed that is kinetically distinct and shows a high quantum output (1, 2). This H202-induced bioluminescence is different from a low quantum yield (~10~5) light emission initiated with H2O2 and FMN in lieu of FMNH2 and O2 (3,4). The mechanism for this high quantum yield H 2 02-induced emission has been the subject of some speculations (1, 2). Interestingly, this intense H 2 0 2 -induced light is obtained only when the reduced flavin is prepared by dithionite treatment but not by photochemical reduction or catalytic hydrogenation (5). In dithionite treatment of flavin models Mager and Tu (5) have detected the formation of three flavin-4a-sulfur adducts (i.e. 5-EtFl-4a-S03", 5-EtFl-4a-SOr, 5-EtFl-4a-S204') in addition to the fully reduced flavin. Consequently they suggest that similar adducts could be formed during the treatment of FMN with dithionite. At least one of these FMN-4a-sulfur adducts, stabilized by binding to luciferase, could react with H2O2 to form the 4a-hydroperoxyFMN intermediate II and subsequently with aldehyde to emit bioluminescence. In this report, evidence is presented to support such a proposal.

Results and Discussion Possible active species for this high quantum yield H202-induced bioluminescence have been suggested to be luciferase:4a-hydroxyFMN (intermediate IV) (1) and/or luciferase:FMN generated from the break down of either the intermediate II or IV (1,2). We have prepared these luciferase-bound flavin species and found them active in reacting with

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin-New York-Printed in Germany

282 aldehyde and H2O2 for light emission. However, their peak intensities and total quantum outputs were only 0.5-5% of that in the H2Q2-induced bioluminescence obtained by reacting luciferase with FMN, dithionite, aldehyde, O2, and H2O2. Therefore, neither intermediate IV nor luciferase:FMN generated in the normal bioluminescence reaction (in which the luciferase is conformationally different from the resting state of luciferase) represents the primary active species in the dithionite/H2C>2 (D/H)-induced bioluminescence. On the other hand, evidence has been obtained to indicate that FMNH-4a-sulfite is formed during dithionite treatment of FMN and is involved in the high quantum yield D/Hinduced bacterial bioluminescence. We have observed that the sulfite anion (SC>3=, a common oxidation product of dithionite) is a very effective inhibitor of bacterial bioluminescence. In the in vitro light emission initiated with FMNH2 (photochemically reduced), marked inhibition is evident either with the sulfite present at the onset of the bioluminescence (Fig. 1) or with the sulfite added secondarily during the light emission (Fig. 1 inset). However, a subsequent addition of H2O2 resulted in a much enhanced emission. On the basis of the recent chemical model studies (5), we propose that the sulfite inhibition is due to a nucleophilic displacement of the 4a-group in the intermediate II and/or the 4a-peroxyhemiacetalFMN (intermediate III) by the S0 3 =, and the subsequent emission initiated by H2O2 is the result of a re-conversion of the inactive FMNH-4a-S03" back to intermediate II. Fig. 1. Effects of sulfite and H2O2 on the in vitro bioluminescence. To 0.2 mL 0.05 M phosphate, pH 7, containing 8 jig luciferase, 26.5 |iM decanal, and zero (A) or 4 mM sulfite (S for B, C, and D), 0.1 mL of 50 jiM FMNH 2 prepared photochemically were injected to initiate the bioluminescence. For B, C, and D, 50 nL of 416 mM H2O2 (H) were also injected at the time indicated. Inset: Bioluminescence was initiated at time zero by injecting 1 mL of 50 nM photochemically reduced FMNH2 to a 1-mL solution containing 8 \ig luciferase and 64 (iM octanal. At the times indicated, 50 nL of 0.8 M sulfite and 0.1 mL of 1.25 M H2O2 were sequentially added. One LU = 6.8 x 10 9 q-s"1. Another important piece of evidence in support of our proposal is obtained through an examination of the oxygen effect on the D/H-initiated bioluminescence. Mager and Tu (5) show that, in treating oxidized flavin with dithionite, the yield of 5-EtFl-4a-S03" is greatly enhanced in the presence of oxygen due primarily to the oxidation of 5-EtFl-4a-S02' to form additional flavin 4a-sulfite adduct. This suggests that, if FMNH-4a-S03~ is a major active

283 species as we now propose, the D/H-initiated bioluminescence would be more efficient when the dithionite treatment of FMN is carried out in the presence of some oxygen rather than under anaerobic conditions. This predicted effect of oxygen has indeed been observed (Fig. 2, A and B). Moreover, the addition of sulfite to the sample during the treatment of flavin by dithionite under anaerobic conditions resulted in a significant enhancement

of the

bioluminescence induced by H 2 0 2 (Fig. 2, B and C). This is probably due to an additional FMNH-4a-SC>3" formation favored by a higher concentration of sulfite. Fig. 2. Effects of oxygen and sulfite on the efficiency of the dithionite/H202-initiated bioluminescence. To 0.23 mL buffer containing 8 ng luciferase, 0.23 mM decanal, 0.2 mM dodecanol, zero (A, B) or 3.5 mM sulfite (C), and 50 (iM FMN, a slight excess amount of dithionite was added under different conditions. The sample was immediately placed under nitrogen and the bioluminescence initiated by the injection of 0.1 mL deaerated (250 mM) H2O2 solution. (A): The dithionite treatment was carried out using an air-saturated sample solution. (B, C): The sample solution was first freed from oxygen before the dithionite treatment. One LU = 6.8 x 10 9 q-s-!.

so _ 3

20 = c c ~ ,0

w

0

Time

(min)

In another experiment, the 4a-hydroperoxyFMN intermediate II was formed with photochemically reduced FMNH 2 and isolated by Sephadex G-25 column chromatography. This intermediate II was immediately treated with an excess amount of sulfite and passed through another Sephadex G-25 column. The luciferase sample so obtained exhibited no bioluminescence upon reacting with decanal but the subsequent addition of H2O2 triggered an intense light emission. We propose that the intermediate II is converted to FMNH-4aSO3" by the sulfite treatment, thus losing the reactivity with aldehyde to emit light. The subsequent addition of H2O2 converts the flavin-4a-sulfite adduct back to the intermediate II and bioluminescent activity is resumed. The formation of luciferase:FMNH-4a-S03~ during dithionite treatment as documented in this report suggests that the usual preparation of luciferase "intermediate II" using dithionite would lead to the co-formation of the true intermediate II and the flavin-4a-sulfite. To tested this prediction, we have prepared (6) the "intermediate II" samples using reduced FMN obtained through both the photochemical and the dithionite reduction. The dithionitetreated "intermediate II" indeed showed a lower bioluminescence activity than the photochemically prepared sample upon reacting with aldehyde (Table 1). Furthermore, upon

284 Table 1. Relative Quantum Output of "Intermediate II" Prepared by Dithionite and Photochemical Reduction Relative Total Quantum Output (%) Flavin Reduction Photochemical Dithionite

-H2O2

+H2O2

+H2O2/-H2O2

100

102

1.0

77

109

1.4

Absorbance measurements at 380 and 450 nm were taken for all "intermediate II" samples obtained by Sephadex G-25 column chromatography. Quantum outputs were corrected for FMN contents and normalized according to flavin-4a-adduct contents. reacting with H2O2 and aldehyde, enhanced bioluminescence is only observed with the dithionite-treated "intermediate II" sample.

Acknowledgements: This work was supported by grants GM 25953 from NIH and E-1030 from the Robert A. Welch Foundation. We are deeply grateful to Dr. H. I. X. Mager for his catalytic role in this work.

References 1. Cho, K. W„ H. J. Lee, S. C. Shim. 1986. J. Korean Biochem. 19, 151. 2. Watanabe, H„ J. W. Hastings. 1987. J. Biochem. (Japan) KM, 279. 3. Watanabe, T„ T. Nakamura. 1976. J. Biochem. (Japan) 79, 489. 4. Hastings, J. W., S.-C. Tu, J. E. Becvar, R. P. Presswood. 1979. Photochem. Photobiol. 29, 383. 5. Mager, H. I. X., S. -C. Tu. 1990. Photochem. Photobiol. 51, 223. 6. Tu, S.-C. 1979. Biochemistry 18, 5940.

STRUCTURE OF FP3 9 0 INCLUDING ITS PROSTHETIC GROUP (Q-FLAVIN): PHYSIOLOGICAL SIGNIFICANCE OF LIGHT EMITTING REACTION IN LUMINOUS BACTERIA

Sabu Kasai,* Shigeru Fujii,** Retsu M i u r a , " Tadao Nakaya,* and Kunio Matsui****

Shoji Odani,***

* Faculty of Engineering, Osaka City University, Sumiyoshi-ku, Osaka, Osaka 558, Japan, * *Kansai Medical University, Hirakata Osaka 573, Japan, ***Niigata University School of Medicine, Niigata, Niigata 951, Japan, and * * * *Teikoku Women's University, Moriguchi, Osaka 570, Japan

Introduct ion In the previous symposium, we reported that biosynthesis of FP3 9 0 is induced in the cell of Photobacterium phosphoreum under the same conditions which cause luciferase induction (1) As a result of that study, it was inferred that a gene which codes for FP390 should be included in the lux operon. On the other hand, Mancini et al. found a new gene designated as lux in the lux operon (2) and the nucleotide sequence of the gene was determined by Soly et al. (3). Hence, it was postulated that the lux F is the gene coding for FP390. This is also supported by the fact that the gene does not code for any protein related to luminescence system and that the molecular weight of the lux F protein is similar to that of FP390 (1,3). Results and Discussion To demonstrate that the lux F gene codes for FP390, the protei was digested with CNBr and the digest was purified using HPLC. Five fragments (CB-I to V) were obtained and they were identical in number and size with fragments expected from the digest of lux F protein (see Fig. 1). Partial amino acid sequences of three large fragments (CB-I, II and III) were determined and they were identical with those of lux F protein as shown in Fig. 1. The amino acid compositions of FP390 and of the five CNBr fragments were determined and they were identical with those of lux F protein with one exception: a proline residue seemed to be exchanged with an isoleucine in

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin New York -Printed in Germany

286 OB-IV the largest > CBTV ^ fragment (CB-II). CB-I CB-11 CB-I11 It was concluded 26 130 205 215 231 from these results that lux F gene |CB-11 FP3s oMet-Asn-Lys-Trp-Asn-Tyr-Gly-V»1-Phe-Phe-V«1-Asn-Phecodes for FP3 9 o , 1 ux F ATG AAT AAA TGG AAT TAC GGA GTC TTC TTC GTT AAC TTT although in our Tyr-Asn-Lys-Gly-Gln-Gln-Glu-Pro-Sre-Lys-Thr-Met strain (IFO 13896) TAT AAT AAA GGC CAA CAA GAG CCA TCA AAA ACG ATG mutation seems to ICIMI FPj 9 QAsn-Asn-Alfl-Leu-Glu-Thr-Leu-Arg-lle-1le 1 ux F AAT AAT GCA TTA GAA ACA TTA CGT ATT ATT occur at least at one point when ICB-llll FP3 g qLeu-His-Va1-Asn-Va1-Asn-Glu-Ala compared with the luxF TTA CAT GTC AAT GTT AAT GAG GCA Fig. 1 CNBr digestion profile of FP39o strain NCMB 844 (above) and comparison of partial amino which was studied acid sequences with nucleotide sequences of by Meighen's lux F (below) group. FP390 contains a flavin prosthetic group, Q-flavin (QF). The previous preparation of OF seemed to be completely pure but it gave an NMR spectrum of a mixture of two flavins. We tried to further purify QF under quite mild conditions. QF was released from the apoFP3 9 0 using guanidine • HC1 and purified using chromatographic conditions at neutral pH. However, an NMR spectrum of newly prepared QF was mostly identical with that of the old one. The major component was designated as QFi and the minor was QF2. QFs (QFi + QF2 ) are phosphate esters and they were hydrolyzed to free forms (QFi-P + QF2-P) by the action of a phosphatase. QFi-P was separated from QF2-P by HPLC using a reversed phase column. On the FD mass spectrum of QFs, base peak was found at m/z=683 and (M+l)* ion peaks of QFi-P and QF2-P were found at the same mass number, m/z=603 on the mass spectra. Therefore, both QFi and QF2 are monophosphates. Because QFi-P and QF2-P have very similar properties and the same molecular weights, these compounds seem to be isomer. C, H, N-composition of a mixture of QFi-P and QF2-P was as follows; C, 58.20; H, 7.54; N, 8 . 6 0 % . These results give for each compound an empirical formula C3 1 H4 6 N4 Os • 2H2O which requires C, 58.29; H, 7.89; N, 8.77%. The formula suggests that these compounds consist of a molecule of riboflavin (C17H20N4O6) and one of myristic acid (C14H28O2). 1 H - N M R spectra of QFi-P and QF2-P are shown in Fig. 2; only one aromatic proton was found in both spectra. NOE was observed at protons of ribityl and methyl groups when the aromatic proton was irradiated. From these data, it was concluded that myristic acid moiety binds to riboflavin moiety at position 6

287 of the isoalloxazine ring. On the other hand, positions where riboflavin moiety binds to myristic acid moiety were determined by using COSY NMR. On the 1 H-NMR spectrum of QFi-P, resonance of two a methylene

J uukjj

I

vĄ .

i opm protons of myristic acid moiety showed sharp double doublets [6 =3.11 (1H, dd, J = 6 and 8 Hz) and 3.39 (1H, dd, J = 8 and 9 Hz)]. Resonance of these methylene protons showed spin couplings with a /3 methine proton [

[ETFox]

[dHox-P]

[ETFxJ

whereas the free reduced dehydrogenase, EFl2e , is some 30-fold less reactive towards ETF under comparable conditions (6).

Thus product binding markedly accelerates interflavin

electron transfer, despite rendering the dehydrogenase a significantly poorer reductant of ETF in thermodynamic terms (5,6).

This kinetic effect is not unique to ETF, but is shown

by a number of non-physiological acceptors of the dehydrogenase (10; e.g. phenazine methosulfate, ferricyanide and ferricenium salts).

Since ferricenium hexafluorophosphate

(FC+PF6~) proved a facile oxidant of the medium chain enzyme (11), it was selected for detailed study.

A comparison of

pseudo-first order rate constants for reoxidation of substrate and chemically reduced dehydrogenase are collected below (using 200 ^M Fc*PF6" in 100 mM HEPES buffer, pH 7.6, 1°C):

k2

40/sec [dH2e-p]

[dH le P]

[Fc'PF6]

^ [ FC*PF 6 " ]

3/sec [dH2B] [FC*PF6-]

200/sec [dH ox P] [Fc]

0.3/sec [dH0J

[dHla] [Fc]

[Fc*PF6-]

[Fc]

302

The effect of product is manifest in both phases of reoxidation but are most marked at the semiguinone level (13- and 700-fold faster for ka and k2 respectively). The dependence of the observed rate constants on Fc+PF6~ concentration is consistent with a weak precursor complex between dehydrogenase and ferricenium ion, but such interactions could not explain the kinetic modulation summarized above (10). Obviously one might invoke conformational changes induced by product binding which allow a closer approach, or a more favorable alignment of redox centers to explain these rate enhancements, but it is perhaps surprising that these effects are shared by physiological and artificial acceptors. A more direct effect on the chemical reactivity of the flavin is suggested by studies with a number of acyl-CoA analogues which, when bound to the reduced enzyme, generate rate enhancements of from 2- to 30-fold over the free E.Fl2e (10). Clearly then these rate accelerations are not unique to substrate reduced enzyme, neither are they merely a reflection of ligand binding. There does however appear to be a correlation between those ligands which are most effective in accelerating electron transfer and their ability to lower the pK of the blue semiquinone into the physiological pH range (10,12). An explanation of this phenomenon is ventured below. Redox communication between acyl-CoA dehydrogenase and ETF appears to involve obligatory 1-electron steps, and probably proceeds via outer-sphere processes (see e.g. 13) from the comparatively buried dehydrogenase flavin (7) to the exposed dimethylbenzene edge of the FAD in ETF (14). Since electron transfer itself is so much faster than any internuclear motions of reactants or solvent, adjustments in these parameters need to be made prior to electron transfer if the activation energy of the overall process is to be minimized (13). One factor contributing to the overall activation energy of electron transfer would be the protonation state of sites on the isoalloxazine whose pK values are strongly

303

dependent on the redox state of the flavin.

Scheme 1

illustrates various protonic states (horizontal) and redox states (vertical) of free flavin for illustration.

PROTONATION STATE 2H

+

1H+

0H +

pK 6.7

REDOX STATE —1 ro ® ®

FIH2

FIH "

I ! I ! P* 2

FIH^

pK 8.3

FIH-

^-r

o a>

pK

FIH* ox

F! ;

0

Flox

Scheme 1. Thermodynamic cycle showing selected protonated and redox states of free flavin. Preferred routes for 1-electron transfer reactions are in bold. pK values are from Muller (15) . First consider the neutral free dihydroflavin (F1H2; top left) as a potential 1-electron outer sphere reductant. Direct electron transfer would yield the energetically unfavorable semiquinone state (FlH2- n
6 Kcal) was determined using the stopped-flow spectrophotometer. TTie effect of this pK shift on the shape of the potential vs pH profile is two fold. First, the pH independent flat portion of the sq/red line is extended to higher pH, well beyond the normal measurement range. Second, the 60 mV slope portion of the ox/sq line is extended out to pH 12. F i g u r e 3B The measured potential of the ox/sq couple of flavodoxin (-125 mV in M. elsdenii flavodoxin at pH 7.0) is raised by -125 mV ( - 3 Kcal) relative to that of free flavin at pH 7.0. This rise in p o t e n t i a l m e a s u r e s the stabilization of the semiquinone relative to the oxidized form of flavodoxin. The effect of this added stabilization is to shift all the ox/sq lines to more positive potentials. In the absence of a more definitive measure, we have chosen to place the sq/ox line at the position found with M. elsdenii flavodoxin. The energy attributable to the new hydrogen bond at N(5) is of the same order of magnitude as this potential shift. The hydrogen-bond is retained in reduced flavodoxin. Although its strength may change somewhat with reduction, we have not adjusted the vertical position of the lines representing the sq/red potentials.

0

5

pH 10

In reduced M. elsdenii and C. beijerinckii flavodoxin, the pK of N(l) changes from a value of 6.7 for free flavin to a much lower value that cannot be measured directly. NMR (4) and visible spectra (1) show that the pK is below 4.0. The x-ray structure of 1-deaza-FMN flavodoxin shows that addition of a hydrogen at the 1-position causes a perturbation of the

402 protein and suggests that steric hindrance is responsible for the low pK of N(l) in reduced FMN flavodoxin (1). There is a pK=5.8 for reduced FMN flavodoxin found in the sq/red-potential vs pH profile (5). From the preceding paragraph, it is clear that this pK cannot be attributed to the reduced flavin. We have assigned it to the glutamate in reduced flavodoxin which is near N(3). In the oxidized and semiquinone forms of flavodoxin, this glutamate pK is much lower. A value of 3.8 can be calculated from the following thermodynamic cycle: pKi is for reduced 1-deaza-FMN bound to flavodoxin in the dFl^H dFlyH presence of the protonated glutamate and is taken to be equal GluH Glu© to the value of 5.6 for reduced 1-deaza riboflavin free in solution pKj = 5 . 6 p « . =7.6 (6). PK3 is the measured, nonflavin pK which we assigned to ,, the ionization of the glutamate in (JF1 r © the presence of the anionic, dFl£© reduced flavin. pK; 4 is the Glu© GluH measured pK for ionization of pK3=5.8 reduced 1-deaza-FMN bound to flavodoxin in the presence of the anionic glutamate £1). The remaining value pK2=3.8 is calculated, and reflects the pK of the glutamate in those forms of flavodoxin where the flavin has no net charge, ie oxidized and semiquinone. p K 9 =3.8

Placing an anionic reduced flavin next to an ionized glutamate generates a destabilizing charge repulsion in reduced flavodoxin. One consequence of this charge repulsion is an increase in the pK of the glutamate by at least 2 units (3.8 to 5.8). Since no visible evidence for the ionization of the flavin appears above pH 4, the upper limit for the pK of the reduced flavin under any conditions must be 4.0. According to the following thermodynamic cycle, in the presence of uncharged glutamate the pK is lowered to at least pH=2:

FlyH

p K 2 = 3.8

GluH

Fl v r H Glu©

p K 1 = 2.0 r © v GluH

p K 4 = 4.0

PI M

F rI R

PK3 = 5.8

©

'v Glu©

pK2 is the pK for the glutamate ionization in the presence of a neutral flavin, calculated from the previous thermodynamic cycle involving reduced 1-deazaflavin. PK3 is the observed pK of the glutamate in the presence of the reduced anionic flavin. pK4 is the upper limit, pK=4.0, for the reduced flavin. pKi is then the calculated, limiting value for ionization of the flavin in the presence of a neutral glutamate.

403

Sqfc2'

>

£ §"

\jedH3

\

v \

\

• \

I

• •

i •

SqH2

\

RedH2~

\ \ V \

2

.

_

O xJ,H \ SqH2 c - 4

o

Ox"\-

a.

a

- 6

.

• i

5

0

pH

?qH2

10

B OxH-Glu

Figure 4A This shows the effect of the N(1)H pK shift on the potential vs pH profile. The pK shift extends the pH independent, flat portion of the sq/red line down to pH=2. Figure 4B Charge repulsion raises the pK of the glutamate near N(3) in reduced flavodoxin. Between the pK of 3.8 in the semiquinone and the pK of 5.8 in the reduced form, the potential decreases with a slope of -60 mV/pH unit. In this range, oneelectron reduction of the flavin is linked to protonation of the glutamate. The result is a more negative potential for the sq/red couple, reflecting the destabilization of fully reduced flavodoxin relative to semiquinone. The final potential vs pH profile which we have calculated is qualitatively the same as that measured for M. elsdenii flavodoxin (dotted lines).

5qH2-Glu" In view of the approximations involved in the calculations described here, the calculated

S qH2—GluH

profile is pleasingly close to the H2 — —GluH

measured one. We have taken the rise in potential of the ox/sq couple to be equal to that actually

SqH2—Glu

observed

RedH2~—GluH RedH2~-Glu_ -i—i—i—i—I—i—•—i i I i i

5

pH

10

with M.

elsdenii

flavodoxin (~3 Kcal). The large observed shift in pK ( - 6 Kcal) corresponds to about twice the observed shift in potential. This

extra energy includes factors like charge repulsion and may also be used in part by the protein for adjustments to the structure. The shift in pK for N(1)H is large (ApK=4.7) but not unprecedented (7). Charge repulsion-induced destabilization of fully reduced flavodoxin

404 would account for 60% of the observed shift in the sq/red potential in the approximate model described here. If the pK=3.8 of the glutamate (in the presence of neutral flavins) were 1.0 unit lower, 85% of the observed shift could be explained by this interaction. Nevertheless, other factors are also likely to be involved in destabilizing the reduced flavodoxin. For example, additional charge repulsion between the reduced flavin and the 15 negatively charged amino acids (pH 7) distributed over the surface of the protein or the phosphate moiety of FMN could contribute to the destabilization of reduced flavodoxin (8). From the results of this exercise, we feel justified in arguing that the ability of the apoprotein to influence the pKs of the flavin is an important element in the mechanism by which proteins manipulate the flavin potentials. Acknowledgement This work was supported by U.S. Public Health Service Grants GM-111106 (V.M.) and GM-16429 (M.L.L.). References 1. Ludwig, M.L., L.M. Schopfer, A.L. Metzger, K.A. Pattridge, V. Massey. 1990. Biochemistry. Submitted. 2. Draper,R.D., L.L.Ingraham. 1968. Arch.Biochem.Biophys. 125, 802. 3. Fersht, A.R., J-P. Shi, J.Knill-Jones, D.M. Lowe, A.J. Wilkinson, D.M. Blow, P. Brick, P. Carter, M.M.Y Waye, G. Winter. 1985. Nature 214, 235. 4. Franken, H-D., H. Rüteijans, F. Müller. 1984. Eur.J.Biochem. J3§. 481. 5. Mayhew, S.G., G.P. Foust, V. Massey. 1969. J.Biol.Chem. 244, 803. 6. Spencer, R„ J. Fisher, C.Walsh. 1977. Biochemistry 16, 3586. 7. Lewis, S.D., F.A. Johnson, J.A. Shafer. 1976. Biochemistry 15, 5009. 8. Moonen, C. 1983. Thesis, Landbouwhogeschool, Wageningen.

POLARIZED

ABSORPTION

SPECTRA

OF

FLAVIN

MONONUCLEOTIDE

IN

FLAVODOXIN CRYSTALS

Louise Karle Hanson Department of Applied Science, Brookhaven National Laboratory, Upton, NY 11961 Garrott W. Christoph, James Hofrichter Laboratory of Chemical Physics, NIDDK, National Institutes of Health, Bethesda, MD 20892 Martha L. Ludwig Biophysics Research Division, University of Michigan, Ann Arbor, MI 48109

Introduction Although it has long been recognized that protein environments modulate the electronic structure of flavins, the precise nature of the interactions that lead to a specific effect are still poorly understood.

Detailed knowledge of flavin environments

from high resolution crystal structures of flavoproteins, coupled with spectroscopic measurements and molecular orbital calculations should begin to clarify the relative importance of the various possible interactions in a given system. Changes

in optical spectra reflect changes in the

structure.

electronic

Electronic transitions are characterized by their

energy, intensity, and transition dipole moment direction, /x. A

comprehensive

description

of

the

effect

of

the

protein

environment requires a knowledge of all three parameters.

The

first two are readily obtained from isotropic solution spectra of flavoproteins, the third requires oriented samples.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

406 Results Figure crystals

1

presents of

polarized

oxidized

absorption

and semiquinone

spectra

for

(SQ) Anacvstis

single nidulans

flavodoxin (Fd) (1) . These spectra complement results previously reported for the oxidized and semiquinone forms of Clostridium M P Fd (2).

The uniaxial symmetry of the C. M P crystals resulted

in two possible choices for the n of the flavin a and fi absorption bands, given in Table I.

The spectra obtained from the A.

nidulans oxidized crystals resolves the ambiguity because the two Fds crystallize in different space groups.

Oxidized

Semiquinone

Figure 1. Absorption spectra of oxidized (left) and semiquinone (right) A. nidulans Fd in solution (top) and in the Oil face of a single crystal (bottom) with P2^2,2, symmetry. The crystal absorption spectra were obtained with polarized light: E || a axis means that the polarization vector of the incident light w a s parallel to the a axis; E l a axis means that the polarization vector was perpendicular to the a axis. These two directions correspond to the extinction axes, a and 0 refer to the a and 0 absorption bands of the flavin.

407 The transition dipole moment directions, fi, are determined from the polarization ratio, PR, where PR = the OD for E || a axis divided by the OD for E i a axis. the n lie to the a axis.

The larger the PR, the closer

For each absorption band, the A.

nidulans data also yield two choices for n, however,

for the

oxidized Fd one of these corresponds uniquely to one of the C_j_ MP numbers.

The unique values are given in Table I.

Semiempiri-

cal INDO/s molecular orbital calculations on oxidized isoalloxazine in vacuo predict n's within several degrees of the observed values (3). The analysis in the previous paragraph assumes that the fi's will not be affected by the protein environments. found to be valid semiquinones. semiquinone

for the oxidized

This assumption was

species, but not for the

In contrast to the C. MP data, the A. nidulans

spectra

cannot

be

fit with

the

The lack of absorbance

same M ' s as

the

oxidized

spectra.

in the E i a axis

spectrum

(Figure 1) indicates that a significant change in the

orientation of /* has occurred, especially for the a band. resultant

values

of n are given

rotation of 10-15° or more.

in Table

I. They

The

reflect

a

In this case the two-fold ambiquity

cannot be resolved. Table I.

Transition Dipole Moment Directions.

Oxidation

Band

State Oxidized

Semiquinone

C. MP

A. nidulans

(2)

Cal 'd (3)

a

15 or 34 ± 4°

15°

14°

ß

-5 or 53 ± 4°

-5°

2°

a ß

16 or 33 ± 4°

-0 or -34°

-7 or 55 ± 4°

-4 or -38°

1

The angles are relative to the long flavin axis, which lies 17° off the a axis in the A. nidulans crystals. A positive sign denotes a clockwise rotation.

408 Conclusions The polarized absorption experiments on flavodoxin crystals have demonstrated that the protein environment can shift the flavin absorption

maxima

without

affecting

the

transition

dipole

direction

(the a band of the oxidized A. nidulans Fd is red

shifted),

and,

conversely,

can

rotate

the

transition

dipole

direction without affecting the transition energy (the absorption maxima of the two semiquinone Fds occur at the same wavelengths, although effects

their are

intensities

oxidation

state

are

somewhat

dependent.

different). Factors

These

within

the

protein environment that may contribute to the behavior of the optical transitions in Fds include aromatic amino acid residues and charged groups.

Calculations on model systems suggest that

the extent of the interaction depends on the relative orientation of these groups to and their distances from the

isoalloxazine

ring (3).

Acknowledgment This research was supported by an approved Exploratory Research Proposal under the auspices of DOE contract No. DE-AC02-76CH0016 (LKH) and by NIH grant No. GM-16429 (MLL).

References 1. Smith, W.W., K.A. Pattridge, M.L. Ludwig, G.A. Petsko, D. Tsernoglou, N. Tanaka, K.T. Yasunobu. 1983. J. Hoi. Biol. 165. 737. 2. Eaton, W.A., J. Hofrichter, M.W. Makinen, R.D. Anderson, M.L. Ludwig. 1975. Biochemistry 14, 2146. 3. Hanson, L.K.. 1990. J. Mol. Struct, (in press).

STRUCTURAL ANALYSIS OF FULLY REDUCED A. NIDULANS FLAVODOXIN

C.L. Luschinsky, W.R. Dunham, C. Osborne, K.A. Pattridge, and M.L. Ludwig Biophysics Research Division and Department of Biological Chemistry, The University of Michigan, Ann Arbor, Michigan

Introduction

Anacystis nidulans flavodoxin (169 residues) is the structural prototype for "long-chain" flavodoxins (1).

Comparison with the smaller flavodoxin

from Clostridium beijerinckii (139 residues) shows that the interactions between the flavin and adjoining residues differ in the two structures (1,2).

Thus the mechanisms that these members of the flavodoxin family use

to control the redox potentials of the bound flavin may not be identical. In C. beijerinckii (aka C. MP) flavodoxin, formation of a hydrogen bond between a backbone carbonyl and N(5)H of the flavin is important in stabilizing the semiquinone species (3), and electrostatic interactions have been implicated in control ,of the sq/red potential (4,5).

In this report we describe the structure of the FMN binding site in fully reduced A. nidulans flavodoxin.

The conformation we observe in reduced A.

nidulans flavodoxin is a departure from the model for C. MP flavodoxin. The interactions that affect the orientation of the peptide in question (i.e. 58-59) in the fully reduced A. nidulans flavodoxin favor a conformation in which the carbonyl points away from the flavin, rather than toward N(5)H as in flavodoxin from C. MP.

We also report the construction

of mutants that will be used in further studies of the energetics of FMNprotein interactions in A. nidulans flavodoxin.

Flavins and Flavoproteins 1 9 9 0 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

410 Results

A crystal of recombinant A. nidulans flavodoxin (6) was reduced using a concentrated dithionite solution in buffered ammonium sulfate at pH 8 under anaerobic conditions, and data were collected on an area detector, with an R

sym Of 0.048.

is 5-fold.

The data set is 99.2% complete to 2.2 A, and the redundancy

To determine the oxidation state of the crystal at the end of

the data collection, EPR spectra were measured. After exposure to oxygen, the radical signal rose and then disappeared (Figure 1), indicating that the crystal had been fully reduced throughout the data collection.

-l I—I

1

1

1

-

1 1

1

r

1

n—i—i

r

1—

1.5

1.0

i

io

—I

0.0

-0.5 -20

L. 330 Hogn«t>o Fi.ld (.1)

0

1 1

0

0

1

1 1

0.5

c ia

1

20

40

i A I 1 60

80

100

Time After Exposure To Oxygen (min)

FIGURE 1. Appearance and decay of semiquinone signal after oxygenation of the flavodoxin crystal. The eventual disappearance of the signal after oxygenation was accompanied by the appearance of the characteristic yellow color of the oxidized protein. The inset shows the room temperature flavodoxin signal generated by difference techniques. Quantitation of the EPR signal required estimation of crystal volume and calibration of the spectrometer sensitivity with ImM copper perchlorate. We estimate the errors associated with these measurements at 25%.

411 We began the data analysis by computing difference maps with amplitudes (|Fredl - |F0X|).

By analogy with the clostridial flavodoxin, we expected

to find peaks indicating that the peptide connecting Asn 58 and Val 59 was oriented to form a hydrogen bond between the N(5)H of FMN and the backbone O 58 (i.e. in the O "up" conformation), but the maps did not support thi3 interpretation.

Maps were then computed with amplitudes |Fre S Q ) Wildtype -146 Y98F -148 Y98W -152 Y98H -175 -173 Y98R Y98M -206 Y98A -178 valúas in mV vs N.H.E, p H 7.0

E

Flavodoxin

transfer interactions.

(SQ

1

>HQ) -443 -414 -449 -268 -272 -302 -304

In contrast, tryptophan-90 w h i c h occupies a

generally equivalent location in the CI. beljerinckii flavodoxin is not coplanar w i t h the FMN nor is a long wavelength absorbance band observed (1).

The stability of such complexes is apparently dependent on the redox state of the flavin (11).

For example, tyrosine has been reported to associate

more strongly w i t h the flavin semiquinone than the oxidized or fully reduced forms, while phenylalanine and histidine (imidazole) apparently show little preference.

Tryptophan seems only to bind to oxidized flavin (12).

O n the

basis of these observations, one might anticipate that the midpoint potential of the OX/SQ redox couple (E2) to be dependent on the nature of the aromatic residue at position 98 in this flavodoxin.

However, the redox

potential of this couple was found not to be appreciably affected by such a substitution.

In fact, all mutations w h i c h retain aromaticity (i.e.

Y98W,

Y98F) resulted in potentials nearly identical to wildtype protein (Table I). However, the presence of an aromatic residue at this location does apparently stabilize the flavin semiquinone.

Substitution of non-aromatic,

neutral amino acid residues such as methionine and alanine or positively charged amino acid residues such as histidine (assumed to be at least partially charged at p H 7) and arginine at position 98 result in significantly more negative midpoint potentials for the OX/SQ redox couple.

Pronounced effects on the midpoint potential of the SQ/HQ couple (E^) were noted for all substitutions introduced at position 98 except tryptophan (Table I).

The midpoint potential for the Y98W mutant was comparable if not

slightly m o r e negative than wildtype.

This observation is consistent w i t h

419

reports that in solution, where charge transfer-type interactions are noted, complex formation with tryptophan significantly decreases the redox potential of FMN (12).

This and more direct solution studies provide

evidence that indoles form complexes with oxidized FMN but not FMNH2.

In a

charge transfer complex, where the electron-rich indole serves as the electron donor and the FMN the acceptor, reduction of the flavin would reduce its capability as an acceptor, significantly weakening the complex. The very negative SQ/HQ potentials for both wildtype (tyrosine) and Y98W mutants reflect a weaker association of the fully reduced FMN with the apoprotein than the more oxidized forms. The Y98F mutant has a midpoint potential for the E^ couple which is significantly more positive than either wildtype or Y98W.

Phenylalanine

would be expected to provide a less electron-donating environment when adjacent to FMN than tyrosine and tryptophan.

This might result in a

more energetically favorable situation for the introduction of a second electron into the flavin isoalloxazine ring, especially in the flavodoxin where the FMN hydroquinone is known to retain a formal negative charge (1). Indeed, electrostatic repulsion has been proposed as an explanation for the poor association of FMNH" to flavodoxin and, thus, the very negative redox potential of SQ/HQ couple (13).

The absence of compensating charges in the

vicinity of the flavin hydroquinone anion, the negative surface potential surrounding the flavin, and the presence of the uncompensated phosphate dianion on the ribityl side chain of the cofactor are all thought to contribute to this repulsion.

Mutagenesis experiments producing the Y98R

and Y98H mutant flavodoxin were designed to test this idea.

Indeed, as

shown in Table I, the presence of positive charge adjacent to FMNH" resulted in a large increase in the midpoint potential of the SQ/HQ couple.

(Note:

This argument assumes that the histidine is at least partially protonated at pH 7 in this negatively charged environment.

Experiments to determine the

pKa value of this histidine residue and to determine the pH dependency of the redox potentials of Y98H and other mutants are currently in progress.) Substitution of the aromatic side chain at position 98 with neutral aliphatic residues also results in a substantial increase in the midpoint

420 potential of the SQ/HQ couple (i.e. the Y98A and Y98M mutants); however, the extent of this increase is less than for Y 9 8 R and Y98H.

Molecular graphics

models of the Y98A mutant suggest that the FMN would have a significant portion of the isoalloxazine ring exposed to the high dialectric environment of solvent (assuming that its structure is isomorphous to wildtype protein).

The effects of similar substitutions at the equivalent position in the CI. beijerinckii flavodoxin (tryptophan-90) are also under

ii. Glycine-57 of Clostridium beijerinckii

investigation.

flavodoxin

In the flavodoxin sequences thus far established, a glycine residue is almost invariably observed in the second position within the bend situated near the C(4)-N(5)-C(6) edge of the flavin (1).

It is this loop that

undergoes a conformational change upon reduction w h i c h results the rotation of the peptide backbone carbonyl group of glycine-57 and the apparent formation of a new hydrogen bond between the NH(5) of the flavin semiquinone and the carbonyl oxygen (1).

Because of the bond angles involved, a glycine

residue is more acceptable at this position in the turn (3).

In contrast, an

asparagine residue is found at the equivalent position in

the flavodoxin from A. nidulans (3).

The redox potential for the OX/SQ

couple for this protein is significantly more negative than CI. beijerinckii (1).

These observations have lead to the hypothesis that the

OX/SQ potentials may be a function of the relative conformations of the NH(5), 0(4) and the backbone carbonyl oxygen or conformation energy contributions dictated by the preferred configuration of a glycine residue within this bend as opposed to an asparagine residue or other residues having straight or branched side chains (3).

(For a more complete

discussion, see a related article in this volume (7)).

In an attempt to test this hypothesis, several single-site mutations were introduced within this portion of the coding region in the synthetic gene for the Clostridium flavodoxin using the cassette mutagenesis approach. Glycine-57 has been substituted w i t h an alanine, asparagine, aspartate, and a threonine residue.

The substitution of an alanine residue introduces the

421 Table II. Oxidation-Reduction Potentials and Calculated Ratios of Association Constants for CI. beijerinckii Flavodoxin Mutants Flavodoxin 300 41 46 19 0.3

- 92 -143 -IAO

Wildtype G57A G57D G57N G57T

-162

-270

a

values in mV vs N.H.E, pH 7.0 ^calculated using an Em, 7 for FMN of -238 mV ( D

minimal side chain, an asparagine would duplicate, in principle, the situation in A^ nidulans, an aspartate tests the additional effect of charge, and a threonine introduces a small hydrophilic residue with a branched side chain.

These residues, having side chains of varying

properties, might be expected to interfere with either the conformation change itself or to alter the conformational energy of this bend and/or hydrogen bonding strength between the carbonyl oxygen of this residue and NH(5) of the flavin semiquinone.

In particular, it was reasoned that the

branched side chain of threonine might interfere sterically with the rotation of the adjacent carbonyl group upon reduction of the flavodoxin to the semiquinone. As the preliminary redox potential data provided in Table II suggest, each of the mutant proteins exhibit thermodynamically less stable blue neutral flavin radicals.

Particularly striking is the G57T mutant which displays a

midpoint potential which is approximately 180 mV more negative than wildtype.

The midpoint potential of the G57N mutant, while more negative

than the wildtype protein, is not as negative as in the A. nidulans flavodoxin (Em,7 = -221 mV)(l).

Included in Table II are the ratios of the

association constants for the oxidized and one-electron reduced FMN as calculated from the observed redox potentials.

These data suggest that the

presence of a side chain at position 57, even as small as a methyl group, may significantly alter the interactions between protein and cofactor which stabilize the semiquinone complex. The structural consequences of these amino acid substitutions as determined by X-ray crystallographic analysis of these site mutants and their contributions to the changes in the observed

422 redox properties of the bound FMN cofactor are discussed in greater detail elsewhere in this volume (7).

Acknowledgements This research was supported by a grant from the National Institutes of Health (GM36490).

The authors also acknowledge the support, encouragement,

and helpful discussions of Dr. Martha Ludwig of the University of Michigan. References 1. Mayhew, S.G. and Ludwig, M.L. (1975) in The Enzymes (Boyer, P.D., ed.), 3rd edn., Vol 12, pp 57-118. 2.

Watenpaugh, K.D., Sieker, L.C., and Jensen, L.H. (1973) Proc. Natl. Acad. Sci. USA 70, 3857.

3.

Laudenbach, D.E., Straus, N.A., Pattridge, K.A., and Ludwig, M.L. (1987) in Flavins and Flavoproteins (Edmondson, D.E. and McCormick, D.B., eds), Walter de Gruyter & Co., pp. 249-260.

4.

Fukuyama, K., Wakabayashi, S., Matsubara, H., and Rodgers, L.J. (1989) Abst. M141, 3rd Symposium of The Protein Society, Seattle.

5.

Krey, G.D., Vanin, E.F., and Swenson, R.P. (1988) J. Biol. Chem. 263, 15436.

6.

Eren, M. and Swenson, R.P. (1989) J. Biol. Chem. 264, 14874.

7.

Ludwig, M.L., Pattridge, K.A., Eren, M., and Swenson, R.P. (1990), this volume.

8.

Watt, W., Watenpaugh, K.D., Tulinsky, A., and Swenson, R.P. (1989) Abst. PB55, Annual Meeting of American Crystallographic Assoc., Seattle.

9.

Helms, L., Krey, G.D., Swenson, R.P. (1990) Biochem. Biophys. Res. Commun. 168, 809.

10. Helms, L. and Swenson, R.P. (1990) manuscript submitted for publication. 11. Draper, R.D. and Ingraham, L.L. (1970) Arch. Biochem. Biophys. 139, 265. 12. Wilson, J.E. (1966) Biochemistry 5, 1351. 13. Moonen, C.T.W., Vervoort, J., and Muller, F. (1984) in Flavins and Flavoproteins (Bray, R.C., Engel, P.C., and Mayhew, S.G., eds.) Walter de Gruyter & Co., pp 493-496.

STRUCTURAL CHARACTERIZATION OF SITE MUTANTS OF CLOSTRIDIAL FLAVODOXIN

Martha L. Ludwig and Katherine A. Pattridge Department of Biological Chemistry and Biophysics Research Division, University of Michigan, Ann Arbor 48109 MI Mesut Eren and Richard P. Swenson

Department of Biochemistry, The Ohio State University, Columbus 43210 OH

Introduction In flavodoxin from Clostridium beijerinckii (aka C^ MP), the redox potential for the oxidized/semiquinone equilibrium is about 150 mV more positive than that for free FMN (1).

Comparisons of the structures of the oxidized and

semiquinone states suggest ways in which the semiquinone form may be preferentially bound, increasing E Q (2): formation of a hydrogen bond between the carbonyl oxygen of Gly57 and the N(5)H of the flavin semiquinone is an obvious stabilizing interaction.

Because reorientation of the peptide

bond between Gly57 and Asp58 occurs on reduction of oxidized clostridial flavodoxin, differences in the conformational energies of the oxidized and semiquinone proteins can also affect the observed potentials.

In oxidized

clostridial flavodoxin, Gly57 has ,

+

Q < CL
400

Km NADH

Vmax

NADH / DCPIP pH 8.0

|

558

3 3 (0 a a> E

CM

h c^-

o !g !c _ç t> 3 T3 O

O) ock_ CO o a> "a "a » c

a> T3 O

C

"O c

559 Results and discussion

In previous studies with the pig heart enzyme (3), it has been suggested that a histidyl residue is involved in proton transfer reactions during the catalytic cycle. In glutathione reductase, a histidine was identified in the active site together with a nearby glutamate residue which could act as a charge relay system in this manner. We have mutated this histidine (His450) into tyrosine, phenylalanine and serine respectively and the glutamate (Glu455) into glutamine and aspartate. Furthermore, the cis proline (Pro451), which allows the formation of a hydrogen bond between the backbone carbonyl of His450 and flavin N-3, was mutated into an alanine and a tyrosine (Tyrl6) in the substrate binding site into phenylalanine and serine. In all cases stable, dimeric enzymes were isolated, which were spectrally (absorption and far-UV CD) almost indistinguishable from wild type although the HF and HY mutants showed a somewhat more pronounced shoulder at 480 nm, indicative of a more apolar environment. The spectral properties of the PA mutant indicated a somewhat more polar environment. Table I shows the results of a steady state analysis of these mutants in the physiological reaction (Lip(SH) 2NH2 -> NAD1"), in its reaction with the NAD analog acetylpyridineadenine dinucleotide (APAD*) and in the diaphorase activity (NADH -> DCPIP) . The kinetic data were best fitted with a ping-pong mechanism, both in the wild type enzyme and in the mutants. The relatively small variation of the Km and Vmax values for the diaphorase activity indicate that the mutations did not influence appreciably reactions at the re face of the flavin, the only exception being the YF and YS mutants. With the EQ, ED, YS and YF mutants the physiological reaction could not be measured due to product inhibition (overreduction by NADH). The activity of the PA mutant could not be determined at all due to rapid irreversible inactivation under turnover conditions. With APAD+ as acceptor product inhibition was only observed above pH

560

300

NAD + )

ED m u t a n t ,

rapid

(lip(SH) 2 NH 2 In the

EQ

this

way

In

mutant these

enzyme. Spectral titration with

increase

in redox

potential

of

the

flavin. The rate of the r e d u c t i v e h a l f r e a c t i o n in ED w a s 25% of that

of t h e w i l d t y p e e n z y m e b u t

in EQ

it was d e c r e a s e d b y

f a c t o r of ten, s t i l l not rate l i m i t i n g . The r a t e l i m i t i n g is t h e t r a n s f e r

of r e d u c i n g e q u i v a l e n t s

a

step

f r o m the d i s u l f i d e

to

the f l a v i n a n d v i c e v e r s a . A g a i n no t r a n s i e n t 380 n m a b s o r p t i o n was o b s e r v e d . W i t h b o t h m u t a n t s , h i g h a b s o r b a n c e at 530 n m was observed

(fig

2B) ,

probably

also

e q u i l i b r i u m b e t w e e n EH2

species.

The

indicates

crystal

residue.

One

structure of the

oxygen

atoms

due

the

to

role

of the

a of

shift the

in

glutamate

carboxylate

anion

w i t h i n h y d r o g e n b o n d d i s t a n c e of N1 of H i s 4 5 0 . T h i s m a y in t h e

increase

of t h e p K a

of the h i s t i d i n e

the

a n d in a

is

result correct

o r i e n t a t i o n of the h i s t i d i n e - N 3 t o w a r d s the d i s u l f i d e . T h e o t h e r oxygen

can

form

a

hydrogen

bridge

with

a

NH-group

of

the

563 backbone (Ala452) . The EQ mutant has lost its polarizing and orienting effect on His450, but can still form a hydrogen bridge with the backbone. On the other hand, the results with the ED mutant show that it is probably still able to polarize the histidine and increase its pKa. Formation of a hydrogen bridge with N1 will lead to a distortion of the orientation of the histidine. Flexibility in the bound dihydrolipoamide may allow adaptation to this change in orientation, but the interaction with the interchange thiol will be diminished. Thus also these mutants indicate the important role of His450 in the transfer of electrons from disulfide to flavin. Probably, overreduction is the answer to such a distortion in this part of the molecule, as is also indicated by the overreduction found with the P451A mutant and the C-terminal mutant (see below). Besides the change in redox potential, competition between the rate of overreduction and of reoxidation at the level of EH2 will determine the outcome. Lipoamide dehydrogenase contains a 15 aminoacid C-terminal extension with respect to glutathione reductase. The last 10 aminoacids are not visible in the electron density map and represent probably a mobile structure on the outside of the molecule. Because this structure could be involved in interaction with the E2 component of the complex three deletion mutants were prepared with deletions of respectively 5, 9 and 14 aminoacids. A14 showed quite unexpected properties with respect to wild type enzyme: it dissociated easily (Kd 5 |1M) and lost FAD in this process. The spectral properties indicated a more polar environment and the flavin was easily quenched by I". This mutant shows the importance of the C-terminus for dimer stability. This was unexpected because the dimer contacts are far removed from this part of the molecule. At concentrations where the dimer prevailed, only weak binding to the E2 component of the complex was observed. This may be attributed to a change in the orientation of the subunits in the dimer, because other observations have shown that the dimer is required for binding. The other C-terminal mutants behaved normal in binding.

564 Therefore it is concluded that the C-terminal end of the chain has no direct function in the binding of E2. Docking experiments with dihydrolipoaraide, performed by Mattevi and Hoi (1), indicated a possible role for Tyrl6 in the binding of the substrate. A tyrosine (Tyr99) is found at a comparable position in E. coli glutathione reductase, but replacement by phenylalanine has apparently no effect (5). Tyrl6 was replaced by phenylalanine and serine respectively. The physiological activity could not be determined, again due to rapid product inhibition. With APAD+ as electron acceptor 20% of the wild type activity was obtained with normal Km for APAD+ and lip(SH)2NH2. Therefore an effect on the binding of Lip(SH)2NH2 seems unlikely. These mutants will be further analyzed.

Acknowledgement

This research was supported by the Netherlands Foundation for Chemical Research (SON) with the financial aid from the Netherlands Organisation for Scientific Research (NWO).

References

1.

Mattevi, A. and W.J.G. Hoi, this book.

2.

Westphal, A.H. and de Kok, A. (1988) Eur. J. Biochem. 172, 299-305.

3.

Matthews, R.G., Ballou, D.P., Thorpe, C. and Williams, C.H., Jr (1977) J. Biol. Chem. 252, 3199-3207.

4.

Thorpe, C. and Williams, C.H., Jr 251, 7726-7728.

5.

Deonarain, M.P., Berry, A., Scrutton, N.S. and Perham, R.N. (1989) Biochemistry 28, 9602-9607.

(1976) J. Biol. Chem.

LIPOAMIDE DEHYDROGENASE FROM AZOTOBACTER

VINELAND11

- KINETIC STUDIES ON

WILD TYPE AND MUTANT ENZYMES

Jacques A.E. Benen, Nicole A.H.M. Dieteren, Willem J.H. van Berkel and Arie de Kok. Department of Biochemistry, Agricultural University, Dreyenlaan 3, 6703 HA Wageningen, The Netherlands

Introduction Lipoamide dehydrogena se (lipDH) belongs to the family of flavin containing pyridine nucleotide-disulfide

oxidoreductases.

Its

closest

is

and

relative

glutathione

reductase

structurally

biochemically very similar (1). For both enzymes mechanisms have been proposed and essential residues were identified: except for the disulfide forming cysteines the 'most important' residue is a histidine that serves as a base during catalysis (2,3). Our

first goal was to demonstrate

'proton

accepting'

histidine

and

clear cut the role of the to assess

the

role

of

the

glutamate that forms a strong hydrogen bond with the histidine. Therefore protein

the mentioned engineering

residues were

techniques.

dependent steady-state kinetics of

replaced by

Here A.

we

report

vinelandii

others on

the

by pH

wild type and

mutant lipoamide dehydrogenases.

Results and Discussion Table I of the accompanying paper in this volume) wild type

(Benen et al., see elsewhere

shows the steady-state

kinetic properties of

lipDH and the H450 and E455 mutants

as they

were

determined at the pH optimum for each particular enzyme. Except for mutant E455D where the optimum shifted from pH 8.0 towards

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

566 pH 7.0 no drastic changes in pH optimum were observed. No shift towards higher pH values was observed in mutant H450S as might be

expected

lip(SH)2NH2

from

the

pK a

of

9.35

of

free

substrate

(4). For a better understanding steady-state kine-

tics were studied at different pH values for mutant

the

H450S

strong product

(see Fig. 1) . Except

inhibition/overreduction

observed at high pH values for wild type enzyme using NAD

was

+

and

E4 55D using APAD + . As a result the Km and Vmax values determined at pH values above pH 8.0

(pH 7.5 in case of E455D) are less

reliable. From the Log (Vmax) plots in Fig. la, b a pK a value of

6.3-6.5

was obtained for the rate limiting complex in both wild type and H450S. This value coincides with the pK a of the rate

limiting

EH2.NAD"1" complex reported for pig heart lipoamide dehydrogenase (5) . The pK a

can not be associated

with histidine. A

likely

candidate is the interchange thiol that is subject to deprotonation

during

reoxidation.

The

changes

of

Km

NAD +

and

lip(SH)2NH2 for wild type were too small to allow determination of

pK a

values

associated

with

substrate

binding.

It

is

interesting that Vmax remains constant in H450S between pH 8.0 and

pH 9.0

activity mainly

although

was

lower

at

non

at pH

be attributed

saturating

9.0. The

to an

increase

could be due to a worse binding

fixed

decrease of Km

concentrations of activity

lip(SH)2NH2-

as a result

of

can This

increase

of

negatively charged substrate. Fig.lc shows that Vmax for APAD + reduction by wild type enzyme reached no maximum in the experimental pH interval. Therefore no pK a could be assigned to a rate limiting intermediate. For wild type a pK a value of 8.1 was determined from the pKm APAD + plot representing an ionization of EH2 (since with APAD + a ping-pong mechanism applies). This pK a value of

His450.

histidine

A was

similar

value

for

might reflect the ionization the

reported by Matthews

pKa

of the

et al.

active

(2) . Mutant

site E455D

reached its highest Vmax already at pH 7.0-7.5. Beyond this pH strong inhibition was observed hampering correct determination

567

Measurements were done in triplicate at 25°C at each pH indicated. Reactions were started by the addition of enzyme. All buffers were adjusted to 150 mM in ionic strenght and contained 0.5 mM EDTA. Buffers used: pH 5.5-6.0-6.5, 100 mM MES; pH 7.07.5-7.8, 100 mM HEPES; pH 8.0-8.6, 100 mM HEPPS; pH 9.0, 100 mM CHES. Panel A: A. vinelandii wild type lipDH with NAD + ; panel B: mutant H450S with NAD + ; panel C: wild type lipDH with APAD+; panel D: mutant E455D with APAD + . Curves were drawn by visual inspection.

568 of kinetic parameters. The structural basis for this phenomenon might be twofold: a conformational form a hydrogen bond with the

change of the histidine

shorter Asp455 carboxylate

to

(see

accompanying paper by Benen et al.) and a downward shift of the pKa

of

the histidine

as

a result

of

greater

distance

to

the

electron rich carboxylate of Asp455. As a result a rise of the redox potential of the flavin in the EH2 .NAD + /APAD + complex has been

observed

favouring

overreduction.

The

involvement

of

an

ionizable group is evident from the strong pH dependence of the 'overreduction' this respect

in wild type by NADH and in E455D by APADH.

In

it is very interesting that this overreduction

is

not observed in H450S.

Acknowledgement s This

work

was

supported

Chemical Research

by

the

Netherlands

Foundation

for

(SON) with financial aid from the Netherlands

Organisation for Scientific Research

(NWO).

References 1. Williams, C.H. Jr. 1976. In: The Enzymes, 3rd ed. (P.D. Boyer, ed.) vol.XIII, Academic Press, New York, p. 89. 2. Matthews, R.G., D.P. Ballou, C. Thorpe, C.H. Williams, Jr. 1977. J. Biol. Chem. 2£2, 3199. 3. Pai, E.F., G.E. Schulz. 1983. J. Biol. Chem. 258. 1752 4. Williams C.H. Jr., N. Allison, G.C. Russell, A.J. Prongay, L.D. Arscott, S. Datta, L. Sahlman, J.R. Guest. 1989. Ann. N.Y. Acad. Sci. 573. 55. 5. Matthews, R.G, D.P. Ballou, C.H. Williams, Jr. 1979. J. Biol. Chem. 2 M , 4974.

BINDING STUDIES OF THE DIHYDROLIPOAMIDE DEHYDROGENASE COMPONENT (E3) IN THE PYRUVATE DEHYDROGENASE COMPLEX FROM AZOTOBACTER VINELAND11

E. Schulze, J.A.E. Benen, A.H. Westphal, W.J.H. van Berkel, A. de Kok Department of Biochemistry, Agricultural University, Wageningen, The Netherlands

Introduction The pyruvate dehydrogenase complex consists of multiple copies of three enzymes: pyruvate dehydrogenase (El), dihydrolipoyl transacetylase (E2) and (dihydro)lipoamide dehydrogenase (E3). In the complex from Azotobacter vinelandii El and E3 bind as dimers to the E2-component causing a dissociation of the 24meric E2-core into tetramers. The lipoamide dehydrogenase catalyzes the reoxidation of dihydrolipoyl groups which are covalently attached to the E2-component of the complex. The genes for the E3 from A. vinelandii (1), E. coli (2) and P. fluorescens (3) have been cloned and sequenced. From the three-dimensional model of the crystal structure of A.vinelandii E3 (4) it is known, that the substrate binding site of E3 is constituted from both subunits. The putative binding region for E3 on E2 contains many charged residues. It is also known, that E3 binds to E2 in a very mobile way (5). Therefore from the 3D-structure of lipoamide dehydrogenase from A. vinelandii two possible regions for the binding to the E2-core can be predicted: i) a helix on the surface of the E3-dimer with lysine in every fourth position (residues 221-236) and ii) the C-terminus of the enzyme.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

570 Results and Discussion The lysine in position 233 (in the helix mentioned above) and lysine 373 (situated in the vicinity of the helix) are conserved among lipoamide dehydrogenases but not among glutathione reductases. Therefore the mutants K233Q and K373Q were prepared and analyzed for binding on E2 by detecting the dissociation of the 24meric E2-core into tetramers on Superose-6 gel filtration (FPLC). The E3/E2-interaction was not changed for both mutants. Chemical modification of all accessible lysines on the surface of the E3-dimer decreased strongly the physiological activity and the enzyme started to aggregate. Even this form showed normal binding to E2. Therefore it can be stated, that lysine residues on the surface of the E3-dimer are not involved in binding to E2. In glutathione reductase, showing strong active site amino acid sequence identity with E3 but no binding to a core enzyme in an enzyme complex, the C-terminal tail of E3 (residues 460-476) is not present. In the X-ray structure of A.vinelandii E3 the C-terminus could not be clearly detected until now. This is possibly due to a high mobility (4). The importance of this C-terminal tail was checked by preparing a set of deletion mutants (A5, A9, A14). Some kinetic properties of these mutants are shown in table 1. Table 1: Relative activities of wild-type E3 and of the C-terminal deletion mutants. Enzyme

Lip(SH) 2/NAD+ activity [% wt]

wt

100

180

200

A5

110

200

300

5

70

250

-S^

• FAD •SH

>

-S-

Sc. J

s st 25 C is found to be 500 s _ 1 for X0 8_cl and 90 s - 1 for X0 8-SH . The wavelength dependence of the kinetics was consistent with intramolecular electron transfer and not reoxidation of the partially reduced enzyme species by contaminating oxygen (data not shown) . For both X0 8-cl and X0 8-SH , the rate constant for the observed transient is large relative to k c a t (20 s - 1 for X0 8-cl , 10 s _ 1 for X0 8_SH ; R. Hille and V. Massey, unpublished) .

SECONDS

Figure 1. Plots of fAA-Fe/S v s. fAA-FAD for X0 8_cl (circles; 550nm vs. 450nm) and X0 8 - S H (squares; 450nm vs. 580nm) at pH 6.0 (open symbols) and 10 (closed symbols).

Figure 2. Transients seen in the pH 10^6 pH jump for X0 8-cl (525 nm; Upper Panel) and X0 8_SH (450 nm; Lower Panel).

680 The temperature-dependence of the two pH jump experiments give activated energies of 8.7 and 8.8 kcal/mol for X0 8 - c l and X0 8_SH , respectively. These values are significantly smaller than observed with native enzyme at the same pH (19.8 kcal/mol; ref. 4), but quite high for simple electron transfer events and consistent with the involvement of prototropic equilibria in the redox equilibration process (expected since proton uptake must be associated with the uptake of reducing equivalents by FAD) . Our results demonstrate that electron transfer remains fast relative to turnover in XO containing 8-chloro or 8-mercapto FAD, and that by appropriately varying the FAD reduction potential the Fe/S center involved in the observed equilibration process driven by pH jump can be changed (Fe/S I in the case of native XO and X0 8-SH , Fe/S II in the case of X0 8-cl ) . The rapid equilibration rate between Fe/S II and the FAD of X0 8 - c l cannot be entirely accounted for simply by the larger thermodynamic driving force of electron transfer.

References 1. Hille, R., V. Massey. 1985. In: Molybdenum Enzymes (T.G. Spiro, ed.), John Wiley & Sons, New York, pp. 443. 2. Bhattacharyya, A.,G. Tollin, M. Davis, D.E. Edmondson. 1983. Biochem. 22, 5270. 3. Edmondson, D.E., J.T. Hazzard, G. Tollin. 1987.In: Flavins and Flavoproteins, (Edmondson, D.E., and McCormick, D.M., eds.) Walter de Gruyter, New York, 403. 4. Hille, R., V. Massey. 1986. J. Biol. Chem. 261, 1241. 5. Anderson, R.F., R. Hille, V. Massey. 1986. J.Biol.Chem. 261. 15870. 6. Porras A.G., G. Palmer. 1982. J.Biol.Chem. 257, 11617. 7. Hille, R., J.A. Fee, V. Massey. 1981. J.Biol.Chem. 2J2Ü, 8933.

STUDIES OF ELECTRON TRANSFER IN XANTHINE OXIDASE Russ Hille and Robert F. Anderson Department of Physiological Chemistry, Ohio State University, Columbus, OH 43210; and the CRC Gray Laboratory, PO Box 100, Mount Vernon Hospital, Northwood, Middlesex HA6 2JR

Introduction Xanthine oxidase (XO) contains molybdenum, two 2Fe/2S ironsulfur centers and FAD in each of its two identical subunits (1). Xanthine reduces the enzyme by two electron equivalents at the molybdenum center; the electrons are removed by O2 at the FAD (2,3) . Electron transfer between the molybdenum and FAD, presumably mediated by the Fe/S centers, is thus an integral aspect of the catalytic cycle, and XO serves as a useful system in which to examine electron transfer in an entirely biological context. Previous work has indicated that radiolytically generated viologen radicals introduced reducing equivalents into XO at the FAD of the enzyme, followed by electron equilibration between the FAD and Fe/S centers. In order to obtain rates for equilibration between the molybdenum center arid the other sites of the enzyme, we have used the radiolytically generated radical of N-methylnicotinamide (NMN•; E°'ie- = -1.01 V vs. SHE) to introduce reducing equivalents at the molybdenum center.

Results The reaction of desulfo XO with radiolytically generated NMN- at pH 6.0 is shown in Figure 1. As with the alloxanthine-blocked enzyme (data not shown), the transient consists of a very rapid absorption decrease on the timescale of 100 H-s, and a slower absorption increase on the ms timescale. On the basis of the observed rate constant of 125 s~l, its wavelength dependence and

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin-New York-Printed in Germany

682

lack of dependence on enzyme concentration, the slow phase of the reaction is attributed to the transfer of an electron equivalent from an Fe/S center to the FAD. The fast phase of the reaction of desulfo XO with NMN• is both faster and more complex than is observed with the alloxanthine-blocked enzyme (where the molybdenum center is rendered redox—inert; ref. 4). It is best described by two exponentials having ki = 34,000 s - l and k2 = 9000 s~l, with the faster component accounting for the all of the decrease in absorbance due to consumption of NMN-. We propose that NMN- reacts at the molybdenum center of the enzyme (first component of the fast phase, which is dependent on enzyme concentration), followed by internal electron transfer from the molybdenum center first to one (or both) of the Fe/S centers (second component of the fast phase) and finally to the FAD (slow phase). Upon increasing the pH to 8.5, the slow phase of the reaction is lost, consistent with the known pH dependence of the distribution of reducing equivalents within XO (5), where at high pH Fe/S reduction predominates over that of the FAD. Experiments have also been performed using radiolytically generated 5-deazalumiflavin (dFl-) to reduce desulfo XO. The transient is again biphasic, with k3i0w = 145 s - 1 (Figure 2) . As with the NMN- reaction, the wavelength dependence of the slow phase is consistent with equilibration between Fe/S and FAD, resulting in substantial formation of FADH-. The behavior of the fast phase is consistent with the direct reduction of both Fe/S and FAD by dFl-. At pH 8.5, the slow phase of the transient exhibits a wavelength dependence consistent with equilibration between Fe/S and FAD, but in this case the result is Fe/S reduction (consistent with the known pH dependence of the distribution of reducing equivalents within XO). The observed k s i ow at pH 8.5 is 220 s -1 . In none of the experiments performed with radiolytically generated dFl- were slow spectral changes observed with k < 20 s -1 , as have been seen in flash photolysis experiments (and which have been ascribed to an unspecified artifact).

683

Figure 1. Reaction of NMNwith desulfo XO at pH 6 (525 nm).

Figure 2. Reaction of dFlwith desulfo XO at pH 6 (525 nm).

The present experimental results.are summarized in Scheme I. Fe/S I is most likely the Fe/S center equilibrating rapidly (kobs=9000 s"1) the molybdenum, given the strong magnetic interaction observed by EPR between these two centers (7). The slow phase of the reaction of both NMN- and dFl- must be the direct equilibration between the Fe/S centers and FAD. Since kobs does not change significantly with the level of enzyme reduction prior to pulse (data not shown), we conclude that the Fe/S centers equilibrate at comparable rates with the FAD. 9,000

Mo

*

s" 1

Fe/S I I | Fe/S II

1 2 5 s -!

FAD

Scheme I

684

The pH dependence of the slow process observed in the reaction of dFl- with desulfo enzyme (k0bS=145 s - 1 at pH 6 and 220 s - 1 at pH 8.5) is in reasonable agreement with previous observations using the pH jump technique monitoring equilibration between the FAD and Fe/S I (8) and radiolytic studies using viologen radicals as enzyme reductant to monitor equilibration between FAD and Fe/S II (9).

The work presented here using

radiolytically generated NMN• to rapidly reduce XO at the molybdenum center clearly demonstrates that transient reduction of the Fe/S centers is observed in the transfer of reducing equivalents between the molybdenum and FAD of XO.

Both NMN- and

dFl• are found to react cleanly with XO when generated by the methods used in the present study.

References 1. Hille, R., V. Massey. 1985. In: Molybdenum Enzymes (T.G. Spiro, ed.), John Wiley & Sons, New York, pp. 4 43. 2. Hille, R., V. Massey. 1981. J.Biol.Chem. ZML, 9090. 3. Porras, 9096. A.G., J.S. Olson,, G. Palmer. 1981. J.Biol.Chem. 4. Massey, V., H. Komai, G. Palmer, G.B. Elion. 1970. J.Biol.Chem. 241, 2837. 5. Porras A.G., G. Palmer. 1982. J.Biol.Chem. 257, 11617. 6. Komai, H., H., V. Massey. 1971. In:Park Flavins andBaltimore, Flavoproteins (Kamin, ed.) University Press, 399. 7. Lowe, D.J., R.M. Lynden-Bell, R.C. Bray. 1972. Biochem.J. 130 f 239. 8. Hille, R., V. Massey. 1986. J.Biol.Chem. 2£1, 1241. 9. Anderson, R.F., R. Hille, V. Massey. 1986. J.Biol.Chem. 261. 15870.

EVIDENCE FOR PARTICIPATION OF THE PHOSPHOSERYL RESIDUE OF XANTHINE OXIDASE IN THE HYDROXYLATION EVENT DURING ENZYME CATALYSIS

Susan C. D'Ardenne and Dale E. Edmondson Department of Biochemistry, Emory University, Atlanta, GA 30322 USA

Int roduct ion

The hydroxylation mechanism for the conversion of xanthine to uric acid by bovine milk xanthine oxidase (XO) and the role of the Mo center in this process has been the subject of a number of studies. (1) have shown that

18

Mass spectral data

0 from solvent is transferred to a group on the

enzyme before its subsequent transfer to a substrate intermediate to form uric acid.

Although the chemical nature of this group has not been

determined, possible clues to its identity arise from the following data. ESR experiments by Bray's group have demonstrated a strong Mo (V) on incubation of reduced XO in as a possibility.

17

17

0 coupling to

0-H20 (2) suggesting a Mo=0 group

A previous communication from our laboratory (3) has

shown a reduction in peak intensity of the phosphoseryl on incubation of the enzyme with substrate in

17

0-H20.

31

P NMR resonance

This observed

reduction in intensity of the phosphoseryl resonance (relative to an

16

0

control) could be interpreted as a result of line-broadening of the phosphorus resonance due to quadrupolar relaxation by a bound

17

0 nucleus.

This result implicates a possible catalytic function for the phosphoseryl residue.

To investigate further the role of the phosphoseryl residue in

the catalytic mechanism of XO, we report here experiments that demonstrate 180 incorporation from solvent into the phosphoseryl phosphate moiety on incubation of functional enzyme with substrate in

18

0-H20.

8

Control experi-

ments with non-functional desulfo XO show no 1 0 incorporation.

These data

provide support for a functional role of the phosphoseryl residue in the catalytic mechanism of XO.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

686 Methods Bovine milk xanthine oxidase (56% functional) was dissolved in 97% l80-H20 0.1 M NaPPj, pH 8.5, and incubated with a 5-fold molar excess of xanthine/catalytic site for approximately 5-7 minutes to allow several catalytic turnovers to occur.

The enzyme was then precipitated with

trichloroacetic acid to remove all cofactors (FAD, MoCo and Fe2/S2> and buffer salt.

The protein precipitate was suspended in 10 mM (NH4>2C03 and

dialyzed against two 3-liter volumes of the same buffer.

The enzyme was

then subjected to alkaline conditions in which the covalent phosphate was released from the phosphoseryl residue by P - e l i m i n a t i o n .

The liberated

phosphate was purified by Dowex cation exchange chromatography, dried in vacuo, and dissolved in D2O containing 0.1 mM EDTA, pH 7.1 for analysis by 31

P NMR.

Desulfo enzyme was prepared as described (4) and then subjected

to identical turnover conditions in form.

18

0-H 2 0 as described for the functional

Subsequent conditions for liberation and purification of the

covalent phosphate moiety for NMR analysis were as described above.

Results The use of

31

P NMR approaches to probe for the incorporation of

18

0 into

the phosphate moiety relies on the observation of an isotope shift of 0.02 ppm per

18

0 atom.

Each shift is additive so that P18C>4 exhibits a chemical

shift 0.08 ppm upfield from P16Oi) (5) .

Small shifts such as this cannot be

monitored with enzyme-bound phosphates and therefore necessitates the removal of the phosphate moiety from the enzyme. field

31

Figure 1A shows a high-

P NMR spectrum of the inorganic phosphate released from 56% func-

tional XO after treatment as described above.

Four resonances are observed

which are separated by 0.02 ppm as is expected for

18

most prominent peak observed corresponds to a single the phosphate moiety.

0 isotope shifts. 18

The

0 incorporated into

Smaller peaks are also observed and are assigned to

P160,j, P160218C>2, and P16018C>3.

Demonstration of the correct assignment of

these resonances is shown in Figure IB in which 0.5 mM P1604 (final concentration) was added to the sample, resulting in an increase in intensity of flnly the low-field resonance.

From integrated intensity comparisons of

687 P1603180

chemical shift

0.02 ppm/div

Fig. 1A: 31 P NMR spectrum of phosphate purified from 56% functional XO after several catalytic turnovers in 18 0-H 2 0. Chemical shifts are observed at -0.779 ppm, -0.801 ppm, -0.823 ppm, and -0.846 ppm. All phosphate samples were dissolved in D2O containing 0.1 mM EDTA, pH 7.1. All spectra were recorded at 202.4 MHz; 45° flip angle; 5K Hz spectral width; 32K set of data points. Trimethyl phosphate was used as an external chemical shift standard. B: Spectrum after the addition of 0.5 mM 1 6 0 phosphate (final concentration) to the sample in A. The increase in intensity at -0.77 9 ppm corresponds to P16C>4. Each spectra is the result of 800 acquisitions.

0.01 ppm/div

0.02 ppm/div chemical shift

Fig. 2A and B: 3lp nmr spectrum of the phosphate purified from nonfunctional desulfo XO after treatment as described in Methods section. Chemical shifts are observed at 1.802 ppm and -8.850 ppm. A: Expanded representation of the resonance at 1.802 ppm. B: Expansion of the resonance observed at -8.850 ppm. The peak at 1.802 ppm corresponds to the chemical shift of a P16C>4 standard under the same conditions. The absolute chemical shift difference of the Pi resonance relative to Fig. 1 is due to slight differences in pH. This spectrum is a result of 5000 acquisitions under the same instrumental settings as described in Fig. 1.

688 Fig. 1A and IB, we estimate the yield of phosphate based on total XO in the experiment to be > 95%.

Figures 2A and 2B show the spectrum of the cova-

lent phosphate isolated from non-functional desulfo XO after treatment identical to that described for the functional enzyme.

Rather than a

simple phosphate resonance attributable to inorganic phosphate, two resonances are observed.

One resonance at 1.8 ppm

40% of the total

integrated intensity) is due to inorganic phosphate (Figure 2A).

The

upfield resonance at -8.8 ppm, which constitutes - 60% of the total integrated phosphate intensity,

is an unknown chemical entity (Figure 2B)

and is not observed in the sample prepared from functional XO. isotope shifts are observed for either resonance.

No

18

0

These data demonstrate

solvent oxygen incorporation into the covalent phosphoseryl residue of XO to be dependent on its state of functionality.

Discussion Our laboratory first published a detailed NMR study of the non-covalent phosphorus groups (the FAD and the Mo cofactor), and the single covalentlybound phosphorus residue per active center Mo, which was identified as a phosphoseryl residue (6).

Several

31

P NMR experiments suggest the phospho-

seryl residue to be in close proximity to the Mo center.

A recent ENDOR

study (7) has suggested a phosphorus coupling to the Mo (V) center from functional enzyme prepared by formaldehyde treatment.

It remains for

future work to prove this assignment and to determine whether it arises from the phosphoseryl residue or from the phosphorylated MoCo.

The results

of this study provides strong preliminary data for the involvement of the phosphoseryl residue in XO catalysis.

Solvent exchange with phosphate

oxygen requires prolonged incubation at elevated temperatures (8).

There-

fore, solvent exchange into a phosphoseryl phosphate is neither expected nor observed without its catalytic involvement. reported experiments merit further study.

Several aspects of these

Although chemical analysis shows

the expected yield of phosphate after P - e l i m i n a t i o n from functional XO as well as desulfo XO, in general, the

31

P NMR signals were stronger with the

sample prepared from the functional enzyme.

Secondly, the finding of an

additional resonance at ~9 ppm upfield from the inorganic phosphate resonance for the desulfo enzyme suggests it to be a chemical entity differing from Pi.

Whether the 60:40 ratio of the two phosphate peaks in Figs. 2A

689 and 2B represents a distinction between naturally-occurring desulfo and desulfo formed on CN~ treatment also awaits further study.

While the

answers to the above questions and the detailed role of the phosphoseryl residue in catalytic turnover in XO warrants further investigation, we feel that a recent model study on Ir (III) and Co (III) phosphate complexes by Sargeson's group (9) merits consideration as a possible model for portions of the XO catalyzed reaction.

Mechanistic experiments to

test further this model are underway in this laboratory.

Acknowledgement This work was supported by grants from the NSF (DMB-8616952) and from the NIH (GM-29433).

References 1. Hille, R. and H. Sprecher. 1987. J. Biol. Chem. 2£2, 10914-10917. 2. Gutteridge, S. and R.C. Bray. 1980a. Biochem. J. 1 M , 615-623. 3. Edmondson, D.E., M.D.Davis, F. Muller. 1984. Flavins and Flavoproteins (R.C.Bray, P.C. Engel, and S.G. Mayhew, eds.). Walter de Gruyter & Co., Berlin, pp. 309-317. 4. Massey, V. and D.E. Edmondson. 1970. J. Biol. Chem. 245. 65956598 . 5. Cohn, M. and A. Hu. 1978 . Proc. Natl. Acad. Sci. USA. 23., 203.

200-

6. Davis, M.D., D.E. Edmondson and F. Muller. 1984. Eur. J. Biochem. 145, 237-243. 7. Pinhal, N.M., R.C. Bray, N.A. Turner and D.J. Lowe. 1989. Highlights in Modern Biochem., Vol.1 (A. Kotyk et al., eds) pp. 273-280. 8. Cohn, M. 1957. Methods in Enzymology, Vol. 4 (S. Colowick and N.O. eds.) Academic Press, N.Y. pp. 905-914. 9. Hendry, P. and A.M Sargeson. 1989. J. Am. Chem. Soc. Ill, 2521-2527

ENDOR STUDIES OF THE MOLYBDENUM CENTRE OF XANTHINE OXIDASE

B.D. Howes, B. Bennett, D.J. Lowe, and R.C. Bray University of Sussex, Brighton, BN1 9QJ, U.K.

The technique of ENDOR (electron nuclear double resonance)

spectroscopy

makes possible the evaluation of hyperfine couplings for nuclei interacting too weakly with a paramagnetic centre to permit their study by e.p.r.

Under favourable circumstances it is possible to evaluate the

dipolar coupling component of the hyperfine coupling from ENDOR spectra, and so to obtain information on interatomic distances.

ENDOR

investigations are long established in the flavin field.

The technique

is particularly informative when it can be used in conjunction with samples labelled by substitution with stable isotopes introduced into enzymes at specific sites.

Most relevant earlier work relates to liquid

solution studies of flavin radicals.

This communication

summarizes

recent ENDOR work at liquid helium temperatures on the molybdenum centre of xanthine oxidase.

We have used 'H,

31

P and

13

C ENDOR to probe the

local structure (1) around the metal in this enzyme.

A number of different configurations of the molybdenum centre of xanthine oxidase in the Mo(V) state are known.

They are distinguished

by characteristic e.p.r. spectra, that have been given the names Very Rapid, Rapid, Slow, Inhibited and Desulpho Inhibited.

Considerable

structural information on each of these is already available, from e.p.r. (1) and less directly from EXAFS, eg (2). (3) *H ENDOR spectra for four of these species.

number of distinct groups.

We have recorded

Nine or more protons or

sets of equivalent protons are observed, with hyperfine constants ranging from 0.08 to 40 MHz.

directly

coupling

The protons fall (3) into a

Those showing couplings in the range of

0.08-4.0 MHz are present for all the reduced forms of the enzyme and

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

692 all, or almost all, of them do not exchange with water of the solvent. They are presumed to be due to protons, either from amino acid residues or present in the pterin molybdenum cofactor.

Contrary to claims from

other laboratories, at present it is not possible to deduce further information on the nature of these protons.

This position could change

however, if it became possible to remove and replace the cofactor reversibly or if site-directed mutagenesis could be employed to change amino acid residues in the active centre region.

The more strongly

coupled protons detectable in the ENDOR spectra include those studied by e.p.r. in earlier work.

Those most strongly coupled, which are presumed

to be from -OH or -SH ligands of molybdenum, exchange with the solvent. Others which do not exchange include those derived from substrate molecules bound at the active centre.

Specific findings (3) are that

the aldehyde residue of the Inhibited species has been oxidized and that the four protons derived from the ethylene glycol residue of the Desulpho Inhibited species are not all equivalent.

31

P ENDOR has recently contributed significantly (4) to information on

the structure of the xanthine oxidase molecule.

This contains

phosphorus only (4,5) as the phosphate group of the pterin cofactor and the pyrophosphate of the FAD.

A single ENDOR resonance from

observed (4) for all reduced xanthine oxidase species.

31

P is

This remains

unchanged, when FAD is removed from the enzyme, indicating that the phosphorus atoms from this are more than 10 A from molybdenum. Evaluation (4) of the dipolar coupling gives a value of 7-10A for the distance between the metal and the phosphorus of the pterin cofactor. This makes it clear that the side chain of the cofactor is in an extended conformation in the enzyme molecule.

We have also made a start on examination of

13

C ENDOR spectra from

xanthine oxidase species that bear bound substrate residues.

These

include the stable Desulpho Inhibited species (1) prepared from reaction of the desulpho enzyme with

13

C-ethylene glycol and the Very Rapid

693

a

b

7

10

ENDOR spectra for the Very Rapid signal obtained with: (a) xanthine labelled in the 3 8-position with 1»3 C and (b) unlabelled xanthine. The reaction time was 11 ms. at pH 10.2 and 21"C. The recording temperature was 15K, microwave frequency 9.49 GHz, microwave power 2mW, scanning rate 0.52 MHz/s and modulation depth 200 kHz. The spectrum was recorded at a g value of 1.966, ie near to the g 2 feature of the e.p.r. spectrum.

species, which is a transient intermediate in enzyme turnover.

In the

former, the two carbon atoms are not equivalent, confirming the conclusion

(3) noted above of the inequivalence of the glycol

protons.

Fig 1(b) shows the ENDOR spectrum for the Very Rapid species, when unlabelled xanthine was employed to generate the signal. xanthine labelled in the 8-position with

13

In Fig 1 (a),

C was used and the features

between 7.0 and 9.5 MHz are clearly due to coupling to Mo(V) of this of the bound xanthine residue.

across the e.p.r. spectrum, indicate the M o Such a value excludes the possibility

is short for a

Mo-0-C system

I3

points

C distance to be about 2.6

(7) of a direct Mo-C bond, but

(8), unless a 4-membered ring is

involved.

We are therefore currently seeking to establish by '*N, and

perhaps by

1S

N , ENDOR whether the xanthine N-9 is also coordinated to

molybdenum, so completing such a ring in the Very Rapid species. results have implications

17

These

(cf.2) regarding the mechanism of xanthine

oxidase action and tend further to confirm the earlier mechanism.

C

Preliminary estimates of the dipolar

coupling, obtained by recording the ENDOR spectrum at different

A.

13

(cf.l)

However, it will be important in further work to confirm, by

0 ENDOR, the conclusion

(cf.7) that the Very Rapid species bears an

oxo ligand in addition to the oxygen ligand discussed

here.

694 Acknowledgement

We thank Dr D.L Hughes for helpful discussions on the structures of molybdenum complexes.

The work was supported through a Linked Research

Group from the AFRC and by the Wellcome Trust.

E.p.r. equipment was

provided by the SERC.

References

1.

Bray, R.C. 1988. Quart. Rev. Biophys. 21, 299-329.

2.

Turner, N.A., R.C. Bray and G.P. Diakun. 1989. Biochem J. 260, 563-571.

3.

Howes, B.D., N.M. Pinhal, N.A. Turner, R.C. Bray, G. Anger, A. Ehrenberg, J.B. Raynor and D.J. Lowe. 1990. Biochemistry 29, 6120-6127

4.

Howes, B.D., B, Bennett, A. Koppenhofer, D.J. Lowe and R.C. Bray. 1990. Biochemistry (submitted)

5.

Johnson, J.L., R.E. London and K.V. Rajagopalan. 1989. Proc. Nat. Acad. Sci. USA. 86, 6493-6497.

6.

Tanner, S.J., R.C. Bray and F. Bergmann. 1978 Biochem. Soc. Trans. 6, 1328-1330.

7.

George, G.N. and R.C. Bray. 1988. Biochemistry. 27, 3603-3609

8.

Gutteridge, S. and R.C. Bray. 1980. Biochem. J. 189, 615-623.

MILK XANTHINE DEHYDROGENASE

Jennifer Hunt and Vincent Massey Department of Biological Chemistry, University of Michigan Ann Arbor, Michigan

Introduction Milk xanthine oxidase

(MXO), a dimer containing one FAD, two

2Fe/2S clusters and one molybdenum per subunit, catalyzes the oxidation of xanthine to urate. Previous studies (1,2,3) have compared various properties dehydrogenase

of this enzyme

with

xanthine

(XDH) from chicken liver, which also converts

xanthine to urate but requires NAD rather than 02 as oxidant. It has been NAD-dependent

reported

(4) that MXO can be converted

dehydrogenase

form

by

reducing

to an

agents;

by

including dithiothreitol in all steps of the MXO preparation, we have been able to purify the milk enzyme as nearly

90%

dehydrogenase form. One of the most striking differences between MXO and chicken liver XDH is the stabilization of neutral 8-SH-FAD by XDH; free 8-SH-FAD has a pK of 3.8, yet at pH 7.5 the 8-SH-FAD is bound

to

the

chicken

liver

XDH

in

the

neutral

form.

In

contrast, MXO binds 8-SH-FAD in the anionic form at pH 8.5. In the experiments described below, the pK 1 s of

8-SH-FAD

when bound to MXO and to MXDH respectively were determined. Another characteristic of MXDH is its ability to stabilize the blue neutral semiquinone; photoreduction of MXDH FADH°

shows

present throughout most of the reduction, whereas MXO

shows very little semiquinone.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter&Co., Berlin • New York-Printed in Germany

696 Results and Discussion Deflavo MXO was reconstituted with 8-SH-FAD at pH 8.5; at this

pH,

the

enzyme-bound

8-SH-FAD

is

in

the

anionic,

benzoquinoid form C (scheme 1), as evidenced by the spectral shape. As the pH was progressively citric

acid,

spectral

changes

lowered with

showed

protonated neutral form of the flavin

powdered

conversion

to

the

(form A) . From this

titration a pK of 5.1 was determined for MXO-bound 8-SH-FAD, only a small perturbation

from the pK of the free

flavin

(3.8).(Figure 1)

Scheme 1

fcl x I iX ma-560-600nm

[U^.SZOnm

Deflavo MXDH reconstituted with 8-SH-FAD was incubated with 5mM

DTT

to

obtain

the

maximum

XDH

form.

The

initial

spectrum at pH 6.5 closely resembled that of chicken liver XDH-bound spectrum

8-SH-FAD.(Figure shifted

to

that

2). of

With the

pH

adjustment

anionic

form

the

(after

titration, addition of citric acid caused a return to the original spectral shape, showing that conversion of MXDH to MXO had not occured) .

A pK of 9.0 was found for 8-SH-FAD

bound to MXDH, indicating a significant difference between the flavin binding sites of MXO and MXDH. Another evidence of different flavin environments in MXO and MXDH is the stabilization of the blue neutral semiquinone of native FAD by MXDH. During anaerobic reduction of MXO little or no

semiquinone

is observed,

but

in the

case of

semiquinone appears in early stages of reduction and is

MXDH,

697 0.5

350

450

550

650

Wavelength (nm)

750

Fig. 1: Spectra of 8-SH-FAD milk xanthine oxidase at different pH values, starting in 0.1M NaPPi, 0.3mM EDTA, pH 8.52 at 4°C and adjusting pH with powdered citric acid. Spectrum 1, pH 8.52; 2, pH 5.80; 3, pH 5.30; 4, pH 4.82; 5, pH 4.09. Inset: Plot of absorbance at 570nm vs pH. 0.4

(1) a c o _a Lo CO _a
300

at 6°C). The rate constant of flavinsemiquinone

oxidation

803 does not

depend on ferricyanide concentration

but decreases

(from 20 to 7 s

-1

at

(in the range 15-75 |1M)

6°C) when pyruvate

concentration

increases (from 5 to 50 mM, Fig 2).

Fig. 2 : Time course of the flavosemiquinone rapid

oxidation mixing

of

at

439 nm,

partially

after

reduced

flavocytochrome b2 with ferricyanide,at 5,

10

and

50

mM

pyruvate

(100

mM

phosphate buffer, pH 7.0; t= 6°C) .

The simulation indicates that this dependence is expected when the rate limiting step is pyruvate dissociation

(hyp. 2) and not the electron

transfer from Pyruvate-Fg to H Q or to ferricyanide (hyp. 1). In a parallel rapid mixing study at 10 mM pyruvate in the presence of Hansenula

anomala

cytochrome c, Janot et al

(6) have found that the

limiting step of semiquinone oxidation by heme

is equal to 5 s - ^ at 5

°C. This rate constant is in good agreement with the present results with ferricyanide.

Conclusion Pyruvate binding to the semiquinone flavin decreases at least ten times the rates

of electron

transfers

from the

'Pyruvate-Flavosemiquinone'

complex both to the heme partner in the same subunit and to the external acceptor ferricyanide. In fact, at the present time we cannot exactly know how slow is the electron

transfer

of

the

pyruvate

liganded

flavinsemiquinone

intermediate. Indeed, the results and simulation of the rapid kinetic experiments

at variable concentration of pyruvate

indicate that the

process observed could be pyruvate dissociation regenerating the fully reactive non liganded F g intermediate. The actual electron transfer from the liganded form should thus be very slow and completely shunted through pyruvate dissociation, that would become the rate limiting step.

804 The present

theories on the mechanism of electron transfer

within

donor/acceptor complexes indicate that four factors could play a role in the

flavocytochrome

b2

system.

Their

relative

importance

in

the

modification afforded by pyruvate binding has been evaluated taking into account a hypothetical model of interaction between ferricyanide and the known (7) structure of S.cerevisiae flavocytochrome b 2 : 1 - Minimal distance between acceptor and donor molecules : the distance between F 3

and the heme b 2

as well as the distance between F g

and

ferricyanide do not appear modified in the structure by the presence of Indeed ferricyanide is too large to fit into the pyruvate

pyruvate.

binding crevice near the isoalloxazine plane and must

stand at the

periphery of the flavodehydrogenase assembly in the presence as in the absence of bound pyruvate. 2 - Probability of residence of the acceptor at the position of minimal distance

cannot be evaluated. It is known that the cytochrome moiety

fluctuates relatively to the flavodehydrogenase moiety. 3 - The driving force AF.1U (acceptor-donor) is modified in our system by 140 mV by pyruvate binding. From published data (8) this alteration could provide a factor approx.7 in the decrease of rate constant. This factor alone could then account for our data if the rate measured

5 s - ^ were

actually the rate of electron transfer. 4 - The intrinsic reactivity of the flavosemiquinone could be modified as a result of the different distribution of the electron density on the isoalloxazine ring induced by the presence of pyruvate lying close to the ring.

This factor cannot be estimated at the light of the present data.

References 1. 2. 3. 4. 5.

Gervais, M., Y. Risler, S. Corazzin. 1983. Eur.J.Biochem. ISA, 253. Capeill6re-Blandin,C. 1982. Eur .J.Biochem. 123., 533. Tegoni, M., J.M. Janot, F. Labeyrie. 1990. Eur .J.Biochem. 12fl., 329. Tegoni, M., J.M. Janot, F. Labeyrie. 1986 Eur.J.Biochem. 155, 491. Tegoni, M., M.C. Silvestrini, F. Labeyrie, M. Brunori. 1984. Eur. J. Biochem. 1411, 39. 6. Janot, J.M., C. Capeillfere-Blandin, F. Labeyrie. 1990. Biochem. Biophys. Acta 1016r 165. 7. Xia,Z., N. Shamala, P.H. Bethge, L.W. Lim, H.D. Bellamy, N. Xuong, F. Lederer, F.S. Mathews. 1987. Proc.Natl.Acad.Sci. U.S.A. M , 2629. 8. Tollin, G., T.E. Meyer, M.A. Cusanovich. 1986. Biochim. Biophys.Acta M l , 29.

CRYSTAL STRUCTURE OF P-CRESOL METHYLHYDROXYLASE AT 3.0 A RESOLUTION

F. S. Mathews and Z.-w. Chen Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO 63110,. USA W. S. Mclntire Molecular Biology Division, Veterans Administration Medical Center, San Francisco, CA 94121, USA, and Departments of Biochemistry-Biophysics and Pharmaceutical Chemistry, University of California, San Francisco, CA 94143

Introduction p-Cresol methylhydroxylase (PCMH) is a flavocytochrome c found in certain pseudomonads (1). It catalyzes the hydroxylation of p-cresol first forming p-hydroxybenzyl alcohol and then phydroxybenzaldehyde. It is a tetramer of Mr 115,600 containing 2 flavoprotein subunits each of Mr 49,000 and 2 cytochrome subunits each of Mr 8,800. The flavoprotein subunit contains FAD covalently bound to a tyrosine side chain (2). The amino acid sequence of the cytochrome subunit is known (3) but that of the flavoprotein subunit is unknown, except for the Nterminal 56 residues and the flavin binding heptapeptide. The structure of PCMH was solved initially at 6.0 A resolution (5) and extended to 3.0 A resolution (6) using data recorded by area detector from crystals of the native protein and 2 heavy atom derivatives. The x-ray phases were filtered by a solvent leveling procedure (7) and the final map was averaged about the molecular 2-fold axis (8). The electron density map has now been improved by including a third derivative and remeasuring one of the earlier derivatives, with care being taken to record full anomalous scattering

data. The final

figure of merit for the protein phases improved from 0.55 to

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

806 0.62. The phases were further refined by cyclic averaging (8). The earlier model (6) was generally verified and some marked improvements in the stereochemistry, particularly in the cytochrome domain, were made.

Results The structures of the flavoprotein and cytochrome subunits are shown in figure 1. The flavoprotein subunit contains about 463 residues and consists of 3 domains. The first domain contains about 215 residues and binds the adenosine and ribose moiety of the flavin in a grove formed by extended chains. The second domain, of about 210 residues, contains the covalent flavin attachment site and forms a dome covering the flavin ring. The re face of the flavin ring is packed against the chain while the

polypeptide

face is open to a pocket in the dome. The

topology of folding of the second domain is unusual with an 8stranded antiparallel 0 sheet flanked by 4 o-helices on one side and by a single helix on the other. The third domain is

Fig. 1 Stereograph showing the flavoprotein and cytochrome subunits of p-cresol methylhydroxylase. The polypeptide chain tracing shows the C a positions. The N- and C-termini are indicated. The heme and FAD are also shown.

807 a b o u t 38 r e s i d u e s long a n d lies a g a i n s t d o m a i n 1, a p p r o x i m a t e l y c o v e r i n g the a d e n y l a t e p o r t i o n of the FAD The s e q u e n c e of the c y t o c h r o m e s u b u n i t h a s b e e n f i t t e d the e l e c t r o n d e n s i t y . T h e r e are 3 h e l i c a l 8-14,

segments,

group. into

residues

36-45 a n d 6 2 - 7 2 . The heme g r o u p is b o u n d c o v a l e n t l y

thioether

by

l i n k a g e s to Cys 15 a n d Cys 18 a n d by c o o r d i n a t i o n

the i r o n a t o m t h r o u g h His 19 a n d M e t 50. The heme

g r o u p s are b o t h q u i t e e x p o s e d to s o l v e n t . The s t r u c t u r e

is

generally

such

similar

as c y t o c h r o m e

c5

to other small b a c t e r i a l

cytochromes,

(9), b u t c o n t a i n s a 9 - r e s i d u e d e l e t i o n

o n l y a single h e l i x b e t w e e n the h i s t i d i n e a n d l i g a n d s c o m p a r e d to c y t o c h r o m e

to

propionate

and

methionine

c5.

The c y t o c h r o m e d o m a i n is o r i e n t e d w i t h the e x p o s e d h e m e c o n t a i n i n g the 2 t h i o e t h e r l i n k a g e s to the heme v i n y l in c o n t a c t w i t h the f l a v o p r o t e i n s u b u n i t . The heme a n d g r o u p s are a b o u t 8 A a p a r t at the c l o s e s t p o i n t s a n d p l a n e s are i n c l i n e d to e a c h other by a p p r o x i m a t e l y

edge,

groups, flavin

their

65°.

Fig. 2 S t e r e o g r a p h of FAD a n d the C a b a c k b o n e for r e s i d u e s 215 to 395 of the f l a v o p r o t e i n s u b u n i t , w h i c h forms m o s t of d o m a i n 2. The d i f f e r e n c e d e n s i t y close to the f l a v i n ring f r o m a c r y s t a l s o a k e d in p - c r e s o l , w i t h a m o d e l for p - c r e s o l s u p e r i m p o s e d , is a l s o shown. The n u m b e r i n g for the C a b a c k b o n e of the f l a v o p r o t e i n s u b u n i t b e g i n s at 101.

808 A crystal of PCMH has been soaked in a solution containing the substrate p-cresol. The difference electron density indicates that the substrate moiety is located in the pocket formed by domain 2 opposite the ^

face of the flavin ring (figure 2).

The p-cresol molecule is centered over the N-5 atom of the flavin ring and lies roughly parallel to it.

Acknowledgement This research was supported by NSF grants No. DMB-8718741 and DMB-8816618, USPHS grant no. HL-16251 and the Veterans Administration.

References 1

Keat, M. H. and Hopper, D. J. (1978) Biochem. J. 175, 649-658.

2

Mclntire, W. S., Edmundson, D. E., Hopper, D. J. and Singer, T. P. (1981) Biochemistry 20, 3068-3075.

3

Mclntire, W., Singer, T. P., Smith, A. J. and Mathews, F. S. (1986) Biochemistry 25, 5975-5981.

4

Mclntire, W., Hopper, D. J., and Singer, T. P. (1985) Biochem. J. 228, 325-335.

5

Shamala, N., Lim, L. W., Mathews, F. S., Mclntire, W., Singer, T. P. and Hopper, D. J. (1986) Proc. Natl. Acad. Sci. U. S. A., 83, 4626-4630.

6

Bellamy, H.D., F.S. Mathews, W.S. Mclntire and T.P. Singer, (1987) In: Flavins and Flavoproteins (D.B. McCormick and D.E. Edmondson, eds) Walter de Gruyter, Berlin, 673-676.

7

Wang, B. C. (1982) Methods Enzymol. 115, 90-112.

8

Bricogne, G. (1976) Acta Crystallographica A32, 832-847.

9

Ghosh, D., O'Donnell, S., Furey,S., Jr., Robbins, A. H. and Stout, C. D. (1982) J. Mol. Biol. 158, 73-109.

INTERACTIONS IN MULTI-SUBUNIT REDOX PROTEINS: NMR STUDY OF THE CYTOCHROME SUBUNIT OF p-CRESOLMETHYLHYDROXYLASE W. S. Mclntire Veterans Administration Medical Center, San Francisco, California 94121; Dept. Biochemistry and Biophysics, and Dept. Anaesthesia, University of California, San Francisco, CA 94121 F. S. Mathews Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110 S. Bagby, J. A. Charman, H. A. O. Hill Inorganic Chemistry Laboratory, University of Oxford, Oxford, England P. C. Driscoll Department of Biochemistry, University of Oxford, Oxford, England G. L. McLendon Department of Chemistry, University of Rochester, Rochester, New York 14627

Introduction p-Cresolmethylhydroxylase, (PCMH, E.C. 1.17.99.1), an a 2 (3 2 flavocytochrome, contains (1) flavin catalytic subunits ( a ) (M r 48,600) and c-type cytochrome subunits (p) (M r 8780). Although these subunits are strongly bound (K d ~ 10 M), they can be separated (2) by isoelectric focusing to give independently stable proteins. The crystal structure of the intact enzyme has (3) recently been determined. Knowledge of the structures of the isolated subunits could provide unique information on molecular recognition and interactions in multi-subunit redox proteins. To this end, we

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

810

have undertaken the determination of the structure in solution of the cytochrome c subunit of PCMH, using N M R methods. Herein we present assignments of some of the heme proton resonances and also of some residues which are situated close to the heme. These assignments indicate that there is a significant structural difference between the active sites of the isolated subunit in solution and of the bound subunit in the crystalline state.

Results The general approach used in these studies has been outlined (4) by Williams et al. In the 2 D N O E spectrum of the cytochrome subunit, the pattern of heme meso proton resonances (Figure 1) can be clearly understood: the y proton should give four single proton N O E s , the 8 proton should give two three-proton (methyl) NOEs, the a proton should give one strong three proton (methyl) and one weak three-proton (methyl) NOE, as should the P-meso proton. Hence, from the Figure, we can assign resonance

"0 O) T) 3

'

4

-•

'

00 CO 1.0

ppm

0.0

Figure 1. The contour plot of part of the N O E S Y spectrum of the reduced cytochrome c subunit of PCMH is shown. The pattern of N O E s for each of the four heme meso protons is indicated.

811

1 = Y and 2 = 8. Peaks 3 and 4 could be assigned to either the a or P protons. However, it was noted that resonance 4 has an N O E to the e C H 3 of M 5 0 , the heme axial ligand. Thus, we reasoned that inspection of the heme stereochemistry might distinguish the a and p protons. The X-ray structure (3) of the intact enzyme shows that only one meso proton, yH, is close enough (270 nm ~ 0) rules out energy transfer as a possible mechanism for initiating dimer cleavage.

However, model

studies show that thymine dimer radical anions are unstable and spontaneously monomerize (20), suggesting that dimer cleavage in the photolyase reaction might be initiated by electron transfer from 1FADH2* to generate FADH* plus thymine dimer radical anion.

FADH 2 would be regenerated after monomerization of the

dimer radical.

Acknowledgement This work was supported by NIH Grant GM 31704.

References 1.

Wang, B., S.P. Jordan and M.S. Jorns. 1988. Biochemistry 27, 4222-4226.

2.

Wang, B. and M.S. Jorns. 1989. Biochemistry 2g, 1148-1152.

3.

Johnson, J.L., S. Hamm-Alvarez, G. Payne, G.B. Sancar, K.V. Rajagopalan and A. Sancar. 1988. Proc. Natl. Acad. Sci. USA 85, 2046-2050.

4.

Jorns, M.S., B.Y. Wang, S.P. Jordan and L.P. Chanderkar. 1990. Biochemistry 22, 552-561.

5.

Jorns, M.S. 1987. In: Flavins and flavoproteins (Edmondson, D.E. and D.B. McCormick, eds.). Walter de Gruyter, Berlin, pp. 233-246.

6.

Sancar, G.B., M.S. Jorns, G. Payne, D.J. Fluke, C.S. Rupert and A. Sancar. 1987. J. Biol. Chem. 2£2, 492-498.

7.

Jorns, M.S., G.B. Sancar and A. Sancar. 1984. Biochemistry 23, 2673-2679.

8.

Payne, G., P.F. Heelis, B.R. Rohrs and A. Sancar. 1987. Biochemistry 26,

7121-7126. 9.

Jordan, S.P. and M.S. Jörns. 1988. Biochemistry 22, 8915-8923.

10. Jörns, M.S., B. Wang and S.P. Jordan. 1987. Biochemistry 26, 6810-6816. 11. Van Berkel, W J . H , W.A.M. Van den Berg and R. Müller. 1988. Eur. J. Biochem. 12S, 197-207. 12. Yasui, A., M. Takao, A. Oikawa, A. Kiener, C.T. Walsh and A.P.M. Eker. 1988. Nucleic Acids Res. lfi, 4447-4463. 13. Harbury, H.A., K.F. LaNoue, P.A. Loach and R. Amick. 1959. Proc. Natl. Acad. Sei. USA 45, 1708-1717. 14. Müller, F., M. Brustlein, P. Hemmerich, V. Massey and W.H. Walker. 1972. Eur. J. Biochem. 25, 573-580. 15. Kay, L.D., MJ. Osborn, Y. Hatefl and F.M. Huennekens. 1960. J. Biol. Chem. 235. 195-201. 16.

May, M., T J . Bardos, F.L. Barger, M. Lansford, J.M. Ravel, G.L. Sutherland and W. Shive. 1951. J. Am. Chem. Soc. 22, 3067-3075.

17. Jörns, M.S., E.T. Baldwin, G.B. Sancar and A. Sancar. 1987. J. Biol. Chem. 262. 486-491. 18. Turro, N J . 1978. Modern molecular photochemistry. The Benjamin/Cummings Publishing Co., Menlo Park. 19. Jörns, M.S. 1987. J. Am. Chem. Soc. 1Q9, 3133-3136. 20.

Lamola, A.A. 1972. Mol. Photochem. 4, 107-133.

CHEMICAL MODIFICATION HITH IODOACETAMI DE

Haruo

SUZUKI

OF CORYNEBACTERIUM

and Yasuko

SARCOSINE

OXIDASE

KAWAMURA-KONISHI

D e p a r t m e n t of B i o p h y s i c a l C h e m i s t r y , K i t a s a t o S c h o o l of M e d i c i n e , S a g a m i h a r a , K a n a g a w a 228,

University JAPAN

Introduction Sarcosine oxidative

oxidase

d e m e t h y 1 a t i o n of

CH3NHCH2C00H The B,

enzyme

Mr.44,000;

is a t t a c h e d

C,

"oxidase"

flavins, to

B.

enzyme

FAD

modifies

inhibitor

Therefore,

we p r o p o s e d

to

B (4.) •

FAD m o i e t i e s

and

Mr.110,000:

contains The covalent

that

the

is,

FAD

noncovalent and

electrons

covalent

by a s s u m i n g subunit

an

transfer

with that

the

roles

reported

on that

of

to

that

and

the

the

IAM

that

with sodium

the n o n c o v a l e n t

the

IAM

acetate,

a

sarcosine(4jj>). FAD

we d e t e r m i n e d

is a l s o

the

Cys residues,

IAM m o d i f i c a t i o n ,

of the m o d i f i e d

the d i f f e r e n t

observed

respect

IAM-reactive

oxygen

B specifically

B is p r e v e n t e d

In t h i s work,

on the

we

FAD

we

is d e p e n d e n t

Moreover,

of s u b u n i t

around

H202

"dehydrogenase"

Previously

inhibition

competitive

acetate

a

r e s p e c t i ve 1 y (2.) . T h a t

modification

subunit

the

oxygen:

mod i f i cat i on (.4.) .

sodium

as

m o i e t i e s (3.) .

of

sequences

Jorns proposed

function

+

subunits(A,

b o u n d FADs (X) •

a kinetic mechanism

i o d o a c e t a m i d e (I AM)

U-96 catalyzes

+ H2NCH2C00H

D, M r . 1 0 , 0 0 0 )

noncovalent

flavin

extent

HCHO

of 4 n o n i d e n t i c a l

FADs

sarcosine

We p r o p o s e d

•

Mr.21,000;

Sarcosine

of

+ H20

to s u b u n i t

sp.

sarcosine:

and covalently

covalent

from

+ 02

consists

noncovalently and

from C o r y n e b a c t e r i u m

Flavins and Flavoproteins 1 9 9 0 © 1991 by Walter de Gruyter&Co., Berlin - N e w Y o r k - Printed in Germany

acid

the e f f e c t

a n d the f u n c t i o n

enzymes.

bound

amino of

of the

828 Results

and

Discussion

The enzyme C

in 20

B

extracted

was fro»

separated t h e gel.

chymotrypsin

purified digests further

purified

earlier

fractions

of

in t h i s

UV

gas-phase

labeled mainly

containing

TC-2

N-terminal

Cis-Gly-Thr-Pro-Gly-Ala-Gly-Tyr

al. (5)

covalent Since

the

TC-2

fluorescent, peptide

Previously

enzyme 50

sodium

on t h e was

proteolytic

As

with

acetate

acid by

a of

sequences

an

Applied

were:

(TC-2)

acid as

contained

sequence

of

the

A s p - H i s ( F A D ) - V a l - A 1 a.

this

the covalent

digests

is

a the

that

of

Cys

sequence

and

FAD

is b o u n d

of

subunit

with

were

analyzed with

competitive Cys above,

was to

the

B

is

of

of

by H P L C . sodium the TC-1

sodium

peptides,

The

labeled

Labeling acetate. with

the

and a b s e n c e

and chymotrypsin,

inhibitor

residue IAM

above.

trypsin

inhibited

in t w o

in t h e p r e s e n c e

as d e s c r i b e d

at t h e s a r c o s i n e - b i n d i n g described

labeling

residues

[14C]IAM

digested

was g r e a t l y

s a r c o s i n e (A) > located

showed

presence

a c e t a t e (J.) • To s e e t h e e f f e c t

labeling

labeled

mM s o d i u m

acetate

enzyme

that

reported

we

with

was extensively peptide

peptide

we c o n c l u d e

later

(TC-1)

the amino

s i t e of

at

TC-2.

prevented acetate

reported

FAD-binding

the

and

eluting

that

amino

Ala-Gly-Ile-Ala-Cys-Xaa-Asp-Xaa-Val-Ala Shiga

tryptic

radioactive

peptide

sequences

were

separately

determined

The

and

trypsin

and

two

and

indicating

were

sequencer.

with

^C-peptide

as T C - 1

30

labeled

peptides

were collected

light,

peptide.

The

at

SDS/PAGE

the c h y m o t r y p t i c

HPLC column.

peptides

by

was d i g e s t e d

B showed

named

Fractions

purified

Biosystem

was

by

of

30 m i n

mM E D T A .

proteins

The

peak

by a n o t h e r

as T C - 2 . FAD

the

8.0)-0.1

subunit

pattern

fractions

[ 1 4 C ] I A M for

1 mM other

subunit

of e a c h

fluorescence

covalent

from The

HPLC

^C-labeled

Fractions

yellow

with

extensively.

by H P L C . of

peaks. at

labeled

mM p y r o p h o s p h a t e - H C l ( p H

subunit and

was

enzyme and of

As

the TC-1

sodiun

respect

sequence

of

must

to be

site.

modified

the Cys

residues

of

the

829 TC-1

and

TC-2

know

how

these

flavin against hrs

enzyme and

absence C.

with

absorbance

fractions

with

performed

of

the

IAM-treated

rates

catalyzed

with

If the

activity

of

but

for

described

the

enzyme

according

site (possib1y residues

be c o m p l e t e l y a covalent

Remarks:

are

This

with

to 9 0 %

for

without inhibited that

sites,

o n e at

IAM, of

only

FAD

site)

overall

reaction

not

noncovalent

and

FAD

be

mM

inhibited

sulfite

that

the

enzyme, the

10% mM

idea

of

FAD,

20

two C y s

the since

sulfite

residues

of

sarcosine-binding the

By t h e m o d i f i c a t i o n

s a r c o s i n e (see f i g u r e

of

oxidation

with

showed

site.

treatment

complex(_2).

work

noncovalent

of

covalent

FAD-sulfite

observed

covalent

support

is s u g g e s t e d

were

absence

the native

These

enzymes

in t h e

20

at t h e d i f f e r e n t

FAD-binding

the o x i d a t i o n

it

sarcosine

should

and

of

with

IAM

the

enzymes

reduction

in t h e only

with

enzymes.

Moreover,

should

that

inhibited

oxidizes

to Kp of

Concluding

was

enzymes

found

it w a s of

to f u n c t i o n case,

the

acetate.

treated

and

the

enzyme)

experiments

phase

as t h o s e

time

phases.

absorbance

for

the

sulfite,

by

The

total

Similar

that

is the

was

of

treated

slow

of

for

conditions

the native

absence

to r e a c t

became

IAM-treated

of

the f a s t

indicate

IAM-treated

above.

enzyme

20 mM of

the

phase

EDTA

at 4 5 5 nm.

respectively and

the same

this

It

enzyme

of

FAD

to

the

dialyzed

mM the

and

to the

in

then

anaerobic

fast

of

I AM

apparatus»

forms

6.3 s

is k n o w n

data

sulfite.

20%

the f a s t and

were

noncovalent

sarcosine.

phase

and

rates

sulfite

these

covalent

of

and

enzymes

only

enzyme

55%

function with

8.0)-0.1

under

showed

reduced

interesting

acetate,

monitored

in the p r e s e n c e

to be 7.4

As

F A D (2),

native

fast

in t h e p r e s e n c e

sulfite.

was

change

minus

to be

IAM

the f r a c t i o n s

not

change

oxidized

respective

with

on t h e

mM s o d i u m

a rapid

is

modified

1 mM s a r c o s i n e

the a b s o r b a n c e

were estimated

the

Using

mixed

of

treated

of

of 50

was

mixing

the

enzyme

affect

enzyme

It

at 5

were determined

that

The

B.

buffer(pH

change(the

the

subunit

20 mM p y r o p h o s p h a t e - H C l was

course The

and

of

modifications

moieties.

presence 12

sequences

became

in t h e n e x t

other of

at

the

these

Cys

functional page).

in

830

1AM . Acetate.-treated

IAM-treated

Native

Sar

Figure. S c h e m a t i c r e p r e s e n t a t i o n of the o x i d a t i o n of s a r c o s i n e with the native, IAM-treated and acetate-IAM-treated enzyies. N and C r e p r e s e n t n_oncovalent and c o v a l e n t flavin moieties» respectively. Sar indicates s a r c o s i n e and the larger arrow m e a n s t h e l a r g e r a m o u n t of e l e c t r o n s t r a n s f e r e d f r o m sarcosine to o x y g e n .

Acknowledgements

We

like

Noda

to

Institute

sarcosine for

express

his

for

our

Scientific

oxidase.

valuable

sincere

We

are

thanks

Research,

also

discussions

to

Dr. for

indebted

during

the

to

Masaru his

kind

Prof.

course

Suzuki,

of

supply

Shigeo this

of

Horie

work.

References

1.

Suzuki,M..

1981.

J.Biochem.

2.

Jörns,M.S..

3.

Kawamura-Konishi,Y 915. 3 4 6 - 3 5 6 .

4.

Suzuki,H. proteins.

5.

Shiga,Y., S.Hayashi, M.Suzuki, K.Suzuki and 1983. B i o c h e m . I n t e r n a t i o n a l . 6., 7 3 7 - 7 4 2 .

6.

Hayashi.S., M.Suzuki, Biochin.Biophys.Acta.

1985.

8£.

Biochemistry. and

H.Suzuki.

599-607. M .

3189-3194.

1987.

Biochim.Biophys.Acta.

a n d Y. K a w a r n u r a - K o n i s h i . 1 9 8 7 . F l a v i n s W a l t e r de Gruyter, B e r l i n New York.

and 42,

S.Nakamura. 630-636.

1983.

and pp.

Flavo917-920

S.Nakamura..

Flavoproteins of Medical Relevance

MOLECULAR STUDIES OF TRYPANOTHIONE REDUCTASE: AN ANTIPARASITIC TARGET ENZYME

C. Walsh, M. Bradley, S. Sobolov, F.X. Sullivan Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School Boston, Massachusetts 02115

This chapter describes recent advances in the characterization of an enzyme that catalyses the reduction of a cyclic disulfide derivative of oxidized glutathione, termed trypanothione because of its occurrence in trypanosomatid parasites. It seems appropriate to start with a brief introduction on the parasites, some of global significance in human pathogenicity, then to examine the structure and function of trypanothione to understand the metabolic context in which the enzyme trypanothione reductase may be considered an antiparasitic target. In the protozoan phylum there is an order termed kinetoplastida wherein each representative contains a subcellular organelle, the kinetoplast, which has analogies to mitochondria but contains maxi and minicircles of DNA. The trypanosomatid family encompasses Crithidia which are insect parasites and Leishmania and Trypanosoma which parasitize vertebrates including man. Thus the leishmanial diseases, including cutaneous, mucocutaneous, and visceral forms, are caused by these kinetoplastid trypanosomatid leishmanial species. In Africa, trypanosomes are causative agents of sleeping sickness in man (T. brucei gambiense and T. brucei rhodesiense) and in cattle (T. congolense) while in South America T. cruzi causes Chagas' disease, a chronic and often fatal illness.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

834

Figure 1 Oxidized o xidized Trypanothione

T COO' ' " ^ C - Y ^ a ^ O H3N H O V NH

\ 0

(

(cp 2 ) 3

NH 2 + (CH2)4 H NH NH

H COOData exists suggesting trypanosomes are sensitive to oxidant stress generated during host metabolism (1). In particular while trypanosomatids have superoxide dismutase, they lack detectable hemeproteins such as catalase so their ability to detoxify and scavenge peroxides should depend heavily on the glutathione peroxidase/glutathione reductase couple. On initial examination of parasite glutathione reductase to assess any possible distinctions from host enzyme that might offer the prospect of differential inhibition and trypanocidal potential, Cerami and colleagues (2) discovered that in all trypanosomatids examined, glutathione levels were low and the bulk of (oxidized) glutathione was present as a cyclic N1,8 bisglutathionyl spermidine, trypanothione (fig. 1). Further, there was no detectable glutathione reductase activity (3) (eq. 1 in fig. 2) in trypanosomatid extracts. The biogenesis of trypanothione via the acyclic glutathionyl spermidine intermediate has been determined (4) but why trypanosomatids contain trypanothione is not yet understood.

As an initial probe for host vs parasite specificity in this oxidant stress response pathway we undertook the purification of the trypanothione-specific reductase (eq. 2 in fig. 2). To obtain sufficient biomass as a starting point and to minimize potential pathogenicity the starting organism for enzyme isolation was Crithidia fasciculata. Adaptation of purification procedures for glutathione reductase (GR) led to

835

Figure 2

°

/ o' \ .s

H

NADPH v V

S

+

0 \

H

HsN^v^M^N^COr co 2 H o Oxidized Glutathione Ci O , "

„H 0

o u

)

+ CO," Y H H u0 J H3N ^fNVlN^C02o \ h (i)

NADP / 2 y ^

Glutathione Reductase

® Reduced Glutathione

(GSSG)

n H

s

CO,-

nh

CH2)3

\

JHK V 2>4 „J 2 V M IH H a N ^ ^ N ^ o C0 2~ H O Oxidized Trypanothione

H

0 nadph

V^

O

\

NADP+

S

A

h

Trypanothione Reductase (TSST)

hs

H

NH

W

VyHj) 4 + 0 \ H nh H 3 N ^ n \ n ^ 0 C02- H o

Reduced Trypanothione

Table 1 Comparison of Key Catalytic and Structural Features in Human Erythrocyte Glutathione Reductase and Crithidia fasciculata Trypanothione Reductase Glutathione Reductase FAD NADPH Yes 52,000 dimer 460nm Yes 530nm

Property Flavin Pyridine Nucleotide Redox Active disulphide M r of monomer Oligomeric structure E 0 x ^max Charge Transfer in EH2 EH2 X m ax

Trypanothione Reductase FAD NADPH Yes 53,000 dimer 464nm Yes 530nm

straight forward purification to homogeneity of milligrams of trypanothione reductase (TR) (5).

Initial characterization

revealed many similarities to human glutathione reductase as noted in Table 1.

Each is an FAD-containing homodimer with a

836

Table 2 Specificity Constants for Human Erythrocyte Glutathione Reductase and T.Congolense Trypanothione Reductase. Trypanothione Glutathione Reductase Reductase 0.84M -1 s _1 GSSG 188xl0 6 M -1 s -1 505xl06 TSST 6x10® Ratio >2xl0 * Assumming a K m >10raM reducible active-site disulfide and high homology in the active site bi-cysteine-containing peptide. A striking distinction, however, emerged in the specificity assays for human red blood cell glutathione reductase vs trypanothione reductase (table 2). Each reductase displayed >105 to 10® fold preference in kcat/Km for homologous over heterologous substrate, suggesting the possibility of specific inhibition of parasite enzyme in the presence of host reductase. The next stage of analysis of the molecular basis of host/parasite enzyme specificity required the availability of primary sequence information for a trypanothione reductase for comparison to human GR enzyme. To this end peptide sequence data on the crithidial TR were used to construct oligonucleotide guessmers (based on uncertain codon usage in trypanosomatid genomes) to screen trypanosomatid cDNA libraries (6). The most promising hybridization response was with DNA from the cattle parasite T. congolense and subcloning and sequencing was pursued to yield a 1476 bp open reading frame with 41% homology to human GR. While it seemed highly probable the T. congolense TR had indeed been cloned, this expectation was then experimentally validated by heterologous expression in E. coli. Thus the T. congolense gene yielded overproduction of a novel enzyme activity in E. coli, the anticipated NADPH-dependent trypanothione reductase (7). It has been possible to purify the T. congolense TR from E. coli extracts to homogeneity in quantity and so sufficient

837 pure enzyme is now available for structure/function The T. congolense

studies.

TR, like crithidial TR, displays negligible

glutathione reductase activity while showing a high kcat (9,000 min~l) for oxidized trypanothione. At this point with a cloned, sequenced TR gene in hand and TR heterologously expressed, overproduced, purified, and kinetically characterized, one could proceed to the next stage of inquiry into why these trypanothione reductases are not glutathione reductases and vice versa. Because of the existence of a high resolution X-ray structure of human GR (8) and GR-NADP, and GR-NADP-GSSG complexes, functional assignments of key residues in GR catalysis have been made. As table 3 demonstrates there is very good conservation of cognate amino acids in TR and in GR with one notable exception; Arg 37 in GR has Trp 21 in a homologous location in T. congolense

TR.

Arg 37 makes a close contact with the glyl-carboxylate of the GSl-moiety of GSSG bound in the active site of human red blood cell GR. This gly-COO® anion is not present in TSST for a similar electrostatic interaction with a TR side chain, the glyl carbonyl of TSST now being an amide link in the spermidine bridge. The second notable distinction in Table 3 Table 3 Comparison of Key Residues in Human Erythrocyte Glutathione Reductase and T.Congolense Trypanothione Reductase. Glutathione Reductase GGGSGGL-35 Cys-58 Cys-63 Tyr-114 His-467 Glu-472 Arg-37 Arg-347

Function from X-ray Structure FAD binding site Redox Active disulphide Redox Active disulphide Stacks between GS moieties in glutathione Active site base H bonds to active site His Binds glycyl carboxylate of GSi via H bond Binds y-glu carboxylate of GSi via H bond Unknown

Trypanothione Reductase GAGAGGL-47 Cys-52 Cys-57 Tyr-110 His-461 Glu-466 Trp-21 Ala-343 20 aa tail

838

Figure 3

J L

Hi 8467

N-

S-S

His 461i

N-

s-s _L_L

493* wt (-5) P488* (-10) K483* (-15) Y478 * (-20) T473*

(GR)

C

(TR)

RTPSHYYIKGBKMETLPDSSL RTPSHYYIKGEKMETL RTPSHYYIKGE RTPSHY R

is the addition of an extra 20 amino acids at the C-terminus of TR with no analog in GR. As noted in figure 3 the 20 residue tail of TR has three negatively charged side chains. It is known that the C-terminus of GR loops back to provide His 467' as the key active site base to interact with Cys 58 and Cys 63 on the other subunit of the dimer. Thus, the possibility existed that the anionic C-terminal tail had a substantial role in establishing the specificity of TR for TSST, for example by electrostatic interaction with the cationic central ammonium group in the spermidine bridge of TSST. In ongoing studies in our group we are assessing the role of the C-terminal tail of TR by deletional mutagenesis and of the R37 residue of GR by site-directed mutagenesis. By systematic deletion of 5 residues at a time, we have constructed, expressed, and purified the -5 mutant TR (P488*) and analogously the -10 (K483*), -15 (Y478*) and -20 mutants (T473*), this last corresponding to "glutathione reductase" length. In the event all four of the truncated mutant TRs have essentially wild type catalytic efficiencies (kcat/Km) for TSST reduction (9). Also, none had regained detectable glutathione reductase activity. Obviously this 20 residue

839

Figure 4

tail does not control TSST/GSSG recognition or discrimination. We did note that TR 478 and TR 473 were purifiable in -103 fold lower amounts than the longer mutants and wildtype TR. Whether this low yield reflects unstability

(e.g. to

proteolysis) and/or folding problems is not yet determined. In further analysis of contacts of enzyme to substrate in the GSSG-GR complex Karplus and Schulz noted that Arg 347 made a close contact to the glul-a-COO®

of bound GSSG. They also

pointed out that 14/19 residues lining the GSSG binding site are conserved in TR. In addition to the lack of Arg 37 and Arg 347, TR was also observed to contain a potential binding residue homolog to the Ala 34 found in GR, a glutamic acid residue at position 18. The orientation of these three residues with respect to bound GSSG is displayed in figure 4.

840

In T. congolense TR the corresponding residues are (GR:TR):A34:E18; R37:W21; R347:A343. To determine whether replacement of Glu 18, Trp 21, or Ala 343 by the side chains found in GR would now permit acquisition of glutathione reductase activity by trypanothione reductase we have made the mutations E18A, W21R, A34R in the T. congolense TR gene as single mutants, double'mutants, and triple mutant and have purified all seven mutant enzymes to homogeneity. Happily, one double mutant, E18A W21R, and the triple mutant TR did indeed gain glutathione reductase activity. For example the triple mutant, which is ca. 4 fold better than the double mutant, has a kcat/Km for GSSG reduction 17,000 fold up from wild type GR (kcat/KmcssG 0.84min~lM~l). In the process the TSST reduction catalytic efficiency dropped ca 103 fold. While the catalytic efficiency of wild type TR for TSST/GSSG was ca 5 x 1 0 8 / 1 the triple mutant is down to 28/1. One is beginning to learn the determinants of host vs parasite enzyme specificity. It remains to translate these findings into structural information both for antiparasitic drug design and for predicting recognition rules from enzyme structure. Wild type T. congolense TR does crystallize (10), although not yet in crystallographically useful form and ultimately an X-ray structure of both wild type TR and E18A,W21R,A343R TR (with TSST or GSSG bound) as a hybrid TR/GR would be helpful in going to the next stage of knowledge. Whether TR will indeed prove to be a trypanosomatid parasitic enzyme that can be selectively attacked in leishmaniases, sleeping sickness, and Chagas' disease remains to be proven as does the question of whether selective TR inactivation will prove a trypanocidal strategy. Nonetheless the combined approaches of modern molecular biology and enzymology have

841

rapidly moved this prospective target for antimetabolites from an undefined biological activity to a well-characterized molecular entity, suitable for rational medicinal chemical strategies.

References 1.

Meshnick, S.R., S.H. Blobstein, R.W. Grady, A. Cerami. 1985. J.Exp.Med. 148r 159. Boveris, A., H. Sies, E.E. Martino, R. Docampo, J.F. Turrens, A.O.M. Stoppani. 1980. Biochem.J. 1 M , 643.

2.

Fairlamb, A.H., A. Cerami. 1985. Mol.Biochem.Parasitol..14. 187.

3.

Fairlamb, A.H., P. Blackburn, P. Ulrich, B.T. Chait, A. Cerami. 1985. Science 227. 1485.

4.

Henderson, G.B., M. Yamaguchi, L. Novoa, A.H. Fairlamb, A. Cerami. 1990. Biochemistry 22, 3924.

5.

Shames, S.L., A.H. Fairlamb, A. Cerami. C.T. Walsh. 1986. Biochemistry 25, 3519.

6.

Shames, S.L., B.E. Kimmel, O.P. Peoples, N. Agabian, C.T. Walsh. 1988. Biochemistry 22, 5014.

7.

Sullivan, F.X., S.L. Shames, C.T. Walsh. 1989. Biochemistry 2fi, 4986.

8.

Karplus, P.A., E.F. Pai, G.E. Schulz. 1989. Eur. J. Biochem. 123., 693.

8.

Karplus, P.A., G.E. Schulz. 1989. J.Mol.Biol. 2112, 163. Pai, E.F., G.E. Schulz. 1983. J. Biol Chem 2 M , 1751.

9.

Sullivan, F.X., S.B. Sobolov, M. Bradley, C.T. Walsh, unpublished observations.

10. F.X. Sullivan, E.F. Pai, Luise Krauth-Siegel, C.T. Walsh, unpublished observations.

LIPOAMIDE DEHYDROGENASE AND TRYPANOTHIONE REDUCTASE FROM TRYPANOSOMA CRUZI. THE CAUSATIVE AGENT OF CHAGAS'DISEASE R. Luise Krauth-Siege^ Helmut Lohrer, Klaus-Dieter Hungerer* and Till Schöllhammer Institut für Biochemie II, Universität Heidelberg, FRG Behringwerke Marburg, FRG Medizinische Poliklinik, Universität Heidelberg, FRG INTRODUCTION Trypanosomes and leishmanias do not possess glutathione reductase. The main thiol compounds are conjugates between glutathione and spermidine - like trypanothione - which are kept in the reduced state by the flavoenzyme trypanothione reductase (TR) : TS2 + NADPH + H + T(SH)^ + NADP+. Since this specific redox metabolism is a most promising target for a rational drug design against trypanosomes it should be known which other thiol-regenerating enzymes are available to the parasite. One candidate is lipoamide dehydrogenase+ (LIPDH) : lipoamide + NADH + H + ^ > dihydrolipoamide + NAD because the catalyzed reaction is freely reversible so that the enzyme might reduce lipoyl disulfide at the expense of NADH under certain metabolic conditions. In this context it is of interest that long slender bloodstream forms of T. brucei contain a plasma membrane-associated LIPDH (1). Here we report on the properties of lipoamide dehydrogenase and trypanothione reductase from T. cruzi. the causative agent of American trypanosomiasis. RESULTS AND DISCUSSION Characterization of T. cruzi LIPDH Lipoamide dehydrogenase ana trypanothione reductase can be isolated from the same extract of T. cruzi epimastigotic cells using 2'5'ADP-Sepharose as the first purification step. The NADPH-dependent TR binds to the affinity matrix whereas LIPDH is quantitatively washed through (2, 3). The concentrations of the enzymes in the epimastigotes indicate that they are major proteins of the parasite (Table). J. cruzi LIPDH is very similar to the enzyme from other organisms (4, 5). It is specific for lipoamide as a disulfide substrate; glutathione and trypanothione are not reduced. Lipoate is slowly turned over by the enzyme, the activity being less than 2% with 1 mM lipoate when compared with lipoamide (3). This indicates that the natural substrate of T. cruzi LIPDH is a protein-bound lipoyl residue as it is the case in the 2-oxoacid dehydrogenase complexes of other organisms.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin-New York-Printed in Germany

844

Table: Some chemical and physical properties of lipoamide dehydrogenase and trypanothione reductase from T. cruzi(2.3,6). lipoamide dehydrogenase Subunit M r Cofactor Oligomeric structure Intracellular concentration (/¿M)

55.000 FAD dimer 0.9

trypanothione reductase 50.000 FAD dimer 1.25

Substrates at the pyridine nucleotide site: NADH NADPH activity with 100 (M NADH/ activity with 100 /xM NADPH

K_= 23 MM n.d.

K_= 600 MM ICm= 5 MM

590

0.03

Substrates at the disulfide site: lipoamide glutathione trypanothione

Kj^ 5mM no substrate no substrate

n.d. no substrate K,n= 18 MM

Specific activity (U/mg)

297

100

Catalytic disulfide

yes

yes

Highly reactive cysteine residue in the reduced enzyme species EH 2

yes

yes

Specific inhibition of EH 2 by BCNU

yes

yes

452

461

Absorption band at 530nm in EH 2

yes

yes

A

530nm

of

lmM

EH

2.8

A

530nm

of

lmM

EH

¿max,vis t ™ )

2 2

'

NADH

NADPH

4.9

This is consistent with preliminary immunofluorescence studies on epimastigotic cells using polyclonal antibodies raised against T. cruzi LIPDH and TR, respectively. These experiments gave no indication for T. cruzi LIPDH being localized in the plasma membrane as it has been reported for the enzyme from bloodstream T. brucei (1) .

845 Out of the 30 N-terminal amino acid residues of T. cruzi LIPDH, 21 positions were found to be identical with the sequence of pig heart and human liver LIPDH (3) . The parasite enzyme has the shortest N-terminal extension of all known LIPDHs: it starts only 2 residues in front of the first sheet strand of the FAD-binding domain when compared with the structure of glutathione reductase. Fig. 1: Structures of N 1 -qlutathionvlspermidine and

15-deoxv-

spercrualin. N^—glutathionylspermidine

COOH 0 0 0 i u u 11 HC-CH2-CH2-C-NH-CH-C-NH-CH2-C-NH-(CH2)3-NH-(CH2)4-NH2 nh 2 ch2 SH 15-deoxyspergualin OH 0 0 U I II HN=C-NH-CH 2 -(CH 2 ) 5 -C-NH-CH - C-NH-(CH 2 ) 4 ~NH-(CH 2 ) 3 "NH 2 I NH

Fig. 2: Inhibition of T. cruzi trvpanothione reductase by 15-deoxyspergualin. The kinetics were measured in 40 mM Hepes, lmM EDTA, pH 7.5, the NADPHconcentration being 100 MM. The glutathionylspermidine disulfide concentrations were 44/iM (•), 66 mm (•) and 110/iM

(A) .

846

Inhibition of trypanothione reductase bv 15-deoxvsperqualin. The structural similarities of glutathionylspermidine and the immunosuppressive agent 15-deoxyspergualin (7)(Fig.l) suggested that the latter is an inhibitor of TR. The Dixon-plot (Fig. 2) shows a competitive type of inhibition, the inhibition constant being K^= 60 jXM (8). It will be of interest to study if the glutathionylspermidines of T. cruzi play a role in the immunosuppression observed in acute Chagas' disease. This would be an additional function of the unique trypanothione metabolism of Kinetoplastida. N8-glutathionylspermidine - which was not available to us in pure form - is the best candidate for these studies since its structural similarity with 15-deoxyspergualin is more pronounced than that of the N^-isomer. Crystals of trypanothione reductase. At present we grow crystals of TR from T. cruzi. T. conaolense and Crithidia fasciculata (The enzymes were kindly provided by F.X. Sullivan and C.T. Walsh) . For each species crystallization conditions have been established (2, 9) but the crystals are not yet suited for highly resolved X-ray diffraction analyses. The structural differences between the host glutathione reductase, for which the 3-dimensional structure is known at 0.15 nm resolution, and trypanothione reductase should allow the design of specific inhibitors against the parasite enzyme. REFERENCES 1.) Danson, H.J., K. Conroy, A. McQuattie, K.J. Stevenson. 1987. Biochern. J. 243. 661-665. 2.) Krauth-Siegel, R.L., B. Enders, G.B. Henderson, A.H. Fairlamb, R.H. Schirmer. 1987. Eur. J. Biochem. 164. 123-128. 3.) Lohrer, H., R.L. Krauth-Siegel. 1990. Eur. J. Biochem. submitted. 4.) Williams, C.H. Jr. 1990. In: Chemistry and Biochemistry of Flavoenzymes (Müller, F., ed.). CRC Press, Boca Raton, in press. 5.) Schirmer, R.H., G.E. Schulz. 1987. In: Coenzymes and Cofactors (Dolphin, D., ed.). Wiley, New York, pp. 161-204. 6.) Jockers-Scherübl, M.C., R.H. Schirmer, R.L. KrauthSiegel. 1989. Eur. J. Biochem. 180. 267-272. 7.) Dickneite, G., H.U. Schorlemmer, P. Walter, J. Thies, H.H. Sedlacek. 1986. Behring Inst. Mitt. 80, 93-102. 8.) Schöllhammer, T., MD Thesis, Universität Heidelberg, 1989. 9.) Sullivan, F.X., R.L. Krauth-Siegel, E.F. Pai, C.T. Walsh. 1990. In: Protein and Pharmaceutical Engineering. WileyLiss, New York, pp. 119-134.

BINDING OF FAD TO BCNU-TREATED APOGLUTATHIONE REDUCTASE

Katja Becker, Till Schöllhammer and R. Heiner Schirmer Institut für Biochemie II and Medizinische Poliklinik der Universität, D-6900 Heidelberg, FRG.

Introduction As indicated by experimenta naturae (riboflavin deficiency, glucose-6-phosphate dehydrogenase deficiency) an impaired activity of the erythrocytic flavoenzyme glutathione reductase (GR) protects from malaria. Since this antioxidant protein, catalyzing the reaction NADPH + H+ + GSSG NADP+ + 2GSH (EC 1.6.4.2), is well known it is considered as a target molecule for systematic drug design (1). BCNU (carmustine) is a clinically used cytostatic agent which also has antimalarial activity (2). This compound inhibits the 2-electron-reduced form (EH2) of glutathione reductase by carbamoylating Cys58, one of the active site thiols (3); the oxidized form (E) containing Cys58-Cys63 as a disulfide is not affected by BCNU. The effect of BCNU on FAD-free apoglutathione reductase has not yet been investigated although the apoenzyme can be the predominant form of erythrocyte GR in malaria-afflicted countries where riboflavin deficiency is widespread (4). A prevalence of apoGR is assumed to mitigate the course of tropical malaria (4); on the other hand a malaria therapy based on GR-inhibitors would be inefficient if the apoenzyme - being potential active holoenzyme - cannot be modified by substances like BCNU and its less toxic derivatives.

Results and Discussion Modification of hologlutathione reductase and its apoenzyme in vitro. The inhibition tests on isolated apo- and holoGR were carried out in the GR assay system of Worthington and Rosemeyer (5) which was slightly modified (6). As expected (2), 1 mM BCNU inhibited NADPH-reduced holoenzyme by more than 98% (Table 1). Preincubation with dithioerythritol (DTE), which is known to reduce the active site disulfide (1), followed by

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter&Co., Berlin • New York - Printed in Germany

848 Table 1. Inhibition of purified hologlutathione reductase and its apoenzyme by BCNU. The drug (1 mM) was added after the enzyme had been preincubated in assay buffer (S) with different compounds for 15 min. Residual activity was determined after appr. 2 h.

Conditions

Maximal modification by BCNU HoloGR

ApoGR

no reducing agent

9%

87%

400 |i.M DTE

12%

95%

200 |!M NADPH

98%

79%

3 mM GSH

19%

46%

200 llM NADPH + 3 mM GSH

n.d.

61%

200 IIM NADPH + 1 mM GSSG

6%

40%

addition of BCNU resulted in only 12% loss of activity. This finding was not due to a reaction of the drug with DTE (6) and remains to be studied further. Isolated apoGR could also be modified by BCNU (Table 1). Best yields were obtained after preincubation with 400 p.M DTE but also in the absence of added reducing agents. Our conclusion that the Cys58-Cys63 pair of apoGR is present as a dithiol and not as a disulfide (6) was confirmed by crystallographic studies of Ermler and Schulz (7). As shown in Table 1, the two cysteines are protected by glutathione. Since the flavin barrier is absent (Fig. 1) the BCNU-derivatives might reach the active site thiols also via the NADPH-compartment. This route, however, seems unlikely in view of the insignificant protecting effect of NADPH. This substrate of holoGR binds tightly to apoGR (unpublished results). Spectroscopic characteristics of the BCNU-treated protein after complementation with FAD. The absorption spectrum of this yellow protein species is identical with that of oxidized holoGR. In contrast, when holoGR is treated with BCNU the resulting spectrum corresponds to that of the orange-coloured EH 2 form (3, 8). This difference may be due to a modification of both active site thiols in the apoenzyme. Alternatively, one of the cysteines might be modified in such a way that the resulting geometry rules out the charge transfer

849

Figure 1. Sketch of the "active site" in apoglutathione reductase. As revealed by chemical studies (6) and X-ray diffraction analysis (7) the structure of the active site is preformed in the apoenzyme. When FAD is complexed the flavin ring is squeezed in between the nicotinamide-binding site and the redox-active dithiol Cys63-Cys58. The disulfide of Gly-Cys-^Glu at the bottom designates the binding site of GSSG.

300

400

500 600 wavelength

nm

Figure 2. Absorption spectrum of BCNU-treated apoglutathione reductase after reconstitution with FAD in the presence of dithioerythritol and BCNU derivatives. The peak at 414 nm is characteristic for the semiquinone anion of flavoproteins (8,9). Removal of the reagents by dialysis leads to the standard spectrum of oxidized hologlutathione reductase (6,8).

850 complex between thiolate 63 and flavin which is colourfully manifested in the Massey-Williams spectrum of EH2 and EHR forms (8). When DTE and excess BCNU had not been removed before adding FAD a spectrum with an additional absorption maximum at 414 nm was observed (Fig. 2). As this peak is characteristic for the anionic flavin semiquinone (8,9), the experiment indicates that BCNU can give rise to radical formation in a flavoprotein; this is a new aspect in the pharmacology of nitrosoureas. Modification of apoglutathione reductase by BCNU in intact erythrocytes. Red blood cells containing a high proportion of apoGR were identified by using a standardized EGRAC-test (10). When exposing these cells to 500 nM BCNU (2, 6), the holoenzyme could, as expected, be inhibited by > 90%; in contrast, the apoenzyme was modified by only < 25%. This different behaviour of apoGR and holoGR was manifested in the EGRAC value which was 1.5 before and > 5 after treating the cells with BCNU. The resistance of the intracellular apoenzyme against modification by BCNU remains to be explained. As a first step we plan to determine the dithiol/disulfide ratio for the redox pair Cys58-Cys63 under in vivo conditions.

References 1. Schirmer, R.H., R.L. Krauth-Siegel and G.E. Schulz. 1989. In: Glutathione, Vol. A (D. Dolphin, R. Poulson and O. Avramovic, eds.). John Wiley and Sons, New York. pp. 553-596. 2. Zhang, Y„ E. Hempelmann and R.H. Schirmer. 1988. Biochem. Pharmacol. 37. 855-860. 3. Karplus, P.A., R.L. Krauth-Siegel, R.H. Schirmer and G.E. Schulz. 1988. Eur. J. Biochem. 171, 193-198. 4. Bates, C.J.. 1987. Wld. Rev. Nutr. Diet. 5Q, 215-265. 5. Worthington, D.J. and M.A. Rosemeyer. 1976. Eur. J. Biochem. SL 231-238. 6. Becker, K.. Dissertation, Universität Heidelberg 1988. 7. Ermler, U.. Dissertation, Universität Freiburg 1989. 8.Williams, C.H.Jr. 1976. The Enzymes XIII, 89-173. 9. Massey, V. and G. Palmer. 1966. Biochemistry 5, 3181-3189. 10. Becker, K., B. Krebs and R.H. Schirmer. Internat. J. Vitam. Nutr. Res., submitted.

THE EGRAC AS A MEASURE OF THE RIBOFLAVIN STATUS IN MAN. TITRATION OF HEMOLYS ATE FAD WITH APOGLUTATHIONE REDUCTASE

Katja Becker, R. Heiner Schirmer Institut für Biochemie II der Universität, INF 328, D-6900 Heidelberg, FRG.

Introduction Human erythrocyte glutathione reductase (GR) is a homodimeric flavoenzyme of 105 kDa (1, 2). In vivo

GR is not fully saturated with its prosthetic group FAD but the inactive

apoenzyme can be completed in vitro according to the equation: apoGR + FAD — h o l o G R . This is the hypothetical basis for the determination of the erythrocyte glutathione reductase activation coefficient (EGRAC) which is defined as the ratio of FAD-stimulated to unstimulated activity of erythrocytic glutathione reductase (3). The activation coefficient is a reliable index for riboflavin deficiency in man (1). The multifacetted medical aspects, including interference with embryonic development and impaired protection from noxious chemicals, of this widespread hypovitaminosis have been reviewed comprehensively (1,4).

Results

and

Discussion

The clinical interpretation of the EGRAC test is based on a set of equations

(Eqn. 1):

EGRAC = total GR / holoGR = (holoGR + apoGR) / holoGR = 1 + apoGR / holoGR. In order to test this hypothetical interpretation, the interaction of apoGR and FAD in hemolysates was studied by titration. The individual steps of the experiment which is summarized in Table 1 are to be explained here. We started out with a concentrated hemolysate, its EGRAC being 1.25. According to the law of mass action the free FAD concentration was expressed as a multiple of the dissociation constant: [free FAD] = K Il 3 C 3 -u Ä *0 ld c O IS «I i> M ai m > c u « i O II04JH •w G a O ja -h «C O 3 J3 o H m i C

fl KO CM CM CN

, > o o r- m x o • -p -u

M ti o m o ä ra id -P — >, X >, H II M öl O 1141 E II •H II X! G C G J3 +1 O ld 3 •HE Ol 41 3 • J3 •P •H }

CM tM I I

(N (N r— t— I I

i l

i l

u < s D

id J3 e o tN

H >1 c — 4 1 « N II •H Ol c o> c Eh -H II O O m "O -H >i-H O C -P XI -H II 3 3+1 .-I h a aw o •P >1 O 01 id >i M u h o X C4i >.X II O X rH Cr in j a -h D O "H • E 4 J 4 I e S m 3 id Il m c kl « S E -P.C O T3 kl -H 4J C M II 41 4J •H K4 G > C •H O II -H D ' O n c i < 41 JJ kl 4J "O 6. M C 3 II (ti O -P £ G Il O. O -H -H si ai D X K C £

h

•u O c O •h •P

id

0.0 h •H n 4-> • Id 3 « S H Ü o o — oi m in o — c X

o

N r - r - (N

m t - tN m

en T- ro m a» \o oo cm

m cm n i-i-oin O O O (N

O O O n N r - o o o o m

^ ^ o O

^ o CM es en o o co O O 1-

id

(IH

«

tt X X il 0> -H ttl JC M n m Id Ol

«H

e « -H ~ Il -rt •H IS -P IH O Id e o n) o -h 4J v a n e s a ) « a 4i c 4> Q •P -H II II 4> VO II • Q C O û 41 •H H • C • «H E T- O II • r- M — 4J E id • •H D 4J m TI kl i-I 01 • 4J II II XI > , — - H E a Id 14 M c 3 X EH U — 3 01 U

•8 5 ? u S id 8 3 H

41 O r t R s

-P

3

853 the apoenzyme activity (0.80 cryptic U/ml) was by Ac = - 0.53 U/ml (103 nM) lower than expected. "Expected" means that added apoGR would not readily associate with FAD which is contained in the hemolysate. The data, however, indicate that a proportion of the added apoGR was indeed complemented with its prosthetic group to give holoenzyme. Accordingly, the increase in holoGR (Ac = 106 nM) corresponds numerically to the decrease in the level of FAD. The free FAD concentration equals 4 K^ in the untreated hemolysate and 2 K^ after adding apoenzyme. These relations yield the equation Ac = 106 nM = 4 K^ - 2 K^ from which a K-

L

60

RETENTION TME (n*i) Fig.1: Anion Exchange HPLC of Neutrophil Cytosol on DEAE-5PW. Fresh cytosolic fraction from dormant cell was applied to a column. A linear gradient was run from buffer I (50mM Hepes, pH 7.6, containing 1mM MgCl2 and 20% glycerol) to buffer II (buffer I plus 0.5M KCI). Inset shows SDS-PAGE of the cytosolic fraction A (left lane) and marker proteins of molecular weight standards (116, 92.5, 66.2 and 45 kDa, respectively on right lane).

region in this fraction. Cytosolic proteins involved in fraction A are susceptible to undefined endogeneous proteases and almost completely converted to proteolysis products upon incubation at 36°C for 18h, resulting in loss of activation of superoxide anion production and NADPH:NBT reductase activity. In contrast, the plasma membrane fraction prepared from phorbol 12-myristate 13-acetate (PMA)-stimulated cell provided high NADPH:NBT reductase activity (33 nmol/min/mg protein) as well as NADPH oxidase activity (41 nmol/min/mg protein).

The two enzymatic activities of the membranes from

dormant cell were low. Octylglucoside(OG)-solubilized plasma membranes, granules and cytosol from stimulated cells were loaded on polyacrylamide gel electrophoresis in the presence of 35mM OG and after PAGE the gels were stained for NADPH:NBT reductase activity (Fig. 2). Thus, we could see three major staining bands in extracts of plasma membranes whose molecular weights were approximately 95, 45 and 40kDa, respectively. Amoung them the protein with 45kDa showed markedly stronger NBT reductase activity, but its staining intensity from resting cells was considerably lower. On the HPLC column (DEAE-5PW) the membrane-bound 45kDa protein with NBT reductase activity also appeared at high molarity of KCI (0.32-0.35M) in the presence of 35mM OG and 20% glycerol, corresponding to a protein with acidic isoelectric point.

879 Fig. 2: OG- PAGE of Cytosol, Plasma Membrane and Granule Fractions Separated from PMA-activated Neutrophils. After electrophoresis the gel was cut into two fragments, one of which was used "for protein staining (gel lane 1 to 3) and another was stained for NADPH:NBT reductase activity (gel lane 4 to 6), respectively. Lanes 1 and 4, cytosol (0.22mg); Lanes 2 and 5, OGsolubilized granules (0.12mg); Lanes 3 and 6, OG-solubilized plasma membranes (0.12mg). Lane M contains marker proteins of known molecular weight.

10

20

30

40

50

60

Fig. 3: Effect of Cytosolic Fraction A on Superoxide Anion Production in a Cellfree System. The membrane fraction (0.17mg) from resting cells was preincubated with (curve C, 60jig; curve B, 15p.g) or without (curve A) cytosolic fraction A. Immediately after preincubation for 2min at 30°C, 0.2mM NADPH was added to both sample and reference cuvettes (indicated by arrow). Curve D shows a time course for 0 2 generation of membrane fraction (0.15mg) prepared from_ PMA-activated neutrophils (38 nmol 02/min/mg of membrane).

CYTOSOLIC FRACTION A (pg)

Figure 3 shows that the addition of cytosolic fraction A containing mainly 45 and 40kDa proteins to the cell-free activation mixture significantly enhances the extent of NADPH oxidase activity with its concentration dependence, although the duration of the stimulatory effect is less than 2 min. Thus, it appears that the cytosolic 45 or/and 40kDa proteins is involved in the oxidase activation (4), but simultaneous incubation of membrane fraction with both cytosolic fraction A and B which is unbound to DEAE-5PW ion exchange column gives no more remarkable effects. On the basis of these data, we suggest that during cell stimulation the cytosolic 45kDa component may be transformed into an activated form in the process of either

880 phosphorylation, dissociation or cleavage of precursory peptide(s) which shows higher affinity for NADPH rather than NADH and translocated to the membrane (Fig. 4).

In

addition, it appears that the lability of NADPH oxidase system of phagocytic cells is due probably to proteolysis of activated 45kDa protein by potent proteases derived from granules after stimulation of cells.

"Phosphorylation (PKC), Dissociation (arachidonate, SDS or DG) Cleavage of precursor (proteases)

Fig. 4: 45kDa Protein-dependent Activation Mechanisms of Superoxide Anion Production. Fp, flavoprotein; PKC, protein kinase C; DG, diacylglycerol.

Acknowledgement

We are grateful to Dr. M. Yamada (Osaka University, Protein Research Institute) for his generous gift of HL-60 cells.

References 1. Curnutte, J.T.,

R. Kuver,

P.J. Scott. 1987. J. Biol. Chem. 262, 5563-5569.

2. Volpp, B.D., W.M. Nauseef, R.A. Clark. 1988. Science 242, 1295-1297. 3. Heyworth, P.G., C.F. Shrimpton, A.W. Segal. 1989. Biochem. J. 260, 243-248. 4. Yea, C.M., A.R. Cross, O.T.G. Jones. 1990. Biochem. J. 265, 95-100.

COVALENTLY-BOUND FLAVIN IN PEROXISOMAL L-PIPECOLIC ACID OXIDASE FROM PRIMATES

S.J. Mihalik and M.C. McGuinness Kennedy Institute and Depts. of Pediatrics and Neurology, Johns Hopkins University School of Medicine, Baltimore, MD 21205

Introduction

L-Pipecolic acid oxidase, a component of the lysine degradation pathway via L-pipecolic acid (1), catalyzes the dehydrogenation of L-pipecolic acid to a-aminoadipic acid semialdehyde (2) in the peroxisome (3).

To date, this peroxisomal enzyme has been

found only in tissues from monkeys and humans (4).

In

patients with disorders of peroxisomal biogenesis, such as Zellweger syndrome (5), the activity of L-pipecolic acid oxidase is decreased and patients accumulate L-pipecolic acid. Patients with these disorders present with severe neurological dysfunction and profound mental retardation.

Their tissues

lack normal peroxisomes, as well as most peroxisomal metabolic functions (6).

Since L-pipecolic acid and its metabolites are

reported to be neuroactive agents, they may play a role in the pathophysiology of these disorders.

We report here on our

studies of the purification and characterization of L-pipecolic acid oxidase derived from monkey liver.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin-New York-Printed in Germany

882 Results L-Pipecolic acid oxidase has been purified, l i v e r , t o 96% h o m o g e n e i t y as j u d g e d f r o m

from frozen

densitometric

s c a n n i n g o f a 10% S D S - p o l y a c r y l a m i d e g e l of t h e protein.

purified

M o l e c u l a r w e i g h t of t h e p u r i f i e d p r o t e i n w a s

Da on SDS-polyacrylamide gels and molecular sieve s h o w e d t h a t it e x i s t s a s a m o n o m e r . (3).

Michealis-Menten

o f 3.7 m M w i t h t h e s u b s t r a t e

Benzoic acid competitively

o x i d a s e w i t h a K^ o f 0 . 7 5

was

described

The purified enzyme followed

kinetics and had a acid.

assay

46,000

studies

Enzyme activity

determined using a spectrophotometry previously

macaque

L-pipecolic

inhibited L-pipecolic

acid

mM.

The results of substrate specificity studies are s h o w n T a b l e 1.

L-Pipecolic acid was the major substrate

in

identified,

w h i l e i - p r o l i n e a n d s a r c o s i n e h a d 23 a n d 10%, r e s p e c t i v e l y , the activity of L-pipecolic acid.

However, the enzyme

of

showed

n o r e a c t i v i t y t o w a r d D - p i p e c o l i c a c i d a n d o n l y a b o u t 6%

toward

D-proline. The purified enzyme was a yellow protein with an

absorption

m a x i m a a t p H 7 . 5 o f 452 a n d 383 n m w i t h a s h o u l d e r a r o u n d nm.

T h i s s p e c t r u m is c o n s i s t e n t w i t h a p r o t e i n w h i c h

a tightly-linked

flavin.

When the protein was

w i t h 10% t r i c h l o r o a c e t i c a c i d ,

contains

precipitated

followed by centrifugation,

s u p e r n a t a n t d i d n o t a b s o r b a t 452 o r in t h e 3 4 0 - 3 8 0 n m T h e r e s i d u a l p r o t e i n p e l l e t , w h i c h w a s y e l l o w in c o l o r , resolubilized

in 5 M guanidine HC1.

a b s o r p t i o n m a x i m a a t 452 a n d 363 n m . filtered through a Centricon

This solution

was

had was

filtrate

These results suggest

L - p i p e c o l i c a c i d o x i d a s e c o n t a i n s a f l a v i n w h i c h is

the

range.

After this solution

(30,000 D a c u t o f f ) , t h e

d i d n o t a b s o r b a t 452 o r 363 n m . bound.

480

that

covalently

883 Table 1. Substrate Studies using Purified L-Pipecolic Acid Oxidase. Substrate

Relative Activity

L-Imino acids L-Pipecolic acid L-Proline L-Hydroxyproline

Substrate

Relative Activity

Sulfur containing (100) 23 0

amino acids L-Methionine L-Cysteine

0 0

D-Amino/Imino acids

Neutral amino acids L-Alanine

0

L-Serine L-Isoleucine

0 0

Glycine

0

Acidic amino acids L-Glutamic acid

0

Basic amino acids L-Lysine

2

D-Pipecolic acid D-Proline

0

D-Lysine

0

D-Aspartic acid D-Alanine

1 0

Other compounds Sarcosine Dimethylglycine

6

10 0

L-Histidine

0

Nipecotic acid

0

L-Glutamine

0

Glycolytic acid

0

L-Arginine Aromatic amino acids

0

L-Phenylalanine

0

On SDS-polyacrylamide gels, L-pipecolic acid oxidase had a positive periodic acid-Schiff (PAS) stain.

Carbohydrate

analysis, using the Dionex HPLC carbohydrate system, showed that the protein did not contain any of the common glycoprotein moieties.

However, the Dionex system will not

separate ribose, the carbohydrate moiety of flavin.

The lack

of other carbohydrates is not surprising since, to date, no

884 peroxisomal glycoproteins have been found.

We have concluded

that the positive PAS stain was probably associated with the ribose of the covalently bound

flavin.

The only other covalent flavoproteins thus far identified eukaryotes include succinate dehydrogenase, monoamine

oxidase, and L -

sarcosine dehydrogenase, dimethylglycine dehydrogenase, gulono-T-lactone oxidase mitochondria.

in

(7), all of which are located

in

To our knowledge, L-pipecolic acid oxidase

is

the first peroxisomal protein to contain a covalently-bound flavin and the first eukaryotic covalent flavoprotein outside the mitochondrion.

located

(Supported by NIH grants no.

HD10981, HD24061, and RR05808.)

References

1. Grove, J. and L. M. Henderson. 1968. Biochim. Biophys. Acta 165. 113-120. 2. Rao, V. V. and Y. F. Chang. 1990. Biochim. Biophys. Acta 1038. 295-299. 3. Mihalik, S. J. and W. J. Rhead. 1989. J. Biol. Chem. 2509-2517. 4. Mihalik, S. J. and W. J. Rhead. 1990. J. Compar. (in press).

264.

Physiol,

5. Mihalik, S. J., H. Moser, P. A. Watkins, A. Poulos, D. M. Danks and W. J. Rhead. 1989. Pediatr. Res. 25, 548-552. 6. Lazarow, P.B. and H. W. Moser. 1989. In: The Metabolic Basis of Inherited Disease, sixth edition (C. R. Schriver, A. L. Beaudet, W. S. Sly and D. Valle., eds.) McGraw Hill, New York, pp 1479-1509. 7. Edmondson, D. E. and R. DeFrancesco. 1990. In: Chemistry and Biochemistry of Flavoenzymes, Vol.1. CRC Press, Boca Raton, Florida, (in press)

DIFFERENCES

IN

PROPERTIES

OF

XANTHINE

DEHYDROGENASE

AND

XANTHINE OXIDASE

Takeshi Nishino Department

of

Biochemistry,

Yokohama

Fukuura 3-9, K a n a z a w a - k u , Yokohama

City

University

School

of

Medicine,

236, J a p a n

Introduction Xanthine

dehydrogenase

(EC

1.1.1.

204)

catalyzes

the

oxidation

h y p o x a n t h i n e t o x a n t h i n e o r x a n t h i n e to uric acid a t t h e e x p e n s e of of

NAD

to

NADH.

Mammalian

to the oxidase type

enzymes

including

milk e n z y m e

by proteolysis or by oxidation

of

reduction

convert

easily

of s u l f h y d r y l r e s i d u e ( s )

of

e n z y m e molecule (1,2) . The oxidase t y p e e x h i b i t s low x a n t h i n e :NAD r e d u c t a s e activity

but high x a n t h i n e

:0.-, r e d u c t a s e a c t i v i t y even in t h e p r e s e n c e of NAD

(3,4). The o x i d a s e t y p e p r o d u c e s hydrogen peroxide and s u p e r o x i d e (O2 ), while the

dehydrogenase

only

in

the

type

absence

produces

of

NAD

(5).

for cells, the 02"dependent

activity

tissue

in

injury

(G). H o w e v e r ,

a

considerable As t h e s e

amount

active

of

oxygen

these

are

is t h o u g h t t o be a c a u s e of post

contrast

to

mammalian

toxic

ischemic

e n z y m e s avian

a r e always of t h e d e h y d r o g e n a s e t y p e and a r e n e v e r c o n v e r t e d t y p e by any t r e a t m e n t

compound

species

enzymes

to the

oxidase

which has e v e r been a t t e m p t e d . As reviewed

previously

(3,4), t h e s e e n z y m e s a r e very similar in many of t h e i r physical and

enzymatic

properties;

they

are

dimeric

150,000 and c o n t a i n s one as

cofactors.

structure

and

This

chapter

functional

in o r d e r t o u n d e r s t a n d t o oxidase. In addition,

proteins

and

each

subunit

has

molecular

FAD, t w o iron s u l f u r c e n t e r s , and one deals

properties

with of

comparative xanthine

informations

dehydrogenase

t h e mechanism of conversion of x a n t h i n e

weight

molybdopterin on

and

protein oxidase,

dehydrogenase

t h e role of this e n z y m e in post ischemic tissue injury

is also b r i e f l y discussed.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter& Co., Berlin • New York - Printed in Germany

886 Structural Changes of Rat Liver Enzyme by Proteolysis or Sulfhydryl Oxidations Conversion

of

the

dehydrogenase

to

the

oxidase

occurs

reversibly

by

oxidation of sulfhydryl groups or irreversibly by proteolysis of enzyme molecule (1,2).

Although

the

dehydrogenase

type

can

be

easily

converted

irreversible oxidase type during purification, it can be purified as interconvertible

type

by

careful

purification

procedures

(7-9).

to

the

a reversible However,

has not yet been identified what kind of protease converts to the

it

irreversible

oxidase type during purification. Tryptic digestion can also result in conversion from the dehydrogenase type this

trypsin

treatment

to the irreversible oxidase type (2,7,10). Although

cleaved

the

enzyme

protein

without

dissociation

of

peptide fragments with concomitant change of the dehydrogenase type to the oxidase type, carboxymethylation the trypsin-treated and 85kD

in the presence of Guanidine-HCl

dissociated

enzyme into the t h r e e peptide fragments of 20kD, 40 kD

(10). Comparing the N-terminal sequence of each

fragment to

the

whole protein sequence obtained from cDNA, the cleavage sites were identified to be between 184-185 and 539-540 residues. As described in the o t h e r chapter in this volume (11), cysteine responsible

for

reversible

conversion

were

modified

with

residues

FDNB.

reversible conversion from the dehydrogenase type to the oxidase type,

During three

cysteine residues, Cys-525, Cys-980 and Cys-1313, were modified. It is possible t h a t two of these cysteine residues a r e concerned in the reversible conversion to form disulfide bridge.

Fig. 1. The tryptic cleavage sites and the modified sulfhydryl residues during conversion of rat liver xanthine dehydrogenase to oxidase.

887 Although and

which

it is still u n c l e a r t h a t the two c l e a v a g e s i t e s a r e both

cysteine

dehydrogenase by

alternation

the

enzyme

residues

are

concerned

in

the

conversion

type t o t h e oxidase type, t h e process is presumably of

the

positioning

of

the

resulting

in

conformational

molecule,

three

tryptic

peptide

changes

essential from

the

accompanied fragments

of

particularly

in

environments around t h e flavin and NAD.

D i f f e r e n c e in environments of FAD moiety Pronounced were

differences

indicated

reconstituted flavins

6-

by

with

or

displayed

slight

preference On

binding

the

of

the

the

no

residue(s)

an

intact

the of

-OH

2(A)

forms more

or - S H

or

flavin

of than

charged

N( l ) - p o s i t i o n , be

resulted rat

liver

replaced

by

either

xanthine

properties

bound of

substituents

The

xanthine

anionic (6-OH,

flavin 6-SH,

4

four

flavins,

pH

units.

site,

and

at

the

oxidase

(8-OH) and

or

possibly

preference

perturbing These

strong

flavin enzyme

imply

negative

for

the

pK that

charge

change its conformation

the NAD-binding s i t e was lost a t t h e same time as

residue(s) instead

dehydrogenase-oxidase

of

the forms

had a d r a m a t i c

all

NAD-binding

two

(12,13).

neutral the

around

the

l o c a t e d near the flavins N( 1 (-position, could

negatively

such

Fig.

dehydrogenase

substituents

with

in such a manner t h a t

the

for

the

of

ionizable in

form

(protonated)

ionizable

dehydrogenase,

hand,

structure

properties

shown

anionic

other

protein

containing as

preference

for

neutral

in

spectral

flavin

8-position

either 8-SH).

the

was

removed Such

digestion

dehydrogenase.

between

the

vicinity

by a partial positive c h a r g e

conversion.

tryptic

from

two types

a

should

of enzyme

be were

of

noted

change

cysteinyl that

the

flavin

concomitantly

conformational

or modification It

of

with might

residue(s)

differences

observed between

in

chicken

liver xanthine dehydrogenase and milk xanthine oxidase ( 1 2 ) , and also observed between

rat

liver

xanthine

as shown in Fig. 2 (B)

dehydrogenase

and

oxidase

in a

reversible

fashion

(13).

Redox p o t e n t i a l s of Chromophores Another

important

difference

between

the

two forms

is in the order

of

Rat Liver Xanthine Dehydrogenase 2 M CaCl2 Sephadex G-25 FAD Deflavo-enzyme

*

DTT Treatment I Sephadex G-25 Deflavo-enzyme

F A D H 7 F A D H 2 > F A D / F A D H " # F e / S ( I ) » Mo V / M o l V > MoVI/MoV, whereas

for

chicken

liver

xanthine

dehydrogenase

(19)

or

for

rat

liver

dehydrogenase ( 1 8 ) is F e / S ( I I ) > F A D / F A D H " = * F e / S ( I ) $ FADH'/FADH The

stabilization

noted

by

of

many

responsible

for

(3,4,14,15).

It

the

semiquinone

workers

(14,15,20)

the

low

should

be

this

of

that

S*Mo V / M o I V > M o V I / M o V .

xanthine

and

reactivity noted

in

properties

xanthine

raising

dehydrogenase was

has

suggested

dehydrogenase

the

flavin

been to

with

potential

by

be

oxygen replacing

t h e normal flavin with a r t i f i c i a l flavin having a higher p o t e n t i a l such as

2-thio

or 4 - t h i o

FAD does not inhibit stabilization of the semiquinone ( 2 1 ) , suggesting

that

stabilization

such

interaction be

also

due

to

flavin-protein

with t h e o t h e r oxidation-reduction

noted

that

FADH'/FADH2

are

consequently considered

is

the

largely

stability to

redox of

play

potentials

changed the

major

of

by in

and

not

due

to

c e n t e r s of t h e enzyme. I t should both

binding

semiquinone roles

interactions couples of

is

pyridine

changed

defining

of

the

FAD/FADH"

and

nucleotide

and

(18,19,22).

reactivity

of

This

is

xanthine

dehydrogenase.

Steady S t a t e The is

Kinetics

most

that

both

whereas

they

difference

is

compared

with

kinetic

characteristic NADH react

the the

parameters

difference

and

NAD

poorly

or

are not

poor r e a c t i v i t y high of

reactivity the

oxidase

good at

of

between

all

the

the

substrates with

the

two for

and

the

oxidase.

dehydrogenase

of the oxidase. T a b l e type

forms

with

of

enzyme

dehydrogenase, Second

apparent

molecular

oxygen

1 shows steady

dehydrogenase

type

of

state

purified

rat liver enzyme ( 1 8 ) . In this experiments Lineweaver-Burk plots shows parallel lines f o r

a series

of

fixed

concentrations

of t h e o t h e r s u b s t r a t e

in all

cases.

It is n o t i c e a b l e that the V value of x a n t h i n e - 0 „ a c t i v i t y c a t a l y z e d max ¿* oxidase type enzyme and t h a t of xanthine-NAD a c t i v i t y c a t a l y z e d by

by

890 dehydrogenase activity

type

catalyzed

these values.

enzyme

are

close,

by dehydrogenase

The K ^

while

type

the V

enzyme

value of xanthine-0„ max £> is less than one third of

value for O ^ of dehydrogenase type enzyme is about

five times as high as that of oxidase type enzyme. T A B L E 1. Steady state kinetic parameters of rat liver enzyme. Dehydrogenase

Oxidase

Xanthine-NAD activity K m for xanthine (jiM)

1.3

K m for N A D (jjM)

8.5

(mol/min/mol F A D )

810

Xanthine-Og activity K m for xanthine (yjM) K m for 0 2 V

max

Rapid

2.8

(>iM)

(mol/min/mol

Reaction

FAD)

Studies

of

Fully

1.8

260

46

270

1030

Reduced

Xanthine

Dehydrogenase

with

Molecular Oxygen When

a

dehydrogenase appearance process

of

with

observation the rates

reoxidation by

oxygen

the

process was

absorbance

various

(5). The was difficult

kinetic

of

fully

reduced

followed

with

spectrum

of

phases

being

a

chicken

stopped

oxidized

flow

enzyme

dependent

on

the

liver

xanthine

apparatus, are

wavelength

production process is also multiphasic and each to fit to the

reoxidation rate of the enzyme

the slowest one. The apparent discrepancies in rates at different in the reoxidation kinetics and between these and the O ^

the

multiphasic of of

except

wavelengths

production kinetics

can be easily reconciled by application of the principles of rapid equilibration of

the

reducing

equivalents

among

the

various

redox

groups

of

the

enzyme

in the same way as first proposed for xanthine oxidase (23). Fig. 3.

shows

the major enzyme forms which might be expected to occur during reoxidation, based on the observed

kinetics and our

knowledge

of the various redox groups of the enzyme.

of the relative

potentials

891

r-Tt/Srti

rMoVI WAD l-F./SrH

rF«/Sr«d

o2 V

H2O2 f

1

0 F«/Sred I2

r-Ft/Srtd

E

-Mo IV -FAD L F»/Sr«d

L

m.vi F*/Sox FADH-

v

o2 t rF«/Sr«d •MoVI FAD y -Ft/So:

pi•Fe/Sox (-MoVI rFADH» Ff/Sox

6e"

4e"

3e~

l-Fi/Sex •MoVI FAD Ff/Sox

k.

le"

2e"

Fig. 3. Reaction of fully reduced xanthine dehydrogenase with O (ref.5). The enzyme species in each vertical column are envisaged t o be in rapid equilibrium with the proportion of each species determined by the relative redox potentials of the d i f f e r e n t centers (ref. 23). It is envisaged that enzyme forms containing FADH react with O to g e n e r a t e H O with an instric r a t e constant k j , and those forms containing FADH' react t o produce O 2 with an instric r a t e constant k„.

Xanthine-Oxygen Turnover by Xanthine Dehydrogenase The major d i f f e r e n c e between xanthine dehydrogenase and xanthine oxidase is t h a t the

the

xanthine

catalyzes V

max

the

va ue

'

dehydrogenase oxidase

does

oxidation xan

of

has a binding site not.

thine-oxygen

xanthine

dehydrogenase

activity.

The

reaction

is of

In

xanthine 30-40

the

absence

to uric

reductase % of

f o r NAD and of

NAD, the

with

activity

that

reduced enzyme

acid

for the

NADH,

whereas

dehydrogenase

O^ as acceptor.

of chicken or rat xanthine-NAD

The liver

reductase

with oxygen is a relatively

slow

892 and

second

the

order

catalytic

by

process,

rate

xanthine

on

resulting

oxygen

dehydrogenase

considerable

(5,18)

and substantial

This

to

associated

containing

the

be flavin

between

that

expected

might

4-electron

be

reduced

expected

to

steps

one

of

is competition of

NAD

enzyme

with with

be

a

to

of

reduced

enzyme

(19)

and

C>2 t o the reduced f l a v i n of

to

be

lower

than

that

to

the

C>2 with

is

turnover

catalysis

as

major

transfer

form

0 2 . In

f o r the

much

the

faster

than

the

inhibitor of

reaction

(5).

flavin

of

xanthine

Urate Fe/S r i i L H o VI FADHi Fe/S r t i

rapid

^

At

oxidase

the

might

NAD

be

there

reaction

of of

reduced substrate

Accessibility

xanthine dehydrogenase, that might be reduced

in

levels

sequential

of

indicative

the O 2 reaction

forms

shown

(5). The

p r e v e n t e d by pyridine n u c l e o t i d e . It should be noted that binding of

k Ol I

is

form

presence

kinetics

catalysis.

reduced

through

reduced f l a v i n

saturation

turnover

forms.

(B)

oxygen

of

semiquinone

enzyme

reduced and 2 - e l e c t r o n

to

to

flavin

occured during of

dehydrogenase,

shows

binding so that N A D is a potent

O2

the

Xanthine-oxygen

during

and O^

of

of

of

reaction

dependence

xanthine-oxygen

amount

that

transfer NAD

During

strong

xanthine

form

between

O2

(5).

occur

levels

electron

the

the 4 - e l e c t r o n

dominant

observed

formation

with

semiquinone

Fig. 4 shuttling

the

concentration.

a

was observed appears

in

expected

(18),

is

also

NAD

— F e / S ,t( D o IV • Urate FAD —Fe/S rti Xanthine . _ Fe/S r i d C Ho VI _ FAD - Fe/S r t d

NADU + H' k, P - Fe/S , t i fx „ , ^ Ho VI FADH Fe/Srii j rapid

—Fe/S rii D o V FADH • -Fe/S rti

ri:V _ FAD - Fe/S

Fig. 4. S t e a d y s t a t e turnover of xanthine substrate and O^ as a c c e p t o r ( r e f . 5 ) .

r!i

dehydrogenase

with

xanthine

as

893 changes

the

4-electron

redox

potentials

reduced

catalysis (Fig.

level

of

flavin

might

be

couples

expected

Reperfusion

in early

injury,

hypoxic stage

oxyradical

tissue,

the reoxygenation

This

by

catabolism

proposed anoxia.

by

that

occurs

(2,7,8),

hypothesis

of

the

to

strong

of

the

even

inhibitor

cells,

of

pathogenic

reaction

might

of

is

after

It

short

a

process

(6) t h a t

the

responsible

for

superoxide

be

(5).

well

as

of

oxidized

by

conditions,

the

of

in

the

it

is

than

(30),

It

is

well

in

vivo

oxidase

amount

substrates

the

might NADH

(29).

As of

form,

in

tissue

xanthine

absence

of

NAD

reaction

possible

most

the injury

form

to

in normal

that

conditions

minor

a

strongly

hypoxanthine,

anoxic

in

(27,28).

Although

reported

dehydrogenase

from

(27,28). O^

during

reoxygenation

oxidase

conditions

been

hypothesis

xanthine

inhibits

exists occurs

oxidase

conversion

the

20-30%

of

of

the

to

dehydrogenase

However,

ischemic as

the

contain

amounts

enzyme

From

ischemia a

roles

nucleotide

time

normal

of as

the

accumulation

converts

that

important

has

xanthine

whereas

to

the

less

NADH,

ischemia.

time

seems Thus,

that

to explain

mostly

oxygen

ischemia.

of

slow

short

after

reports

suggest

pyridine

with C^

during

exists

be

the

and hypoxanthine

dehydrogenase

significant

the

enzyme

during

hypoxanthine

at

during

reoxygenation

important

oxidase

on

(26)

is t o o

organ

to

accumulation

occur

based

reports

type.

produces

larger amount of

dramatically

in

reversible

presence

because

following

most

xanthine

relatively

normal

suggested

reduced

the

nucleotide

enzyme

injury

a

occur

of

xanthine

oxidase

a

the

a

the

form

or organ inplantation. Hypothesis

also

some

to

although

the

is

(6)

presumably

(5),

one

from

adenine

after

dehydrogenase

major

form

Injury

may

cells of many organs

that

is

the

(A)

and allopurinol diminish reoxygenation injury in some system

However,

established

be

that

injury has been supported by t h e observation t h a t

McCord

dehydrogenase

is

infarct

derived

dismutase, c a t a l a s e (24,25).

which

of organ

species

in endotherial

to

so

4).

Role of X a n t h i n e Oxidase in Recirculation

previously

(19,22)

of

produce

conditions,

be

expected

accumulates NADH

is

a

accumulated

of

oxidase

contained

are

oxidized

xanthine

dehydrogenase to form NADH . However, f u r t h e r detailed studies on this a s p e c t a r e needed using adequate model systems.

894 Acknowledgements The author thanks Drs. K. Tsushima, Y. Amaya, Tomoko Nishino, T. Saito, K. Yamazaki, Yokohama City University, for helpful discussions. He also wishes to thank Drs. V. Massey and L.M. Schopfer, The University of Michigan, for their helpful discussions. This work was supported by Grantin-Aid 62480135, 2044123 for scientific research from the Japanese Ministry of Education, Science and Culture, and in part by a research grant for intractable disease from the Japanese Ministry of Health and Welfare.

References 1. Deila Corte, E., Stirpe, F. 1968. Biochem. J. K)8, 349 2. Delia Corte, E., Stirpe, F. 1972. Biochem. J. 126, 739 3. Coughlan, M.P. 1980. in Molybdenum and Molybdenum-containing Enzymes (Coughlan, M.P. ed), ppl 19, Pergamon Press, Oxford 4. Bray, R.C. 1981. in Flavins and Flavoproteins (Massey, V., Williams, C.H. Jr., eds) pp775, Elsevier, New York 5. Nishino, T., Nishino, T., Schopfer., L.M., Massey, V. 1989. J. Biol. Chem. 257, 2518 6. McCord, J.M. 1985. N. Engl. J. Med. 312, 159 7. Waud, W.R., Rajagopalan, K.V. 1976. Arch. Biochem. Biophys. 172, 354 8. Nakamura, M., Yamazaki, I. 1982. J. Biochem. (Tokyo) 92, 1279 9. Ikegami, T., Nishino, T. 1986. Arch. Biochem. Biophys. 247, 254 10. Amaya, Y., Yamazaki, K., Sato, M., Nöda, K., Nishino, T., Nishino, T. 1990. J. Biol. Chem. in press 11. Nishino, T., Amaya, Y., Nöda, K., Nishino, T. in Flavins and Flavoproteins, this volume 12. Massey, V., Schopfer, L.M., Nishino, T., Nishino, T. 1989. J. Biol. Chem 264, 10567 13. Saito, T., Nishino, T., Massey, V. 1989. J. Biol. Chem. 264, 15930 14. Barber, M.J., Bray, R.C., Lowe, D.J., Coughlan, M.P. 1977. Biochem. J. 153, 297 15. Barber, M.J., Bray, R.C., Cammack, R., Coughlan, M.P. 1977. Biochem. J. 163, 279 16. Spence, J.T., Barber, M.J., Siegel, L.M. 1982. Biochemistry 21, 1656 17. Porras, A.C., Palmer, G. 1982. J. Biol. Chem. 257, 11617 18. Saito, T., Nishino, T. 1989. J. Biol. Chem. 264, 10015 19. Schopfer, L.M., Massey, V., Nishino, T. 1988. J. Biol. Chem. 263, 13528 20. Rajagopalan, K.V., Handler, P. 1967. J. Biol. Chem. 242, 4097 21. Nishino, T., Nishino, T., Schopfer, L.M., Massey, V. 1989. J. Biol. Chem. 264, 6075 22. Nishino, T., Nishino, T. 1989. J. Biol. Chem. 264, 5468 23. Olson, J.S., Ballou, D.P., Palmer, G., Massey, V. 1974. J. Biol. Chem. 249, 4363 24. Shlafer, M., Kane, P.F., Wiggins, V.Y., Kirsh, M.M. 1982. Circulation 66, 185 25. Parks, D.A., Granger, D.N. 1983. Am. J. Physiol. 245, G 285 26. Jarasch, E.D., Grund, C., Bruder, G., Heid, H.W., Keenan, T.W., Franke, W.W. 1981. Cell, 25, 67 27. Groot, H., Littauer, A. 1988. Biochem. Biophys. Res. Commun. .155, 278 28. Wajner, M., Harkness, R.A. 1989. Biochim. Biophys. Acta 991, 79 29. Brosnan, J.T., Krebs, H.A., Williamson, D.H. 1970. Biochem. J. H 7 , 91 30. Kaminski, Z.W., Jezewska, M.M. 1979. Biochem. J. 181, 177

BIOCHEMICAL CHARACTERIZATION OF A MUTANT HUMAN MEDIUM-CHAIN ACYL-COA DEHYDROGENASE PRESENT IN PATIENTS HAVING DEFICIENT ACTIVITY.

Peter Bross, Franz Krautle, Jochen Stiemke, Sandro Ghisla and Ihab Rasched Faculty of Biology, Univ. Konstanz, D-77 50 Konstanz, FRG Niels Gregersen, Brage S. Andresen Molek. Genet. Lab., Skejby Universitetshospital, DK-8200 Arhus Arnold Strauss, Daniel P. Kelly Washington Univ. Med. Center, St. Louis, MO 63110, USA

Introduction Medium-chain acyl-CoA dehydrogenase (MCAD) deficiency is a relatively common inherited metabolic defect causing liver dysfunction and hypoglycaemic coma and accounting for some of the cases of sudden death in infants [1,2]. The mutation responsible for MCAD deficiency in five families is a point mutation causing a lysine to glutamic acid change at position 304 of the mature protein. According to the three-dimensional structure, Lys-304 is not part of the active site but is part of the helix which forms the interface between the subunits [3]. In this report we demonstrate expression of mutant MCAD-glu-304 in E.coli and biochemical characterization of the defective enzyme in comparison with wild-type MCAD.

Flavins and Flavoproteins 1990 © 1991 by Walter de Gruyter & Co., Berlin • New York - Printed in Germany

896 Results Construction o f p l a s m i d s : We h a v e d e s c r i b e d the c o n s t r u c t i o n o f a p l a s m i d (pWTMCAD-2), which d i r e c t s e x p r e s s i o n o f m a t u r e human MCAD i n E.coli [4] . A d e r i v a t i v e of pWTMCAD-2, pBMCK2-, which expresses mature a c t i v e human MCAD under c o n t r o l of the l a c promoter, was constructed ( f i g . l ) . The EcoRI/BamHI fragment of pBMCK2" c a r r y i n g the i n f o r m a t i o n f o r the C-terminal h a l f of MCAD was replaced with an EcoRI/BamHI fragment o r i g i n a t i n g from a PCR clone with the glu-304 mutation ( f i g . l ) . The plasmid encompassing the sequence of MCAD-glu-304 was c a l l e d

Figure

1: Maps of MCAD wild - type

plasmids. orientation transition acid position

Relevant of

restriction

the genes

causing 304 of

a lys

and mutant sites,

location

and the position to glu

codon

the mature protein

expression

of change

are

and

the A to G at

amino

indicated.

897 p985-41. In order to ensure that no PCR errors are contained, the whole EcoRI/BamHI fragment was sequenced and found to be identical to the wild-type sequence with the exception of the lys to glu codon change.

Expression of wild-type and alu-304 mutant MCAD in E.coli: Both plasmids, pBMCK2- (wild-type) and p985-41 (glu-304 mutant), express correctly-sized, immunoreactive MCAD protein (fig.2). In comparison between wild-type and lys304, cells disrupted by boiling in electrophoresis sample buffer display additional and more pronounced degradation bands for the mutant. In extracts prepared by lysozyme disruption of the cells, the band for the mutant protein is much weaker than that of the wild-type. This indicates that after disruption the mutant protein is more susceptible to proteolytic degradation than the wild-type.

Activity in extracts: Extracts of cells expressing, wildtype or glu-304 MCAD respectively were assayed for enzyme activity. Extracts were prepared by lysozyme disruption followed by NH^-sulfate precipitation (20-70%) and activity was measured as described by Thorpe et al (5) In this assay, extracts from cells expressing wild-type MCAD display activities in the range of 20-25 mu/mg soluble protein. No activity could be detected in extracts from cells expressing MCAD-glu-304. Due to the high background in the extract the assay would be sensitive enough to detect >5% of the activity measured for the wild-type extract.

898

0