Flavins and Flavoproteins 1993: Proceedings of the Eleventh International Symposium, Nagoya, Japan, July 27–31, 1993 [Reprint 2020 ed.] 9783110885774, 9783110141658

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Flavins and Flavoproteins 1993

Flavins and Flavoproteins 1993 Proceedings of the Eleventh International Symposium Nagoya (Japan) July 27-31,1993

Editor Kunio Yagi

W G_ DE

Walter de Gruyter • Berlin • New York 1994

Editor Kunio Yagi, M.D. Ph. Director Institute of Applied Biochemistry Yagi Memorial Park Mitake, Gifu 505-01 Japan With 392 figures and 91 tables

© Printed on acid free paper which falls within t h e guidelines of the A N S I to ensure permanence and durability.

Library of Congress Cataloging-in-Publication

Data

Flavins and flavoproteins 1993 : proceedings of the eleventh international symposium, Nagoya (Japan) July 27-31,1993 / editor, Kunio Yagi Includes bibliographical references and index. I S B N 3-11-014165-5 1. Flavins—Congresses. 2. Flavoproteins—Congresses. I. Yagi, Kunio. II. International Symposium on Flavins and Flavoproteins (11th : 1993 : Nagoya, Japan) QP671.F5F54 1994 94-26914 574.19'258~dc20 CIP

Die Deutsche Bibliothek -

CIP-Einheitsaufnähme

Flavins and flavoproteins: Proceedings of t h e international symposium. Berlin ; New York : de Gruyter. 1993. Proceedings of the eleventh international symposium, Nagoya, Japan, July 27-31,1993. - 1994 I S B N 3-11-014165-5

© Copyright 1994 by Walter de Gruyter & Co., D-10785 Berlin. All rights reserved, including those of translation into foreign languages. N o part of this book may be reproduced in any f o r m or by any means, electronic or mechanical, including photocopy, recording,or any information storage and retrieval system, without permission in writing from the publisher. Printed in Germany. Printing: Druckerei Gerike G m b H , Berlin. Binding: Mikolai, Berlin.

PREFACE

This volume is a collection of the papers presented at the Eleventh International Symposium on Flavins and Flavoproteins held in Nagoya, Japan, from the 27th to the 31st of July, 1993. As I mentioned in the special lecture at the last symposium held in Como (cf. Ravins and Flavoproteins 1990, pp. 3-16), our symposia are unique in that these meetings have been held almost regularly for nearly 30 years. If we compare this period with the total age of flavin research, viz., about 60 years, this is surprisingly long. This longevity is obviously due to the international cooperation among the scientists of different disciplines who have taken an interest in flavins and flavoproteins, which substances are of fundamental importance for our life. The present symposium focused our attention on the structure-function relationship of our flavoproteins. Different kinds of approaches through physical, chemical, biological and genetical studies were presented, representing the skeleton of the present symposium; and the papers presented could be classified into those on 1) theoretical and chemical approaches, 2) biosynthesis of flavins, 3) flavoproteins containing dissociable or covalently bound flavin as the sole catalytic center, 4) complex flavoproteins containing flavin and other prosthetic group(s) for catalytic activity, and 5) enzymes using reduced flavin for their activity. Although recent advances since the last symposium are noteworthy, we realize that many things still remain to be clarified. In addition, recent developments in molecular biology now allow us to produce possible mutants of proteins, which will enable us to investigate further the structure-function relationship of our enzymes and probably to construct more idealistic flavoproteins. Thus many problems await our solution. Naturally we cannot predict when the research in this field will terminate. Accordingly, we must expect further advances to be reported in the next symposium that will be held in Calgary in 1996.

VI Another unique feature of our symposia is the friendly and constructive atmosphere generated by the scientists who gather for our meetings. This is a most important attribute, since truly international cooperation is the best way to advance science and thus promote human welfare. Finally, I would like to thank most sincerely all of the members of the international advisory committee as well as those of the regional organizing committee for their help and advice regarding this symposium, the International Union of Biochemistry and Molecular Biology and other organizations for their financial support, and all of the participants for their cooperation in having made this meeting fruitful in the sense of both science and culture. Kunio Yagi Gifu, November, 1993

INTERNATIONAL ADVISORY COMMITTEE: Bruno Curti (Milano) Vincent Massey (Ann Arbor) Masateru Shin (Kobe) Cees Veeger (Wageningen) Kunio Yagi (Gifu) REGIONAL ORGANIZING COMMITTEE: Kenji Aki, Yoshiyuki Ichikawa, Eiji Itagaki, Takashi Iyanagi, Hiroshi Matsubara, Retsu

Miura,

Yoshihiro

Miyake,

Takao

Nakamura,

Takeshi

Nishino,

Yukio Nisimoto, Nobuko Ohishi, Kiyoshi Shiga, Masateru Shin, Kenji Soda, M a s a z u m i Takeshita, Haruhito Tsuge, Keishiro W a d a , K u n i o Yagi, Toshitsugu Yubisui SPONSORS: The International Union of Biochemistry and Molecular Biology Aichi Prefecture Nagoya City Institute of Applied Biochemistry CONTRIBUTORS: Eisai Co., Ltd. Daiichi Pharmaceutical Co., Ltd. Kyowa Hakko Kogyo Co., Ltd. Mochida Memorial Foundation for Medical and Pharmaceutical Research Mochida Pharmaceutical Co., Ltd. Pfizer Pharmaceuticals Inc. Sato Pharmaceutical Co., Ltd. Suzuken Memorial Foundation Tokyo Tanabe Co., Ltd. Tosoh Corporation

and

CONTENTS

THEORETICAL AND CHEMICAL APPROACH Use of molecular orbital calculations in studies on mechanisms of enzyme catalysis I.M.C.M. Rietjens, M.G. Boersma, A.E.M.F. Soffers, N.H.P. Cnubben, J. Koerts, S. Peelen, W.J.H. van Berkel, C. Veeger, and J. Vervoort

3

A theoretical approach of the mechanism of C(4a)-peroxyflavin catalyzed reactions C. Veeger, I.M.C.M. Rietjens, J. Vervoort, and J. Lee

13

Semiempirical calculations on properties of (iso)alloxazines in ground and excited states H. Szymusiak

23

Pathways for a 4 a -hydroxy-4 a ,5-dihydroflavin radical in the hydroxylation of aromatics H.I.X. Mager, and S.-C. Tu

27

Chemical of nekoflavin K. Matsui,synthesis and S. Kasai

31

Novel 5-substituted 5-deazaflavins: Synthesis and applications as active site probes for flavoproteins Y.V.S.N. Murthy, and V. Massey

35

D-Lactate dehydrogenase model. Oxidation of a-hydroxy acid by functionalized oxidation active flavin mimic in the presence of Zn 2 + and base in t-butanol Y. Yano, K. Mitsui, Y. Ohsawa, and T. Nabeshima

39

Mechanism of flavin reduction by Cu(I)-EDTA complex B. Lei, and S.-C. Tu

43

Self-association flavin: Interaction of FMN with albumin T. Ishida, and K.ofHoriike

47

BIOSYNTHESIS OF FLAVINS Studies on the biosynthesis of flavins. Structure and mechanism of 6,7dimethyl-8-ribityllumazine synthase A. Bacher, K. Ritsert, K. Kis, K. Schmidt-Base, R. Huber, R. Ladenstein, J. Scheuring, S. Weinkauf, and M. Cushman

53

X Biosynthesis of riboflavin. Cloning, sequencing, mapping, and hyperexpression of the genes ribA coding for GTP cyclohydrolase II and ribC coding for riboflavin synthase of Escherichia coli S. Eberhardt, G. Richter, H. Ritz, J. Brandt, and A. Bacher

63

Biosynthesis of riboflavin: Enzymatic formation of 6,7-dimethyl-8-ribityllumazine in Saccharomyces cerevisiae J J . García-Ramírez, M. A. Santos, and J.L. Revuelta

67

Electron microscopic studies on the lumazine synthase/riboflavin synthase complex of Bacillus subtilis S. Weinkauf, J. Brandt, and A. Bacher

71

19 F NMR studies on lumazine protein from Photobacterium phosphoreum J. Scheuring, J. Lee, M. Cushman, H. Oschkinat, and A. Bacher

75

NMR studies on the mechanism of riboflavin synthase J. Scheuring, M. Cushman, H. Oschkinat, and A. Bacher

79

OXIDASES Studies on active site mutants of spinach glycolate oxidase P. Macheroux, and Y. Lindqvist

85

High-level expression of rat kidney hydroxy acid oxidase in Escherichia coli: Purification and characterization of the recombinant protein A. Belmouden, and F. Lederer

95

The influence of substrate structure on the redox reactions of monoamine oxidases R.R. Ramsay, S.O. Sablin, and T.P. Singer

99

Quantitative structure-activity relationships in the oxidation of benzylamine analogues by bovine liver monoamine oxidase B M.C. Walker, and D.E. Edmondson

109

Experimental probes of hydrogen tunneling in bovine liver monoamine oxidase B T. Jonsson, J.P. Klinman, and D.E. Edmondson

117

Stereospecificity and deuterium isotope effect in the oxidative deamination catalyzed by flavine and non-flavine amine oxidases P.H. Yu, and B.A. Davis

123

Characterization of L-aspartate oxidaseG. overexpressed E. coli and H.G. Gassen A. Negri, M. Mortarino, T. Simonie, Tedeschi, S.inRonchi,

127

Identification of another subunit in L-phenylalanine oxidase from Pseudomonas sp. P-501 and the primary structure of the subunit E.B. Mukouyama, H. Suzuki, T. Sasaki, and H. Koyama

137

XI

Kinetic isotope effects on reductive half-reactions of flavoprotein oxidases P.F. Fitzpatrick, J.M. Denu, J.J. Emanuele, and V. Menon

141

Effects of ligands on the reactivities of reduced and semiquinoid forms of D-amino acid oxidase Y. Nishina, K. Sato, and K. Shiga

151

Thermodynamic study of FAD binding in D-amino acid oxidase F. Tanaka

155

Structure K. Fukui of human D-amino acid oxidase gene Amino acid sequence of D-amino acid oxidase from the yeast Rhodotorula gracilis L. Faotto, L. Pollegioni, F. Ceciliani, G. Gadda, S. Ronchi, and M.S. Pilone

159 163

Chemical modification of arginine groups in D-amino acid oxidase from Rhodotorula gracilis. Involvement in catalysis and assignment in the sequence G. Gadda, A. Negri, and M.S. Pilone

167

Functional and structural aspects of D-amino acid oxidase from Rhodotorula gracilis probed by limited proteolysis L. Pollegioni, F. Ceciliani, B. Curti, S. Ronchi, and M.S. Pilone

171

The crystal structure of cholesterol oxidase. Mechanistic implications for FAD dependent alcohol oxidation A. Vrielink, J. Li, P. Brick, and D. Blow

175

Nitroalkane-oxidizing K. Soda, T. Kurihara, flavoenzymes M. Tchorzewski, N. Esaki, and N. Ohishi

185

A 340-nm chromophore of nitroalkane oxidase from Fusarium oxysporum T. Kurihara, N. Esaki, K. Soda, and N. Ohishi

195

OXYGENASES Lactate monooxygenase: Studies of active site mutations U. Muh, C.H. Williams, Jr., and V. Massey

201

The mobile flavin of parahydroxybenzoate hydroxylase — A case for major structural dynamics in catalysis B. Entsch, B.A. Palfey, M.S. Lumberg, D.P. Ballou, and V. Massey

211

Structures of mutant p-hydroxybenzoate hydroxylases: Evidence for an alternative mode of flavin binding M.S. Lah, D. Gatti, H.A. Schreuder, B.A. Palfey, and M.L. Ludwig

221

XII Substrate and effector specificity of two active-site mutants of p-hydroxybenzoate hydroxylase from Pseudomonas fluorescens W.J.H. van Berkel, F.J.T. van der Bolt, M.H.M. Eppink, A. de Kok, I.M.C.M. Rietjens, C. Veeger, J. Vervoort, and H. Schreuder

231

Solvent isotope effects on p-hydroxybenzoate hydroxylase B.A. Palfey, D.P. Ballou, and V. Massey

235

Immunological studies on the structure of 4-aminobenzoate hydroxylase from Agaricus bisporus H. Tsuji, T. Oka, M. Kimoto, Y. Natori, T. Ogawa, and Y.-M. Hong

239

Studies on p-hydroxyphenylacetate-3-hydroxylase U. Arunachalam, and V. Massey

243

Fluorescent intermediates and the reaction mechanism of phenol hydroxylase K. Maeda-Yorita, and V. Massey

247

Application of pulse radiolysis to produce absorbing species of substrates for flavoprotein hydroxylases R.F. Anderson

251

Mammalian flavin-containing monooxygenase catalyzed conversion of 4-haloN-methylanilines J. Vervoort, M.G. Boersma, N.H.P. Cnubben, W.J.H. van Berkel, J. Koerts, J.A. Boeren, and I.M.C.M. Rietjens

255

2-Aminobenzoyl-CoA-monooxygenase/reductase, an enzyme with two distinct functions and one active center B. Langkau, and S. Ghisla

259

Characteristic properties and kinetic analysis of neurotoxins for porcine FADcontaining monooxygenase R.-F. Wu, and Y. Ichikawa

263

Genomic DNA structure of a unique flavoprotein, 2-nitropropane dioxygenase from Hansenula mrakii M. Tchorzewski, T. Kurihara, N. Esaki, and K. Soda

267

DEHYDROGENASES AND ELECTRON TRANSFER Three dimensional structures of acyl-CoA dehydrogenases: Structural basis of substrate specificity J.J.P. Kim, M. Wang, S. Djordjevic, R. Paschke, D.W. Bennett, and J. Vockley

273

Mechanism of a,|3-dehydrogenation by acyl-CoA dehydrogenases S. Ghisla, S. Engst, P. Vock, V. Kieweg, P. Bross, A. Nandy, I. Rasched, and A.W. Strauss

283

XIII 13 C- and 1 5 N-NMR studies of medium-chain acyl-CoA dehydrogenase from porcine kidney

R. Miura, Y. Nishina, K. Sato, S. Fujii, K. Kuroda, and K. Shiga

293

Resonance studies on acyl-CoA dehydrogenases Y. Nishina,Raman K. Sato, I. Hazekawa, S. Fujii, K. Kuroda, R. Miura, and K. Shiga

303

Two new mechanism-based inhibitors of the acyl-CoA dehydrogenases J.G. Cummings, and C. Thorpe

313

Studies of the thermodynamic regulation of medium chain acyl-CoA dehydrogenase B.D. Johnson, and M.T. Stankovich

323

Long-chain specific enzyme from medium-chain acyl-CoA dehydrogenase A. Nandy, P. Bross, F. Kräutle, I. Rasched, and S. Ghisla

327

Glutaryl-CoA dehydrogenase from anaerobic, benzoate degrading sp.: An FAD-dependent glutaconyl-CoA decarboxylase U. Härtel, and W. Buckel

331

Pseudomonas

Crystallization and preliminary x-ray diffraction study of the Afunctional flavoenzyme 5-hydroxyvaleryl-CoA dehydratase/dehydrogenase from Clostridium aminovalericum U. Eikmanns, W. Buckel, and E.F. Pai

335

] Essential arginine residue(s) K. of Suzuki, 3-ketosteroid-A -dehydrogenase M. Kadode, H. Matsushita, and E. Itagaki

339

Flavodoxins and nitrogen fixation — Structure, electrochemistry and posttranslational modification by coenzyme A R.N.F. Thorneley, G.A. Ashby, M.H. Drummond, R.R. Eady, D.L. Hughes, G. Ford, P.M. Harrison, A. Shaw, R.L. Robson, J. Kazlauskaite, and H.A.O. Hill

343

Regulation of the expression of flavodoxin and ferredoxin in Anabaena PCC 7120 under iron and nitrogen limiting conditions P. Razquin, M.L. Peleato, C. Gomez-Moreno, M.F. Fillat, S. Schmitz, and H. Böhme

355

Crystallization and x-ray structure determination of E. coli flavodoxin D.M. Hoover, R.G. Matthews, and M.L. Ludwig

359

Cis-trans isomerization of the 57-58 peptide in crystalline flavodoxins from C. beijerinckii M.L. Ludwig, M.M. Dixon, K.A. Pattridge, and R.P. Swenson

363

Electron-transfemng flavoprotein from pig kidney contains an AMP K. Sato, Y. Nishina, and K. Shiga

367

The enigma of Old Yellow Enzyme. II V. Massey

371

XIV Stirring new interest in an old enzyme: Crystal structure of old yellow enzyme K.M. Fox, and P.A. Karplus

381

Effect of deletion and mutation of Old Yellow Enzyme gene from Saccharomyces cerevisiae B.J. Brown, and V. Massey

391

Structure and function of NADH-cytochrome bs reductase in relation to hereditary methemoglobinemia T. Yubisui, K. Shirabe, and M. Takeshita

395

Role of flavin binding motif, RxY(T/S), of NADH-cytochrome b$ reductase in electron transfer reaction K. Shirabe, T. Yubisui, and M. Takeshita

405

The structure of human erythrocyte NADH-cytochrome bs reductase at 2.5A resolution T. Takano, S. Bando, C. Horii, M. Higashiyama, K. Ogawa, M. Sato, Y. Katsuya, M. Danno, T. Yubisui, K. Shirabe, and M. Takeshita

409

Two bound forms of ferredoxin-NADP reductase in chloroplast thylakoids M. Shin, T. Nishikawa, T. Sekido, and N. Sakihama

413

Characterization of active-site mutants of ferredoxin-NADP + reductase A. Aliverti, L. Piubelli, B. Curti, and G. Zanetti

423

Structural study of ferredoxin-NADP + reductase from Anabaena PCC 7119 and of its complex with NADP + L. Serre, F. Vellieux, J. Fontecilla-Camps, M. Frey, M. Medina, and C. Gomez-Moreno

431

Structure-function relationships in ferredoxin: Site-directed mutagenesis and time-resolved absorption spectroscopy applied to the ferredoxin:ferredoxin N A D P + reductase interaction J.K. Hurley, Z. Salamon, T.E. Meyer, J.C. Fitch, M.A. Cusanovich, G. Tollin, H. Cheng, B. Xia, Y.K. Chae, J.L. Markley, M. Medina, and C. Gomez-Moreno

435

N-Terminal structure of mature ferredoxin-NADP reductase N. Sakihama, M. Shin, and S. Obata

439

Refined crystal structures of native, complexed and reduced forms of spinach ferredoxin reductase C.M. Bruns, and P.A. Karplus

443

Overexpression of ferredoxin-NADP + reductase from Anabaena sp. PCC7119 in E.coli M.F. Fillat, M.C. Pacheco, M.L. Peleato, and C. Gomez-Moreno

447

XV The reductase activity of a flavoprotein is changed into a superoxide-forming oxidase by chemical modification M.T. Bes, M.L. Peleato, C. Gómez-Moreno, A. López, and V.M. Fernández

451

FNR-like flavoproteins of chemoautotrophic bacteria and photosynthetic bacteria Y. Fukumori, S. Hikichi, and T. Yamanaka

455

^ NMR investigation of NADPH-adrenoferredoxin reductase with NADP+ and adrenoferredoxin S. Miura, and Y. Ichikawa

459

One electron reduction of adrenodoxin reductase as studied by pulse radiolysis M. Miki, K. Kobayashi, S. Miura, and Y. Ichikawa

463

Structure-function studies on NADPH-cytochrome P450 reductase using urea-perturbation and 19 F NMR spectroscopy R. Narayanasami, P.M. Horowitz, B.S.S. Masters, and J.D. Otvos

467

Cerebellar nitric oxide synthase behaves as a bi-domain structure B.S.S. Masters, E. Sheta, and K. McMillan

471

Studies on NAD(P)H-quinone oxidoreductase G. Tedeschi, S. Chen, and V. Massey

475

Effect of riboflavin deficiency on DT-diaphorase K. Yagi, S. Komura, M. Nakashima, N. Ishida, N. Ohishi, and L. Emster DISULFIDE REDUCTASES

479

Hybrid molecules of glutathione reductase: Tools for investigating protein interactions at the dimer interface N.S. Scrutton, M.P. Deonarain, A. Berry, and R.N. Perham

485

The reductive half reaction of Escherichia coli glutathione reductase P. Rietveld, L.D. Arscott, and C.H. Williams, Jr.

493

Determination of the redox potential of E.coli glutathione reductase using NADH and NAD+ D.M. Veine, L.D. Arscott, and C.H. Williams, Jr.

497

An investigation of engineered co-operativity in interface mutants of Escherichia coli glutathione reductase A. Bashir, M.J. Cockerill, A. Berry, N.S. Scrutton, and R.N. Perham

501

Role of conserved glycine residues in the NADPH binding motif of glutathione reductase M. Rescigno, and R.N. Perham

505

XVI Eukaryotic lipoamide dehydrogenase: Molecular genetic and structural aspects K. Koike, M. Koike, and A. Takenaka

509

Disruption of the gene coding for dihydrolipoamide dehydrogenase in Haloferax volcanii by homologous recombination N.N. Vettakkorumakankav, K.J. Stevenson, L.C. Schalkwyk, and W.F. Doolittle

519

Redox properties of the FAD in the active site mutants C44S and C49S of Escherichia coli lipoamide dehydrogenase N. Hopkins, and C.H. Williams, Jr.

523

R- and S-dihydrolipoic acid derivatives as substrates of lipoamide dehydrogenase L.D. Arscott, and C.H. Williams, Jr.

527

Comparative study on flavin radical formation of lipoamide dehydrogenase and glutathione reductase by photoreduction K. Maeda-Yorita, and K. Aki

531

The pyruvate dehydrogenase complex from Azotobacter vinelandii A. de Kok, A. Berg, W. van Berkel, A. Fabisz-Kijowska, A. Westphal, F. van den Akker, A. Mattevi, and W.GJ. Hoi

535

The 3-dimensional structure of trypanothione reductase from Trypanosoma cruzi as a basis for drug design against Chagas' disease R.L. Krauth-Siegel, E. Jacoby, and C.B. Lantwin

545

Mechanism and structure of thioredoxin reductase T.S.R. Krishna, G. Waksman, J. Kuriyan, S.B. Mulrooney, B.W. Lennon, and C.H. Williams, Jr.

557

E. coli thioredoxin reductase — Potential acid-base catalysts — Properties of HIS-245 and ASP-139 mutants S.B. Mulrooney, A. Nicoletti, J. Yuvaniyama, and C.H. Williams, Jr.

567

Rapid reaction kinetics of Williams, EscherichiaJr.coli thioredoxin reductase B.W. Lennon, and C.H.

571

2'-Fluoro-2'-deoxy-arabino-FAD: Effects on the formation and stability of 2-electron reduced mercuric ion reductase S.M. Miller

575

The alkyl hydroperoxide reductase of Salmonella typhimurium: Redox activity of the cystine disulfide of AhpC and evidence for involvement in peroxide reduction L.B. Poole

583

Flavoprotein peroxide and disulfide reductases and their roles in streptococcal oxidative metabolism A. Claiborne, R.P. Ross, D. Ward, D. Parsonage, and E.J. Crane, III

587

XVII N^-(2-Aminoethyl)-FAD: Synthesis and coenzyme activity with respect to apo-NADH oxidase from Thermus thermophilics and Thermus aquaticus A.F. Biickmann, H. Erdmann, M. Pietzsch, J.M. Hall, and J.V. Bannister

597

A flavoprotein functional as NADH oxidase from Amphibacillus xylanus EpOl Y. Niimura, K. Ohnishi, K. Yokoyama, and T. Nishino

601

COMPLEX FLAVOPROTEINS Probing the structure and function of flavocytochrome ¿2 using protein engineering methods S.K. Chapman, P. White, S. Daff, G.A. Reid, R.E. Sharp, and F.D.C. Manson

607

The L-mandelate dehydrogenase from Rhodotorula graminis is a flavocytochrome ¿2 S.K. Chapman, O. Smekal, and G.A. Reid

617

S-Mandelate dehydrogenase from Pseudomonas putida: Construction of a soluble chimeric mutant of a membrane-bound enzyme B. Mitra, J.A. Gerlt, P.C. Babbitt, G.L. Kenyon, D. Joseph, and G.A. Petsko

621

The D282N and Y254L active-site mutants of flavocytochrome ¿2 are expressed in E. coli as a mixture of holoenzyme and flavin-free protein M. Gondry, M. Schaffner, F.D.C. Manson, S.K. Chapman, G.A. Reid, and F. Lederer

625

Reconstitution of flavin-free flavocytochrome bj with 5-deazaFMN: A carbanion or a hydride mechanism? A. Balme, and F. Lederer

629

Physical studies on phthalate dioxygenase reductase (PDR) D. Ballou, G. Gassner, L. Wang, C. Batie, D. Gatti, W.R. Dunham, and R.H. Sands

639

Structures of oxidized, reduced, and liganded states of the iron-sulfur flavoprotein, phthalate dioxygenase reductase C.C. Correll, D.L. Gatti, and M.L. Ludwig

649

Flavin and iron-sulfur cluster containing hydroxyacyl-CoA dehydratases (I) (7?)-2-Hydroxyglutaryl-CoA dehydratases from Acidaminococcus fermentans and Fusobacterium nucleatum U. Müller, A.-G. Klees, K. Bendrat, M. Mack, and W. Buckel

659

Flavin and iron-sulfur containing hydroxyacyl-CoA dehydratases (II) 4-Hydroxybutyryl-CoA dehydratase from Clostridium aminobutyricum U. Scherf, and W. Buckel

663

Glutamate synthase from Azospirillum brasilense: Structural and mechanistic studies M.A. Vanoni

667

XVIII Cloning and expression of Azospirillum brasilense glutamate synthase R. Pelanda, L. Piubelli, P. Fumagalli, A. Mazzoni, E. Verzotti, M. A. Vanoni, G. Zanetti, and B. Curti

675

Phototrophic bacterial flavocytochromes c M.A. Cusanovich, R.G. Bartsch, M.M. Dolata, J. Fitch, T.E. Meyer, E. Varga, and J. van Beeumen

685

Structure and function of the soluble fumarate reductase, flavocytochrome c, from Shewanella putrefaciens G.A. Reid, S.L. Pealing, F.D.C. Manson, F.B. Ward, and S.K. Chapman

695

Xanthine dehydrogenase: properties T. Nishino, T. Nishino, A.Structure Sato, T.and Page, and Y. Amaya

699

The primary structure of human xanthine dehydrogenase and chromosomal location of the gene K. Ichida, T. Hosoya, O. Sakai, S. Minoshima, N. Shimizu, Y. Amaya, K. Noda, and T. Nishino

707

Overexpression and characterization of xanthine dehydrogenase in a baculovirus-insect cell system T. Nishino, T. Saito, Y. Amaya, S. Kawamoto, Y. Ikemi, and T. Nishino

711

Redox of milk xanthine dehydrogenase J. Hunt,potentials and V. Massey

715

Electron transfer in milk xanthine dehydrogenase as studied by pulse radiolysis K. Kobayashi, M. Miki, K. Okamoto, and T. Nishino

719

A comparison of substrate specificity between milk xanthine oxidase and xanthine dehydrogenase C.M. Harris, and V. Massey

723

Crystallization and preliminary x-ray diffraction of xanthine oxidase isolated from bovine milk B.T. Eger, U. Eikmanns, E.F. Pai, M. Sato, K. Okamoto, and T. Nishino

727

New tight binding inhibitorsand of T. xanthine oxidase K. Okamoto, T. Iwamoto, Nishino

731

Molecular cloning and structural characterization of the gene locus coding for mouse xanthine oxidoreductase G. Cazzaniga, E. Garattini, P.L. Schiavo, F. Segalla, F. Galbiati, and M. Terao

735

Mouse L929 cells are defective in the expression of xanthine oxidoreductase enzymatic activity and molybdenum (VI) salts can complement the deficit F. Falciani, M. Terao, S. Goldwurm, M.L. Calzi, M. Salmona, G. Cazzaniga, E. Garattini, A. Ronchi, A. Gatti, and C. Minoia

739

XIX Physicochemical properties of retinal oxidase purified from rabbit hepatic cytosol M. Tsujita, S. Tomita, Y. Matsuo, S. Miura, and Y. Ichikawa

743

Mutagenesis of the cysteine residues of the FAD domain of nitrate reductase W.H. Campbell, and U.N. Dwivedi

747

Electron transfer and prototropic equilibria in two complex metalloflavoproteins R. Hille, and R.J. Rohlfs

751

The absence of protoheme in active respiratory complex II of the thermoacidophilic archaeon, Sulfolobus sp. strain 7 T. Iwasaki, T. Wakagi, T. Oshima, and K. Matsuura

755

Unique properties of the "rotenone site" of NADH-Q oxidoreductase T.P. Singer, and R.R. Ramsay

759

Reconstitution of a cell-free superoxide generation of human neutrophil Y. Nisimoto, and H.O. Murakami

763

FLAVOPROTEINS CONTAINING COVALENTLY-BOUND FLAVINS Preparation and properties of recombinant corynebacterial sarcosine oxidase L.J. Chlumsky, L. Zhang, and M.S. Jorns

769

One-step cloning and overexpression of the sarcosine oxidase operon from Corynebacterium sp. P-l L.J. Chlumsky, A.J. Ramsey, and M.S. Jorns

779

L-Gulono-y-lactone oxidase — cDNA cloning and elucidation of the genetic defect in a mutant rat with osteogenic disorder K. Yagi, and M. Nishikimi

783

High-level expression of rat L-gulono-y-lactone oxidase in silkworm cells with a baculovirus vector M. Nishikimi, K. Yagi, and J. Kobayashi

791

Structures responsible for FAD binding and substrate recognition of rat liver monoamine oxidase F. Ogata, Y. Tsugeno, I. Hirasiki, J. Mitoma, and A. Ito

795

Vanillyl-alcohol oxidase from Pénicillium simplicissimum : A novel flavoprotein containing 8a-(A/3-histidyl)-FAD W.J.H. van Berkel, M.W. Fraaije, E. de Jong, and J.A.M. de Bont

799

Flavinylation of the trimethylamine dehydrogenase of bacterium W3A1 expressed in Escherichia coli N.S. Scrutton, L.C. Packman, and F.S. Mathews

803

XX A new family of flavoenzymes? R. Brandsch

807

E N Z Y M E S USING REDUCED FLAVIN FOR THEIR ACTIVITY The bacterial luciferase reaction: Model or maverick in flavin biochemistry? J.W. Hastings

813

Kinetic control of folding and assembly of heterodimeric bacterial luciferase T.O. Baldwin, M.M. Ziegler, J.F. Sinclair, A.C. Clark, and A.-F. Chaffotte

823

Kinetic mechanism of the bacterial luciferase reaction W.A. Francisco, H.M. Abu-Soud, A.C. Clark, F.M. Raushel, and T.O. Baldwin

829

Bacterial luciferase: Bioluminescence emission using lumazines as substrates P. Macheroux, S. Ghisla, and J.W. Hastings

839

Studies on Escherichia coli chorismate synthase S. Bornemann, M.N. Ramjee, D.J. Lowe, R.N.F. Thorneley, J.R. Coggins, C. Abell, S. Balasubramanian, T.R. Hawkes, W.W. Nichols, and G.M. Davies

843

Characterization of the Vibrio harveyi expressed in Escherichia coli B. Lei, and S.-C. Tu

847

NADPH:FMN oxidoreductase

LIST O F PARTICIPANTS

851

AUTHOR INDEX

857

S U B J E C T INDEX

865

THEORETICAL AND CHEMICAL APPROACH

Use of Molecular Orbital Calculations in Studies on Mechanisms of Enzyme Catalysis

I.M.C.M. Rietjens, M.G. Boersma, A.E.M.F. Soffers, N.H.P. Cnubben, J. Koerts, S. Peelen , W.J.H. van Berkel, C. Veeger and J. Vervoort Department of Biochemistry, Agricultural University, NL-6703 HA Wageningen, The Netherlands

Introduction Frontier orbital theory has been applied in organic chemistry to explain the orienting effect of substituents in aromatic molecules in electrophilic, radical or nucleophilic reactions in solution (1,2). However, relative differences in the reactivity of various sites in an aromatic compound can be expected to affect the possibilities for conversion of the molecule in the active site of an enzyme as well. The present paper describes results that demonstrate that molecular orbital calculations in combination with frontier orbital theory are a useful additional tool to study mechanisms of enzyme catalysis in not only enzymes with a relative large aspecific active site (the mammalian cytochromes P450) but also for flavin containing monooxygenases with more specific active sites. Results and Discussion Mammalian cytochromes P450. As a first example on the use of molecular orbital calculations in studies on enzyme catalysis a series of non-flavin enzymes was used, namely the mammalian cytochromes P450. This was done because the mammalian cytochromes P450 are enzymes known to have a relatively large aspecific active site. The enzymes catalyze the monooxygenation of a broad range of endogenous as well as xenobiotic compounds. Studying the conversion of relatively small substrate molecules (fluorobenzenes) in the relatively large active site of the cytochromes P450 must be a suitable system to investigate the role of chemical reactivity, i.e. the usefulness of molecular orbital calculations and frontier orbital theory in studies on enzyme catalysis. The fluorobenzenes were chosen as the model substrates because the fluorine substituent has almost the same Van der Waals radius as a hydrogen and will not cause much sterical

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

4 hindrance. Besides, the fluorine substituent m a k e s it possible to determine the regioselectivity of substrate conversion by

,9

F N M R . Finally, the fluorine can be

expected to have significant effects on the molecular orbital characteristics at the various carbon centres in the substituted benzene ring. T h e active site of the cytochromes P450 contains an iron(III)protoporphyrin IX which is converted to a high valence iron-oxo species (P45C)(FeO) 3+ ) during the catalytic cycle. T h e P450(FeC))3 + species contains radical characteristics on the oxygen atom and is generally accepted to catalyze substrate conversion by a radical mechanism (3). T h e exact nature of this reaction mechanism, however, is still a matter for considerable debate, especially the mechanism for the hydroxylation of aromatic rings. Figure 1 presents one of the possible hypotheses for the cytochrome P450 catalyzed aromatic hydroxylation of an aromatic ring taking fluorobenzenes as the model compounds.

Figure 1. Proposed o-addition pathway for the formation of phenolic metabolites f r o m fluorobenzenes by cytochromes P450. The pathway presented assumes that no epoxide intermediates are formed, which implies that the site of initial attack of the P450(FeC)) 3 + on the benzene ring will be the site of hydroxylation. The cationic a-adduct is formed f r o m the radical a-adduct by electron abstraction f r o m the aromatic ring by the Fe atom. Other pathways suggested in the literature include epoxide intermediates or initiation of the reaction by electron abstraction f r o m the aromatic ring (3,4).

In molecular orbital terms the interaction between the single occupied molecular orbital ( S O M O ) of the P 4 5 ( ) ( F e O ) 3 + species and the reactive Ti-electron system in the highest occupied moleculer orbital ( H O M O ) of the fluorobenzene molecule can be depicted as presented in figure 2. Interaction between the P450 ( F e O ) 3 + S O M O and the H O M O of the fluorobenzene results in an energy gain of 2E1-E2, which can be used to overcome the activation energy of the reaction.

5 Energy

IXIMO SCMO

HOMO

-H-

(FeO)3

3+

n

Figure 2. Molecular orbital scheme of the interaction between the active iron o x o P 4 5 0 ( F e O ) 3 + intermediate and the substrate in the active site of a cytochrome P450. T h e energy gained by the interaction (2E1-E2) can be used to o v e r c o m e the activation barrier.

If the m e c h a n i s m depicted in figure 1 is the correct m e c h a n i s m for the reaction, the site of so-called c - a d d i t i o n will be the site of hydroxylation. A n d if there is no influence of the active site imposing a specific orientation of the substrate with respect to the (FeO)3+, the site of G-addition, i.e. the site of hydroxylation, may be predicted by the distribution of the reactive H O M O electrons in the fluorobenzene ring. Table 1 shows that this is indeed the case and, furthermore, that the site of hydroxylation of a series of fluorobenzenes upon their in vivo metabolism by rats correlates (r = 0.960) with the values predicted on the basis of the calculated frontier orbital density for electrophilic attack.

Table 1. Predicted and Observed Regioselectivity of Fluorobenzene Hydroxylation by Male Wistar Rats. Benzene

carbon centers

predicted

observed

by M O calculations

in urine by

0.39 : 0.27 : 0.34

0.34 : 0 . 2 0 : 0.46

,9

F NMR

1-fluoro-

C2/6 : C3/5 : C 4

1.2-difluoro-

C3/6 : C4/5

0.27 : 0.73

0.32 : 0.68

1.3-difluoro-

C2 : C4/6 : C 5

0.16:0.74:0.10

0.12:0.82:0.06

1.2.3-trifluoro-

C4/6 : C5

0.68 : 0.32

0.73 : 0.27

1.2.4-trifluoro-

C3 : C 5 : C 6

0.19:0.60:0.21

0 . 2 5 : 0 . 5 1 : 0.24

6 Table 2. Regioselectivity of 1,2-DifIuorobenzene Hydroxylation by Microsomal Preparations Containing Different Cytochrome P450 Enzyme Patterns. To achieve high enough conversion rates, tert.butylhydroperoxide was used as an artificial oxygen donor. For isosafrole and acetone microsomes (with the highest turnover rates) it could be demonstrated that results were similar to those obtained in a NADPH/oxygen supported reaction. P450 inducer

hydroxylation ratio at C3/6 : C4/5

total rate of conversion nmol/10 min/ nmol P450

none

0.29 : 0.71

3.5 ± 0 . 3

phénobarbital

0.28 : 0.72

2.1 ± 0 . 3

3-methylcholanthrene

0.26 : 0.74

3.1 ± 0 . 8

isosafrole

0.22 : 0.78

17.2 ± 0 . 1

dexamethasone

0.27 : 0.73

1.5 ± 0 . 1

acetone

0.28 : 0.72

11.0 ± 1.3

Table 2 presents results that show that the conversion of a fluorobenzene by microsomal preparations differing in their cytochrome P450 enzyme pattern results in similar regioselectivity for the aromatic hydroxylation. This confirms that the active sites of the cytochromes P450 do not have a dominant role in determining the regioselectivity of the 1,2-difluorobenzene conversion. Thus, the hydroxylation of fluorobenzenes by the mammalian cytochromes P450 appears to be a system in which enzymatic conversion is mainly (>95%) determined by chemical reactivity of the substrate molecules. Altogether, the results provide support for a mechanism in which the aromatic hydroxylation proceeds by an initial electrophilic attack of the P450(FeO)^+ on the 7telectron system of the fluorobenzenes and results in formation of phenolic metabolites without formation of intermediate epoxides. Flavin containing monooxvgenases. The results presented above demonstrate that molecular orbital calculations proved a useful additional tool for studies on the reaction mechanism of cytochromes P450 converting a series of relatively small substrates. However, additional studies demonstrated that the approach is also useful for studies on enzyme catalysis by enzymes with a more specific active site and a narrow substrate

LUMO

.--""f^-

-HC(4a)-peroxyflavin H

H

HOMO

^ reaction intermediate

substrate

O-

Figure 3. Schematic presentation of the reaction taking place in the active site of flavin containing monooxygenases: an electrophilic attack of the C(4a)-peroxyflavin on the HOMO electrons of the substrate, i.e. a nucleophilic attack of the HOMO electrons of the substrate on the LUMO of the C(4a)-peroxyflavin. R= COO" for conversions by 4hydroxybenzoate-3-hydroxylase, R=H for conversions by phenol hydroxylase. The possible deprotonation of the hydroxyl moiety is investigated.

specificity, such as for example the flavin containing monooxygenases. These enzymes are known to catalyze substrate monooxygenation via formation of a so-called C(4a)peroxyflavin intermediate, which is generally accepted to result in substrate conversion through either an electrophilic attack of the C(4a)-peroxyflavin on the substrate (figure 3) or through a radical mechanism proceeding by electrophilic attack of a hydroxyl radical resulting from homolytic cleavage of the C(4a)-peroxyflavin dioxygen bond (5,6). Figure 3 also presents that in molecular orbital terms a nucleophilic attack by the substrate implies that the HOMO characteristics of the substrate will be of importance for its conversion. As a consequence, reactivity of the substrate, and, thus, chances for its conversion, will be dependent on i) the energy of these reactive HOMO electrons and ii) the density of the reactive HOMO electrons on the reaction centre. Using this concept, studies on two flavin containing enzymes, namely 4-hydroxybenzoate hydroxylase and phenol hydroxylase were performed. 4-Hydroxybenzoate-3-hydroxylase. 4-Hydroxybenzoate-3-hydroxylase catalyzes the conversion of 4-hydroxybenzoate to 3,4-dihydroxybenzoate. In addition to the normal substrate the enzyme is capable of catalyzing the 3-hydroxylation of a series of fluorinated substrate analogues, although at a different maximal conversion rate (5).

8 Table 3. Effect of Deprotonation of the Hydroxyl Moiety of 4-Hydroxybenzoate Derivatives on Their Calculated Molecular Orbital Characteristics at The Reaction Centre For Aromatic Hydroxylation by 4-Hydroxybenzoate-3-hydroxylase, i.e. at C3 (7). compound

total charge

çffçctorç

HOMO density

0.01 0.01

4-fluorobenzoate

-0.19

benzoate

-0.17

substrates

OH

4-hydroxybenzoate

-0.21

-0.37

2-fluoro-4-hydroxybenzoate

-0.24

-0.41

5-fluoro-4-hydroxybenzoate

-0.17

-0.36

3,5-difluoro-4-hydroxybenzoate

+0.02 -0.17

tetrafluoro-4-hydroxybenzoate

-0.02

o-

-0.20

OH

0.01 0.01 0.01 0.01 0.01

E(HOMO) eV

-4.88 -4.67

o-

o-

OH

0.21

-4.74

+0.44

0.19

-4.86

+0.13

0.17

-4.95

+0.24

0.20

-5.16

+0.04

0.20

-5.41

-0.49

Substrate analogues without the 4-hydroxyl moiety have been demonstrated to bind to the enzyme and to induce formation of the reactive C(4a)-peroxyflavin intermediate, but they cannot be converted. From the existence of these so-called effectors it has been concluded that the presence of the 4-hydroxyl moiety is essential for substrate conversion. Deprotonation of this 4-hydroxyl function of the substrate has been suggested to facilitate the electrophilic attack of the C(4a)-perxoyflavin -or a hydroxyl radical derived from iton the substrate. Results from molecular orbital calculations (Table 3) appeared useful in providing insight in this activation of the 4-hydroxybenzoates upon deprotonation of their hydroxyl moiety. From these data it follows that the 4-hydroxybenzoates are activated upon deprotonation of thei'/ 4-hydroxyl moiety because of i) an increase in the HOMO electron density for electrophilic attack on C3 and ii) an increase in the energy of the reactive electrons in the H O M O of the substrate. The results also explain that without the possibility for this deprotonation of the hydroxyl moiety (for instance in the effectors benzoate and 4fluorobenzoate) C3 hydroxylation can not occur: the reaction centre at C3 contains no significant reactive electrons, reducing chances for an electrophilic attack at this position to almost zero. In addition, the E(HOMO) of these effectors is too low for catalysis to

9 occur. Furthermore, the results indicate that deprotonation of the hydroxyl moiety of the hydroxybenzoate derivatives does not activate these substrates because of increased total negative charge on C3. This follows from the observation that the total negative charge on C3 in the effector 4-fluorobenzoate equals that on C3 in the deprotonated form of the substrate 3,5-difluoro-4-hydroxybenzoate. That an increase in the energy of the reactive electrons actually results in activation of the substrate, and, thus, increased chances (rates) for conversion is also demonstrated by the results presented in figure 4. Using kcat values reported in the literature by Husain et al. (5) a clear correlation (r=0.992) between the In of the rate of conversion of a series of fluorinated substrates (pH 8, 25 ° C ) and their calculated E ( H O M O ) was obtained. The data support our present working hypothesis that at pH 8, 25 ° C substrate conversion by 4-hydroxybenzoate-3-hydroxylase is dependent on H O M O characteristics of the substrate with the C(4a)-peroxyflavin attack on the substrate being the rate limiting step for catalysis.

Figure 4. Correlation (r=0.992) between the In kcat (values taken from Husain et al. (5)) and the calculated E(HOMO) of the benzoate derivatives, for further details see (7).

Phenol hydroxylase. Phenol hydroxylase catalyzes the conversion of phenol to 1,2dihydroxybenzene (catechol). The results presented here are based on studies with phenol hydroxylase from the yeast Trichosporon

cutaneum. In addition to the parent substrate

the enzyme catalyzes the hydroxylation of various substituted phenols (8). Whether deprotonation of the phenolic substrates is a prerequisite for their conversion by the enzyme is still a matter for debate. This was studied in more detail using molecular orbital calculations and 3-fluorophenol as the model substrate. Conversion of

10 3-fluorophenol by phenol hydroxylase results in formation of both possible catechol products, i.e. 3-fluoro-l,2-dihydroxybenzene and 4-fluoro-l,2-dihydroxybenzene, although to a different extent (Figure 5a). T h e actual ratio between these two products resulting f r o m hydroxylation at respectively C2 and C6 appeared to vary with the p H of the incubation medium (Figure 5a), showing a decrease in the C6: C2 ratio with increasing pH. Furthermore, an increase in the rate of product formation with increasing p H was observed (Figure 5b).

Figure 5. Effect of pH on a) the ratio C6:C2 hydroxylation and b) the total rate of product formation of 3-fluorophenol by phenol hydroxylase. Because the pKa of 3-fluorophenol itself is 9.3, the effects presented in Figure 5 -with an apparent pKa around 6 -, are rather ascribed to a protonation/deprotonation equilibrium of an amino acid residue in the active site of the protein.

19

F N M R binding studies of

3-fluorophenol bound to the reduced form of phenol hydroxylase at pH 7.6 (data not shown) demonstrate that the substrate is not bound in two distinct ways, although at this pH value two different catechol products are observed. This result implies that the substrate rotates at a rate that exceeds the time scale of the N M R measurement, and, thus, at a rate that exceeds the rate of enzyme catalysis. Molecular orbital calculations on 3-fluorophenol (results presented in Table 4) provided additional insight in the mechanism of the conversion of 3-fluorophenol by phenol hydroxylase. T h e calculations revealed that 3-fluorophenol has a higher intrinsic chemical reactivity (increased E ( H O M O ) and increased H O M O density at C2 plus C6) upon deprotonation of its hydroxyl moiety (Table 4). Thus, our present working hypothesis states that an active site amino acid residue acts as a base in the reduced form of the protein. Deprotonation of this residue (pKa around 6) results in hydrogen bond formation with the hydroxyl moiety of the phenolic substrate, leading to (partial) deprotonation of the

11 Table 4. Calculated HOMO Characteristics of 3-Fluorophenol and its Phenolate Anion. compound

E(HOMO)

HOMO density on carbon centre C4

C5

C6

in eV

0.11

0.29

0.01

0.22

-9.35

0.00

0.31

0.00

0.23

-3.05

CI

C2

C3

3-fluorophenol

0.18

0.03

3-fluorophenolate

0.03

0.22

phenolic substrate. This partial deprotonation results in an increased chemical reactivity of the substrate for an electrophilic attack by the C(4a)-peroxyflavin, resulting in higher kcat with increasing pH. This latter conclusion implies that the phenolic substrate is converted in its (partially) deprotonated form. The conclusion also implies that the electrophilic attack of the C(4a)-peroxyflavin, or a hydroxyl radical derived from it, on

E (HOMO)

(eV)

Figure 6. Correlation (r=0.884) between the In k C at for conversion of phenol derivatives by phenol hydroxylase, and their calculated E(HOMO). E(HOMO) values presented are those of the phenols with a deprotonated hydroxyl moiety, because k C at values were determined at pH 7.6 (170 |iM NADPH). Values were corrected for decoupling.

12 the frontier ^-electrons of the substrate is the rate limiting step for enzyme catalysis at physiological pH in the absence of monovalent anions. This assumption is supported by the results presented in figure 6 which demonstrate that, as for the conversion of a series of fluorinated substrate analogues by 4-hydroxybenzoate hydroxylase (Figure 4), the In k c a t for the conversion of a series of fluorinated substrate analogues by phenol hydroxylase correlates (r= 0.884) with the calculated energy of their HOMO electrons (Figure 6).

Conclusions On the basis of studies with series of fluorinated aromatic substrates converted by mammalian cytochromes P450 or by flavin containing monooxygenases, i.e. 4-hydroxybenzoate-3-hydroxylase from Pseudomonas fluorescens and phenol hydroxylase from Trichosporon cutaneum, it has been demonstrated that chemical reactivity is an important factor influencing enzyme catalysis. In the examples presented chemical reactivity of the enzyme cofactor and the substrate even seemed to be the main factor determining the outcomes of enzyme catalysis, such as the regioselectivity and the rate of the conversion. These results now open the way to study other factors influencing enzyme catalysis, such as sterical hindrance, substrate orientation and conversion by mutant enzymes, by studying deviations observed from calculated chemical reactivity.

References 1. Fleming, I., 1976. Frontier orbitals and organic chemical reactions. John Wiley & Sons, Chichester. 2. Fukui, K„ T. Yonezawa, C. Nagata and H. Shingu. 1954. J. Chem. Phys. 22, 1433-1442. 3. Guengerich, F.P. and T.L. MacDonald. 1990. FASEB J. 4, 2453-2459. 4. Rietjens, I.M.C.M., A.E.M.F. Soffers, C. Veeger and J. Vervoort. 1993. Biochemistry 32, 4801-4812. 5. Husain, M., B. Entsch, D.P. Ballou, V. Massey and P.J. Chapman. 1980. J. Biol. Chem. 255, 4189-4197. 6. Anderson, R.F., K.B. Patel and B. Vojnovic. 1991. J. Biol. Chem. 266, 13086-13090. 7. Vervoort, J., I.M.C.M. Rietjens, W.J.H. van Berkel and C. Veeger. 1992. Eur. J. Biochem. 206, 479-484. 8. Neuhjahr, H.Y., K.G. Kjellen. 1978. J. Biol. Chem. 253, 8835-8841.

A T H E O R E T I C A L A P P R O A C H OF T H E M E C H A N I S M OF C ( 4 a ) - P E R O X Y F L A V I N C A T A L Y Z E D REACTIONS.

C. Veeger, I.M.C.M. Rietjens and J. Vervoort Department of Biochemistry, Agricultural University, NL-6703 HA Wageningen, The Netherlands

J. Lee Department of Biochemistry, University of Georgia, Athens, Georgia, USA

INTRODUCTION

Several flavin containing enzymes are known to catalyze their reactions by means of oxygen activation and formation of a so-called intermediate C(4a)-peroxyflavin (figure 1). The present paper focusses on some problems under investigation on the functioning of the reactive C(4a)-peroxyflavin.

Figure 1. Structure of the reactive C(4a)-peroxyflavin intermediate -protonated and deprotonated- formed upon two electron reduction and oxygen binding to the flavin cofactor.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

14 NUCLEOPHILIC VERSUS ELECTROPHILIC MECHANISMS

It is well recognized that the mechanisms of substrate conversion by the C(4a)peroxyflavin can be either electrophilic or nucleophilic (1). In a molecular orbital scheme such reactions can be schematically presented as depicted in figure 2 (ref. 2). This implies that, from a chemical point of view, the nature of the reaction actually catalyzed by the C(4a)-peroxyflavin will be dependent on i) the orbital characteristics of the peroxyflavin and possible influences on these characteristics brought about by the surrounding protein, but as well on ii) the orbital characteristics of the substrate to be converted.

Mf 4f

4f

nucleophilic

R-CH=0

•H-

electrophilic

C(4a)-peroxyflavin

00' luciferase cyclohexanone

- H -

monooxygenase

OOH phenol hydroxylase 4-hydroxybenzoate-3-hydroxylase

Figure 2. Molecular orbital diagram for an electrophilic and a nucleophilic attack by the C(4a)-peroxyflavin on respectively the deprotonated aromatic substrates of phenol hydroxylase and 4-hydroxybenzoate-3-hydroxylase as well as the aldehyde substrates of luciferase and cyclohexanone monooxygenase. For efficient interaction with the LUMO of the aldehydes and the energetically comparable HOMO of the aromatic substrates (Table 1), the form of the C(4a)-peroxyflavin catalyzing the attack must be different. For a nucleophilic attack on the aldehydes the energy of the HOMO of the C(4a)-peroxyflavin has to be increased compared to the situation where the electrophilic LUMO attack on the aromatic substrates occurs. This increase in energy of the molecular orbitals of the C(4a)-peroxyflavin could be brought about by deprotonation of the peroxy moiety.

From the diagram presented in figure 2 it can be derived that substrates with high

15 E(HOMO) orbitals will have a higher tendency to react through an electrophilic attack by the C(4a)-peroxyflavin, whereas substrates with relatively low energy orbitals will be better substrates for reactions proceeding by a nucleophilic attack. As a rule, the LUMO of the C(4a)-peroxyflavin will react with the HOMO of the substrate in case of an electrophilic attack by the C(4a)-peroxyflavin. Whereas in case of a nucleophilic attack the LUMO of the substrate will react with the HOMO of the C(4a)-peroxyflavin. This can be further illustrated by results from MO-calculations on the substrates of the following flavin containing model enzymes, i.e. 4-hydroxybenzoate-3-hydroxylase and phenol hydroxylase, proposed to catalyze monooxygenation of their aromatic substrates via an electrophilic attack by the C(4a)-peroxyflavin on the substrate (3-5), and bacterial luciferase and cyclohexanone monooxygenase both proposed to catalyze the reaction through a nucleophilic attack by the C(4a)-peroxyflavin on their substrates (1,6,7). The apparent difference between the mechanisms proposed for luciferase and cyclohexanone monooxygenase is that in the case of luciferase the C(4a)-peroxyflavin species is suggested to be protonated (1,6), whereas that in the cyclohexanone monooxygenase is suggested to be deprotonated (7). Table 1 presents the calculated MO energies of the respective substrates. The data presented in Table 1 support the view that both the hydroxyl moiety of phenol and 4-hydroxybenzoate have to be deprotonated to increase their nucleophilic reactivity, i.e. increase their E(HOMO), to allow an efficient electrophilic attack by the reactive C(4a)-peroxyflavin. The increase in the energy of the reactive electrons observed upon this deprotonation is about 5-6 eV. The results presented in Table 1 also suggest that for an efficient electrophilic or nucleophilic attack brought along by the same C(4a)-peroxyflavin intermediate the characteristics of the reactive enzyme cofactor need to vary to a certain extent. This follows from the following argument. Figure 3 schematically depicts the rule, given by frontier orbital theory (2), that the overlap between two reactive orbitals, and, as a result, the amount of energy gained by this interaction, decreases when the energy difference between the reacting E(HOMO) of the nucleophile and the E(LUMO) of the electrophile increases. The energy gained and, thus, the chances for reactivity are highest when there is a minimum difference between the E(HOMO) of the nucleophile and the E(LUMO) of the electrophile. Figure 2 already schematically presented the consequences of this phenomenon for the different reactions catalyzed by the C(4a)-peroxyflavin cofactor.

16 Table 1. MO-Characteristics of the Substrates of 4-Hydroxybenzoate-3-hydroxylase, Phenol Hydroxylase, Luciferase and Cyclohexanone Monooxygenase. For both aromatic compounds the values for their forms with a deprotonated hydroxyl moiety are also presented. The value of the M O of importance for catalysis is underlined. enzyme

E(HOMO)

E(LUMO)

in eV

in eV

phenol

-9.11

+0.40

phenolate anion

iZfl

+5.63

substate

phenol hydroxylase

4-hydroxybenzoate-3-hydroxylase

4-OH-benzoate

-4.74

+3.89

4-0"-benzoate

+0.44

+8.81

luciferase

CH3(CH2)12-CH=0

-10.59

+0.87

cyclohexanone monooxygenase

cyclohexanone

-10.30

±022

UUMO

HOMO

um

||

-tf"

-H-

n -H-

-tf -if

M F ' -tf

Figure 3. Molecular orbital scheme for the interaction between the HOMO and LUMO in case of a relatively small (left) and large (right) energy difference between E(HOMO) of the nucleophile and E(LUMO) of the electrophile. The energy gained in case of a small energy gap is higher, resulting in higher chances for conversion, than in case of a large energy gap, i.e. 2E\ is higher than 2E2-

17 As demonstrated before (8) an electrophilic attack of the protonated form of the C(4a)peroxyflavin on aromatic substrates, in for example phenol hydroxylase and 4hydroxybenzoate-3-hydroxylase, requires deprotonation of the hydroxyl moiety of the substrate, resulting in activation of the HOMO electrons of the substrate and, thus, more efficient catalysis. This must be due to a smaller energy difference between E(HOMO) of the substrate and E(LUMO) of the C(4a)-peroxyflavin in its protonated state (figure 2). However, the results presented in Table 1 demonstrate that the energy of the reactive LUMO of the aldehyde substrates of luciferase and cyclohexanone monooxygenase are similar to the energy of the HOMO in the deprotonated aromatic substrates of 4hydroxybenzoate dehydrogenase and of phenol hydroxylase. This implies that, for an efficient nucleophilic attack of the C(4a)-peroxyflavin on the LUMO of the aldehyde substrates, the energy of the molecular orbitals of the C(4a)-peroxyflavin cofactor has to be increased to result in an energy of its HOMO, (not its LUMO), around the E(LUMO) of the aldehyde substrates (Figure 2). Such an increase in molecular orbital energy can be expected upon deprotonation of the peroxide moiety. This supports the conclusion that for an efficient nucleophilic attack of the C(4a)-peroxyflavin cofactor on the aldehyde substrates the peroxide moiety has to be deprotonated, whereas for an efficient electrophilic attack on the aromatic substrates the peroxide moiety should be protonated. These considerations are in accordance with the mechanism for cyclohexanone monooxygenase suggested by Walsh (7) and of 4-hydroxybenzoate-3-hydroxylase and phenol hydroxylase suggested by Maeda-Yorita and Massey (5) and by Vervoort et al. (8) and Rietjens et al. (9).

T H E R A T E L I M I T I N G S T E P IN C(4a)-PEROXYFLAVIN D E P E N D E N T M O N O O X Y G E N A S E CATALYSIS

Using stopped flow kinetics and spectral properties of the various intermediates in the reactions, values for kinetic parameters for the C(4a)-peroxyflavin catalyzed intermediate reactions have been obtained (3, 5). In general these data were determined in the presence of enzyme inhibitors or at inhibiting low pH values, to slow down the rate of the reaction, thus allowing detection of the various reaction intermediates. However, this raises the question whether extrapolation of such kinetic data to the situation of enzyme catalysis

18 under optimal turnover conditions is permitted. A more useful approach to obtain insight in the rate limiting step under optimal conditions for enzyme turnover is, in our opinion, to study the effect of systematic changes in either the flavin cofactor (6) or in the substrate (8, 9) on the overall rate of catalysis under optimal turnover conditions. When a correlation between the maximum reaction rate and the parameter characterizing the chemical change in either the flavin or the substrate is observed, this points to the involvement of the respective chemical characteristics of the flavin cofactor or the substrate in the rate limiting step of catalysis. From this a conclusion can be drawn on the actual rate limiting step in catalysis under optimal conditions for substrate turnover. Such a correlation was found for 4-hydroxybenzoate-3-hydroxylase converting a number of fluorinated 4-hydroxybenzoates. A linear relation was found (8) between In kc at as measured by Husain et al. (3) and the value of the E(HOMO) of the benzoate derivatives. A similar linear correlation exists for the conversion of 17 different substrates by phenol hydroxylase (9). Assuming that the attack of the C(4a)-peroxyflavin on the substrate is in approach bimolecular, its rate constant is k = P (kT/h) exp - AH 0 # /RT . exp AS0*/R in which P is the probability of the attack. Upon comparing the reactivities of the deprotonated substrates (assuming AS* remains constant, in other words the affinities of all substrates for the enzyme have the same order of magnitude, and no gross structural changes -leading to changes in water content, in polarity, or the dielectric constant of the active centre- occur), the effect of the gain of energy (2E) by the interaction of LUMO and HOMO is strictly on AH 0 *. Upon comparing the relations between In k c a t and E(HOMO) of the deprotonated substrates of the enzymes 4-hydroxybenzoate-3-hydroxylase and phenol hydroxylase (8, 9) a striking relation between the slopes of the curves is apparent. In both cases the slope is around 3 eV _1 , i.e. about a 20-fold increase in kc a[ per eV change in E(HOMO). From this relation one can conclude that in both enzymes the substrates pass by identical or at least very similar pathways through their transition states upon attack by the C(4a)peroxyflavin. Despite this identical relation, extrapolation of the activities of the natural substrates from one enzyme to the other is not allowed. In view of the 3 eV difference of the E(HOMO)'s

19 of the deprotonated natural substrates (Table 1) the k c a t of phenol hydroxylase could be expected to be about 1/10,000 of the k c a t of 4-hydroxybenzoate-3-hydroxylase, instead of about 1/10 as observed. This discrepancy can in theory be explained by 3 factors. There may be a difference in the probability factor P or in ASq* for the reactions catalyzed by the two enzymes. P is mainly determined by the vibrational and rotational contributions of each reactant to its partition function and influenced probably in enzyme reactions by factors like orientation and distance of the reactants. Factors contributing to a higher A S 0 * could be substrate-induced flexibility around the active centre, so-called substates as have been observed in other flavoproteins by Bastiaens et al. (10). Besides, there may be a difference in the E(LUMO) of the C(4a)-peroxyflavin in the active site of the two monooxygenases. Figure 4 shows for a series of benzoquinone model compounds that the correlation between their redox potentials and their E(LUMO) demonstrates a change in redox potential of about 1 V connected to a change of E(LUMO)

-3

-2.5

-2 E

-1.5

-1

eV

LUM 0 ( )

Figure 4. Correlation between the midpoint potentials of a series of substituted benzoquinones and the calculated E(LUMO). 2,3 DC-5,6 DCNBC1 is 2,3 dichloro 5,6 dicyano; 2356 T C is tetrachloro; TB is tetrabromo; D C is dichloro; TM is tetramethyl.

The observed correlation between k 0 [, s for the luciferase reaction and the calculated oneelectron oxidation potential of a series of FMN-analogs (6) can also be interpreted in

20 terms of an influence of E(LUMO) of the flavin analog on the reaction rate. From this it follows that a change in redox potential of the flavins, i.e. especially the C(4a)-peroxyflavin, brought about by the protein matrix can influence the reactivity of the cofactor by adjusting its E(LUMO) to the E(HOMO) of its natural substrate. Extrapolation of the slope of the In k c a t - E(HOMO) relation of the depronated substrates of 4-hydroxybenzoate-3-hydroxylase and phenol hydroxylase to the E(HOMO) -value of the protonated substrate (assuming identical P- and AS 0 * -values) may lead to the conclusion that the latter form of the substrate is converted at a rate of around 10"6 - 10"8 of the deprotonated substrate. Thus, on the basis of the theoretical considerations presented, it is concluded that the protonated forms can not be converted. It seems unlikely that the probability factor of the reaction of the protonated form will be much higher than that of the deprotonated substrate. However, compensation by an increase of ASo* in the order of 100-130 J . m o l 1 K could lead to an appreciable activity (about 10%) of protonated substrate conversion. Although such high AS 0 * values have been measured in more complex enzymatic redox reactions (11), it appears rather unlikely for a proton of the substrate to induce such an entropic effect. Thus our tentative conclusion is that the protonated forms of the substrates of both 4-hydroxybenzoate-3-hydroxylase and phenol hydroxylase are not or hardly detectable converted.

CONCLUSIONS

It is generally accepted that enzymes accelerate chemical reactions by lowering the activation energy. In most biochemical text books this is suggested to occur through lowering of the energy through effective binding of the transition state. However, the same effect can be achieved by increasing the reactivity of the substrate and/or the enzyme cofactor involved in the catalytic mechanism. For flavin-containing enzymes which catalyze their reaction through formation of the socalled C(4a)-peroxyflavin intermediate this activation is not only dependent on the chemical characteristics of this cofactor as influenced by the protein chain, but also, and perhaps even to a larger extent, on the chemical characteristics and possible activation of the respective substrates.

21 Present evidence may indicate that within the class of flavin monooxygenases the hydroxylation of the aromatic substrates by the C(4a)-peroxyflavin proceeds through transition states that have similar, may be identical, electronic properties.

ACKNOWLEDGEMENTS

This work was supported by the Netherlands Foundation for Chemical Research (SON) with financial aid from the Netherlands Organisation for Scientific Research (NWO). J.L. is an NIH Fogerty Senior International Fellow.

REFERENCES

1.

Ghisla, S. and V. Massey. 1989. Eur. J. Biochem. 181, 1-17.

2.

Fleming, 1., 1976. Frontier orbitals and organic chemical reactions. John Wiley & Sons, Chichester.

3.

Husain, M„ B. Entsch, D.P. Ballou, V. Massey, and P.J. Chapman. 1980. J. Biol. Chem. 255, 4189-4197.

4.

Anderson, R.F., K.B. Patel, and B. Vojnovic. 1991. J. Biol. Chem. 266, 13086-13090.

5.

Maeda-Yorita, K. and V. Massey. 1993. J. Biol. Chem. 268, 4134-4144.

6.

Eckstein, J.W., J.W. Hastings, and S. Ghisla. 1993. Biochemistry 32, 404-411.

7.

Walsh, C.T., and Y.C.J. Chen. 1988. Angew. Chem. 100, 342-352.

8.

Vervoort, J., I.M.C.M. Rietjens, W.J.H. van Berkel, and C. Veeger. 1992. Eur. J. Biochem. 206, 479-484.

9.

Rietjens I.M.C.M., M.G. Boersma, A.E.M.F. Soffers, N.H.P. Cnubben, J. Koerts, S. Peelen, W.J.H. van Berkel, C. Veeger, and J. Vervoort. 1993. This issue.

10.

Bastiaens, P.I., A. van Hoek, W.F. Wolkers, J.C. Brochon, and A.J.W.G. Visser. 1992. Biochemistry 31, 7050-7060.

22 11.

Mensink, R.E., and H. Haaker. 1992. Eur. J. Biochem. 208, 295-299.

12.

Peover, M.E., 1962. Nature 193, 475-476.

SEMIEMPIRICAL CALCULATIONS ON PROPERTIES OF (ISO)ALLOXAZINES IN GROUND AND EXCITED STATES

Henryk Szymusiak Faculty of Commodity Science, University of Economics in Poznan, Al. Niepodleglosci 10, 60-967 Poznan, Poland

Introduction

Among the molecules which display an excited-state proton transfer (ESPT) reaction, alloxazine is one of the most extensively studied (1-6). The alloxazine Sm')=0.008) (11). Since the quantum yield of this step is much higher than usually observed (clO - 4 for aromatic hydrocarbons) the high yield of Tn'->Sm' for isoalloxazines is of great photophysical and photochemical interest. The results of INDO/S calculations show that in the case of isoalloxazines this mechanism is indeed reasonable. The relative ordering of the calculated singlet and triplet levels of the two tautomeric forms of (A) is shown on Fig. 1.

Fig. 1. Energy level diagram of the calculated singlet and triplet states of the alloxazine form (left) and its isoalloxazine tautomer (right). The various photophysical processes are tentatively indicated for a mechanism of double proton transfer cycle, dynamics of transient absorption (2, 3) and two-step laser -induced fluorescence (TSLIF) of isoalloxazines (11).

26 On the Fig. 1. all the rut* states (usually underestimated in the INDO/S gas-phase calculations) were slightly shifted toward higher energy to obtain better consistency with the experiment (experimental extinction coefficients indicate rather rnz* for Si and Si' states). For this reason the calculated data should be treated rather quantitatively. The larger energy gap is predicted between T2 1 and Ti'. Its size of about 6000 cnr 1 (vs. 1500 cnr 1 in (A) form) is quite unusuall for a gap between triplet states. This should be important for the decay properties of the T2' state. The high yield of T-S ISC is proposed to result from a long-lived T2'(n7i*) state basing on the energy gap low and El-Sayed rules. This excitation is predicted to be localized largely in the N-5-C-4a-0-14 region. Considering the Fermi correlation one could expect some deviations from coplanarity of these atoms in the Tj'fnrc*) state. The investigations the TSLIF produced by tuning the probe wavelength far longer than the 0-0 onset of the (iso)alloxazinic emission will in part verify the above theoretical predictions.

Acknowledgements This work was supported in part by Project no. 2 1308 91 01.KBN.

References 1. Song, P.-S., M.Sun, A.Koziolowa, J.Koziol. 1974. J.Am.Chem.Soc. 96, 4319-4323. 2. Grodowski, M.S., B.Veyret, K.Weiss. 1977. Photochem.Photobiol. 26, 341-352. 3. Maclnnis, J.M., M.Kasha. 1988. Chem. Phys Lett 151,375-378. 4. Koziolowa, A., M.Stroinska, M.M.Szafran. 1990. In: Flavins and Flavoproteins (B.Curti, S.Ronchi, G.Zanetti, eds.) W.de Gruyter & Co. Berlin-New York, pp. 23-26. 5. Szymusiak, H., J.Konarski, J.Koziol. 1990. J.Chem.Soc.Perkin Trans. 2, 229-236. 6. Koziolowa, A., H.Szymusiak, J.Koziol. 1993. Pol.J.Chem. in press. 7. Kasha, M. 1987. Acta Phys.Polon. A71, 717-729. 8. Lipinski, J. 1988. Int.J.Quantum Chem. 34, 423-435. 9. Hall, L H., B.J.Orchard, S.K.Tripathy. 1987. Int.J.Quantum Chem. 31, 195-216. 10. Platenkamp, R.J., M.H.Palmer, A.J.W.G.Visser. 1987. Eur.Biophys.J. 14, 393-402 11. Richter, C., W.Hub, R. Traber, S.Schneider. 1987. Photochem.Photobiol. 45, 671-673.

P A T H W A Y S FOR A 4 a -HYDR0XY-4 a ,5-DIHYDR0FLAVIN RADICAL IN THE HYDROXYLATION OF A R O M A T I C S

Humphrey

I. X. Mager and Shiao-Chun Tu

The Department of Biochemical and Biophysical Sciences, University of Houston, Houston, Texas 77204-5934, U.S.A.

INTRODUCTION a a The postulate on the role of a 4 -hydroxy-4 ,5-dihydroflavin radical as a key transient in bacterial bioluminescence (1) was based upon the results of several model studies proving the existence of N-centered flavin radicals like III. The latter is obtained by one-electron oxidation of a a 5-ethyl-4 -hydroxy-3-methyl-4 ,5-dihydrolumiflavin (II) that is performed either chemically (2a,b), electrochemically (2c) or by pulse radiolysis (3). I l l is well defined by its redox potential, spectrum, the reversibility in an electrochemical process and by a rapid conversion to the corresponding flavinium cation or flavosemiquinone. The purpose of the present paper is to point out that III can also play a key role in a mechanistically different pathway, eventually leading to the hydroxylation of aromatics. Me j ^ N - M e ¿1

T "

6

(1)

v Me \ Y

A ^ N - M e ¿ H l l

_er (2)

Me Y VY A ^ X ^ - M e A

H

+

I (5-EtFl^)

6

+•

n (5-EtFl-4a-OH)

I I I (5-EtFMa-OH)

RESULTS A N D DISCUSSION As indicated by the results of comparable studies on aromatic hydroxylations, O-centered flavinoxy radicals like IV" were proposed to directly attack the substrate (4). So far, this proposal has not been conclusively proven due to the complication that a homolysis of the O-O bond in a dihydroflavin hydroperoxide leads to both the flavinoxy and the (hydroxylating) HO radical. Recently, a it has been emphasized (3) that the O- and N-centered flavin radical formulations like IV

b and IV

actually represent resonance structures of the same radical. It implies that III is the protonated form

Flavins and Flavoproteins 1993 © 1994 Walter de G r u y t e r & Co., Berlin • N e w York - Printed in G e r m a n y

28 Me Y

Y

Me I ,N-Me L

—

r^tS-EtFl^-O ) < ab

Y A t w^ kY m J , * —

•

ffl(s-E^-OH)

(3)

i v 1 5 (5-EtFl (5-EtfW-O )

of the flavin radical IV ' (Eq. 3). Consequently, the one-electron oxidation as formulated by Eq. 2 gives an alternative to the homolysis mentioned. It provides the principle to develop a new hydroxylating model system that does not require a dihydroflavin hydroperoxide as a starting compound. Basically, it has opened the way to unambiguously prove the postulate on the primary attack of an aromatic substrate by a (hydr)oxydihydroflavin radical. S-EtFl-^-OH

2

+

+

5-EtFlox

5-EtFl*

+

H+

5-EtFl^

+

H20

• -

+'

S-EtfW-OH

5-EtFl

5-EtFlH +

•

+

(4) (5)

2 5-EtFlH + "

+

[O]

(6)

The dihydroflavin pseudobase II is in equilibrium with the corresponding flavinium cation I (Eq. 1). The one-electron oxidation of II (Eq. 2) can be coupled to the one-electron reduction of I to the flavosemiquinone (5-EtFl ; Eq. 4). Although thermodynamically unfavorable, the process is easily observed to occur when solutions of I and II are kept under N2, rather slowly at 20 °C, but considerably enhanced at e.g. 50 °C (Fig. 1). It is followed by monitoring the appearance of the specific spectrum of 5-EtFl and/or 5-EtFlH

(Eq. 5) coming to an end due to faster, consecutive

conversions of III. In the pH 2-7 range, a pH / reaction rate profile is found (2b) to be in agreement with the mechanisms as given by Eqs. 1 and 4. The spontaneous formation of flavin radicals also proves to proceed in strong acidic solutions (0.05 - 12 N). The reaction rate increases with increasing acidity of the medium and is further influenced by the nature of the anions present. Upon lowering the pH, III rapidly reacts to give I which is recycled until the process is finished as represented by Eq. 6. Based on this principle, we have anaerobically prepared solutions of 5-EtFlH

and/or 5-EtFl in 85-95% yields in various

solvents without using a reducing agent. The yields of 5-EtFlH

/ 5-EtFl were verified by oxidizing

the flavosemiquinone with H N 0 2 leading to a corresponding recovery of I. As exemplified in Fig. 1, I (curve a) may be converted to a mixture of 5-EtFlH

and 5-EtFl (curve e). The transient

curves show five isosbestic points but, in more acidic solutions, four isosbestic points are obtained

29 with the production of 5-EtFlH

only. When the formation of 5-EtFl and/or 5-EtFlH

(Eq. 6) has

reached an optimal value, the pH of the reaction solution is adjusted to 3.0 - 3.5 to shift the equilibrium following Eq. 5 to the left. 5 - E t F l , that accounted for 85-95% of the total initial flavin, was completely removed by extraction with CHC13 to give an aqueous layer of considerable interest. Since no 0 2 and / or H 2 0 2 are produced in the overall process (Eq. 6), oxidized components of the reaction mixture should account for the missing oxygen [O]. When the flavin radicals are formed in the presence of phenylalanine or benzoic acid as test substrates, the concurrent formation of p-, m- and o-hydroxylated aromatics was established. Starting from e.g. phenylalanine, the results are expressed as "Tyr/Fl"-values (the totals of p-, m- and o-hydroxyphenylalanines per starting flavin molecule). Based on the maximal, theoretical value of Tyr/Fl= 0.5 (Eq. 6), anaerobic hydroxylations were obtained with yields of 2-40% (Tyr/Fl= 0.01 - 0.20), increasing with the acidity of the media. Besides the hydroxyphenylalanines, produced anaerobically, the aqueous layer also contained an intermediate X (Scheme 1) that is not a hydroxycyclohexadienyl radical. Consistent with the high recovery of 5-EtFl , X was only accumulated in relatively low yields. It was shown to further react in a secondary, oxidative chain reaction with 0 2 or H 2 0 2 . This remarkably increased the yields of aromatic hydroxylation without any further supply of flavin. In particular, H 2 0 2 (or the hydroxyl radical derived from it by thermal homolysis) has proven to be a very effective oxidant in the chain

12000

so

c

9000

A

/

/ l^'''

«000 .-''a

3000

\ W e

c

t i

.

5-EtFl-4 a -OH

X

+

b

X~' / / ••••

1

.a

OXIDANT

i

Wavelength (nm) Fig. 1. Spontaneous, anaerobic formation of a mixture of 5-EtFl

and 5-EtFlH

at 50 °C in 0.05 N H 2 S 0 4

+

containing 5-EtFl o x ,C10 4 (1 - 4 x 10"4 M) and phenylalanine (0.08 M); pH= 2.74. The spectra were taken at reaction times of (a) 0 (b) 60 (c) 120 (d) 150 and (e) 210 min. Isosbestic points at: 300, 379, 466, 494 and 617 nm.

SCHEME 1

30

+

5-EtFl-

Me

5-EtFl,'ox

R

/

CM

^OH H

N R H2O

o

A ^ N - M e Et

T v

SCHEME 2 reaction. In an experiment that first had anaerobically given a value of Tyr/Fl= 0.07, a subsequent treatment of the aqueous (5-EtFl -free) layer with H 2 0 2 led to a final, very high value of Tyr/Fl= 58.0 (corrected with the results of the analyses of the appropiate controls). The structure of X remains to be elucidated. Its absorption spectrum resembles the one of III (2c, 3). This gives rise to a tentative assumption that X could represent a new type of adduct (V, 4a

Scheme 2). It is formulated as the result from a homolysis of the C

-O bond in III, coupled with

the bonding of the substrate. Acknowledgements:

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

Robert A. Welch Foundation.

REFERENCES

1.

(a) Mager, H.I.X., R. Addink. 1984. In: Flavins and Flavoproteins (R.C. Bray, P.C. Engel, S. G. Mayhew, eds.) W. de Gruyter, Berlin, pp. 37-40; (b) Mager, H.I.X., S.-C. Tu. 1987. In: Flavins and Flavoproteins (D. E. Edmondson, D. B. McCormick, eds.) W. de Gruyter, Berlin, pp. 583-592.

2.

(a) Mager, H.I.X., R. Addink. 1985 Tetrahedron 4 L 183-190; (b) Mager, H.I.X., S.-C. Tu. 1988. Tetrahedron 44, 5669-5674; (c) Mager, H.I.X., D. Sazou, Y-H Liu, S.-C. Tu, K. M. Kadish. 1988. J. Am. Chem. Soc. U0, 3759-3762.

3.

Merenyi, G„ J. Lind, H. I. X. Mager, S.-C. Tu. 1992. J. Phys. Chem. 96, 1052810533.

4.

(a) Mager, H.I.X., W. Berends. 1974. Tetrahedron 30, 917-927; (b) Mager, H.I.X. 1976. In: Flavins and Flavoproteins (T. P. Singer, ed.) Elsevier, Amsterdam, pp. 23-37; (c) Tu, S.-C., H. I. X. Mager, R. Shao, K. W. Cho, L. Xi. 1991. In: Flavins and Flavoproteins (B. Curti, S. Ronchi, G. Zanetti, eds.) W. de Gruyter, Berlin, pp. 253-260.

CHEMICAL SYNTHESIS OF NE.KOFLAVIN

K. Matsui Department of Home Economics, Osaka International University for Women', Tohda-cho, Moriguchi-shi, Osaka 570, Japan S. Kasai Faculty of Engineering, Osaka City University, Sumiyoshi-ku, Osaka-shi, Osaka 558, Japan

Introduction Nekoflavin (NF) was isolated from cat choroids by Matsui (1). The UV-VIS absorption spectra showed its close similarity to riboflavin (RF) , but precise structure was not yet determined. The 1 H-NMR spectrum of pentaacetate suggested la - or 8a hydroxy structures (7a HORF, 8a HORF, -respectively) (2) , and such compounds were found in human urine (3) . Later the ' HNMR spectrum of NF acetate was found not to be coincident with that of 8a HORF acetate. Therefore, NF was tentatively assigned to be 7a HORF (4) . We synthesized la HORF pentaacetate from o-acetyltoluidine. The structure was confirmed by NMR, mass, UV-VIS absorption spectroscopy and elementary analysis. The comparison of these properties of the two compounds proved the identity of la HORF with NF.

Results The pathway of synthesis is shown in Scheme 1. o-Acetyltoluidine (I) was converted to 4-chloro-2-methylbenzonitrile (IV) by the method of Claus and Stapelberg et al. (5). IV was nitrated to V by the mixed acids. V was treated with Dribitylamine in the presence of pyridine to ribitylaminobenzonitrile compound (VI), which was reduced with hydrazine

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

32 hydrate in the presence of FeCl 3 o-phenylenediamine derivative

and active carbon

(VII).

alloxan

(7) to 7-cyanoflavin

acetate

(IX) was hydrogenated to la -aminoflavin

presence of Ni catalyst.

(VIII), which was acetylated. The

and purified by silica gel column Mp 2 0 0 t •

C,53.26; H,4.96; N,9.29%.

0.25H 2 O:

Calcd. for MH + 01

3

_ "^yy "^YY HjJ-1 J ~~ 'J I II IV " III H,Ç-0H HjC-OH (HO-C-HL (HO-C-H), HX~0H 9"* HjC. .a (HO-C-H), HiyY"Ç1Hj + CHj . NO' NO, NH, ncAANHj Ncr^^-NOj VII VI H2Ç-0H Hrf-OA. (AcO-Ç-H), CHO-Ç-Hlj ÇH, 3 y y "jCyY'Yy — ^ y y ^ y _

"iyv

0

603.56.

HjC-OH (HO-C-H), ?"2 Hi-^VV" I^H, „ 0

Elementary

analysis.

Calcd. for C 2 7 H 3 0 N 4 O,2 -

C,5 3.4 2; H,5.06; N,9.22%.

603.2114.

to la HORF

(XII).

Light yellow fine crystals. Found:

(X) in the

X was treated with HN0 2

(XI), which was acetylated chromatography

(6) to an

VII was condensed with

Mass (FAB) : Found m/Z UV-VIS absorption

in CHCl a

spectra

is shown in Fig. 1.

Though the

s

mM

of NF acetate

was not measured, the wavelengths of absorption peaks of both acetates were perfectly coincident.

The

absorbance of NF acetate in UV region

(e.g. 300nm) was

relatively a little low; this

VIII H£-0H (HO-C-HI, CH, HX-^N^V" OH y. 0

suggests Pflntoac«tat# XII

presence of im-

purities in the preparation.

1

synthesized

H-NMR

spectrum

of the synthesized la HORF acetate is shown in Fig.2.

Scheme 1.

The signals were consistent 0

7aH0RFAc » NFAc

with the structure. The spectrum of the natural NF acetate is shown in Fig.3. The signals of both acetates

£ 20

were coincident except those of 3NH, which were variable. The

C-NMR spectra of both

acetates, not shown here,

•v S00

13

400 Wavelength nm

500

also coincided.

Fig . 1. UV-VIS Absorption spectra of acetates of 7a HORF and NF in CHCI3.

(Shimadzu Spectrophotometer

UV-260)

33 CH3-CO-

7*Hb- -7aHo

I'Hb I'Ho

t 5.5

i i

5'Hb 5'Ho

4H 3'H / 2'H \ / 1 i L

I I 5.0

3NH 1

_r

ui > C 4 1 H Fig. 2. 1 H-NMR Spectrum of la HORF acetate in CDC1 3 . (JEOL ppm

_L

S

10

L

NMR Spectrometer JNM GX-400) CH,-C0r

7dHtr -7dHa

10

ppm

8

6

4

T"

Fig. 3. 'H-NMR Spectrum of NF acetate in CDC1 3 .

34 The data described here proved the structure of NF to be la hydroxyriboflavin.

Acknowledgement The authors thank Prof. S. Kusumoto and Prof. Y. Shimonishi of Osaka University for the elementary analysis and the mass spectrometry, and Prof. F. Mtl lier of Sando Agro Ltd. for discussion and trial of synthesis.

References

1. Matsui,K.1965.J.Biochem.Tokyo

57:201.

2. Matsui,K.,S-Kasai.1984.In : Flavins and Flavoproteins (R.C. Bray,P.C.Engel,and S.G.Mayhew eds.),Walter de Gruyter p.75. 3. Ohkawa,H. ,N.Ohishi,K.Yagi.1983•J•Biol.Chem•258 : 5623. 4. Matsui,K.,S.Kasai.1991.In : Chemistry and Biochemistry of Flavoenzymes (F.Mu lier ed.) ,Vol.2,CRC Press,p.110. 5. Claus,Ad.,E.Stapelberg.1893.Lieblg's

Ann.Chem.274 : 285.

6. Hirashima.T.,O.Manabe.1975.Chemistry

Letters

7. Kuhn,R.,K.Reinemund,F.Weygand,R.Stro Chem.Gesell.68:1765.

be.1935.Ber.Deutsch.

1975:259.

NOVEL 5 - SUBSTITUTED 5 - DEAZAFLAVINS : SYNTHESIS AND APPLICATIONS AS ACTIVE SITE PROBES FOR FLAVOPROTEINS.

Yerramilli V.S.N. Murthy and Vincent Massey Department of Biological Chemistry, Univ. of Michigan, Ann Arbor, Mi-48109, USA.

Introduction To understand the mechanisms which govern flavin reactivity, knowledge of the protein environment at the active centers of different classes of flavoenzymes is necessary. One of the ways to obtain such information is through the replacement of the native flavin with appropriately modified flavins. Although there is a wealth of information about the protein surroundings of various positions through studies with modified flavins, little direct information is available for the important redox-active N(5) center (1). Therefore various flavins, structurally modified at the N(5) position, were designed and successfully synthesized for use as flavoprotein active site probes.

Results and Discussion The flavins that were considered for the present study are shown in Scheme 1. One of the interesting structural features of the flavin ( I I ) is that it could be a good mimic for both a Scheme 1 R

xxxr ii X

ii o

l = X = 0;ll = X = S nucleophilic as well as an electrophilic center at the important N(5) position as a result of its two possible isomeric thioketone and enol forms. Hence it was expected that in addition to providing information on the solvent accessibility to this position, it could also throw some light on mechanistic aspects of flavoprotein reactions by acting as a trap for reactive intermediates that might be formed during the reaction.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

36 It was reported earlier that the 4a-5 epoxide of 5-deaza-flavins can rearrange to 5carbonyl flavins when refluxed in pyridine (2). So the 4a-5- epoxide of our model flavin, 10-ph-5-deaza-flavin, was prepared (3) by reacting with 1.1 molar equivalents of mCPBA in chloroform in presence of NaHC03-Then the epoxide was converted to the corresponding 5-carbonyl flavin by refluxing in pyridine for 2 hours. Interestingly, when the epoxide of 5-deaza-riboflavin was refluxed in pyridine, no rearrangement product was obtained. It was thought that the free hydroxyls in the ribityl side chain might be causing some complications and the epoxide of the corresponding tetra-OAc-ribityl- 5-deaza riboflavin was prepared. This on rearrangement formed the corresponding 5-carbonyl -5deaza-riboflavin in 60% yield (Scheme 2). A simple alternative route was also developed for the 5-carbonyl-5-deaza-flavins by using triphosgene as an excellent one carbon atom reagent. Thus the (N-tetra-OAc-ribityl-xylidino) uracil, on refluxing with 0.5 eq of triphosgene in dichloromethane for 12 hours afforded the flavin ( I ) in 40-50% yield (Scheme 2). When this reaction was carried out in DMF, tetra-OAc-ribityl-5-deazariboflavin was obtained in quantitative yields at room temperature through the Vilsmeir reaction. The ribityl hydroxyl groups were deprotected by treatment with 6N HC1. When the 5-carbonyl flavin was dissolved in buffers or protic organic solvents like alcohols, the spectrum of the flavin collapsed with the loss of the visible absorption. This may be because of the hydration of the a , P - unsaturated carbonyl or due to the formation of the enolate anion of the flavin. Interestingly this was found to be reversible as the complete spectrum of the flavin was restored once it was extracted back into aprotic organic solvents like dichloromethane or chloroform. When 5-carbonyl -5-deaza-riboflavin was titrated with the apoprotein of hen egg white riboflavin - binding protein (RBP) in 0.1M NaOAc, pH 5, large changes in the absorption spectrum occured. With the the binding of the flavin to the protein, the visible absorption changed to one typical of the flavin in aprotic solvent. When the titration was carried out in 0.1M KPi at pH 7, the binding was much weaker. This is consistent with the known preferential binding of neutral flavins over negatively charged ones by this protein. These preliminary results highlight the potential applications of this flavin in understanding the hydrophobic and hydrophilic environments of flavoprotein active sites. It was reported by Yoneda et.al., that the 4a-5-epoxides of 5-deaza flavins can be converted to 5-Cl-5-deaza flavins by reacting with Vilsmeir (POCI3-DMF) reagent (3). But in our hands this reaction, when carried out with 10-ph-5-deaza flavin, failed to yield any isolatable chloroflavin and so we looked for other reagents. The 5-chloroflavin was obtained with thionyl chloride as the chlorinating agent and pyridine as solvent (Scheme 2)

37 Scheme: 2 R I 5- deaza-flavin

mCPBA

Y

II

W \

N

V

V I

T I

°

K'lu

Pyridine Redux

I

O

I R = C6H5 , Tetra - OAc - ribityl , Ribityl

This chloroflavin proved to be extremely reactive and of no use for studies with proteins. However , this extreme reactivity was exploited by using it as an effective precursor for synthesizing various 5-substituted flavins. Two potentially useful 5-deazaflavin derivatives, 5-mercapto and 5-azido flavins were synthesized by reacting the chloroflavin ( I I I ) with aq. solutions of Na2S and NaN3 (Scheme 2; Figs 1 and 2). The 5-azido flavin was found to be photoreactive and stable in aqueous solution in the dark. In line with using these flavins as active site probes, the 5-mercapto flavin was taken up first for detailed study. The 5mercapto-5-deaza riboflavin was synthesized just as for the model, except that the tetraOAc-ribityl-5-Cl-5-deaza riboflavin was not isolated but was converted in situ

to the

mercapto flavin by reacting with aq. Na2S. Finally the ribityl hydroxyls were deprotected by treating with IN NaOH. The p K a values of the mercapto flavin were determined to be at pH 0.5 (flavinium cation) and pH 4.0 (thiol) from the change in absorbance spectrum with pH. The

flavin was found to react with all normal thiol

reagents. It reacts with one

equivalent of methyl methane thiosulfonate to form the 5-SSCH3-deazaflavin, and this reaction is reversible with DTT. The mercaptoflavin on reaction with mCPBA, takes up one equivalent of the oxident to form the S-oxide and further consumes another 2 eq. to form the sulfonic acid derivative of the reduced flavin. The flavin also reacts with one equivalent of iodoacetamide and dimethyl sulfate to give the 5-S-CH2-CO-NH2 and 5-S-

38 CH3 derivatives respectively. It also reacts with two equivalents of sodium cyanide, sodium thiocyanate, sodium sulfite; the spectra suggest the formation of derivatives of the reduced flavin. Attempts to take this flavin to the FAD level by using the

Brevibacterium

FAD synthetase failed. However, FMN was obtained in excellent yields at pH 5.5 in 0.1M NaOAc in this reaction. The 5-mercapto-5-deaza riboflavin was found to bind to apoprotein of hen egg white RBP and the apoproteins of flavodoxin, OYE and L-lactate monooxygenase were reconstituted with the 5-mercapto-5-deaza-FMN.

300

400

500

Wavelength (nm)

600

300

400

500

Wavelength (nm)

600

FIG:2

FIG:1

Fig. 1: Reaction of 10-Ph- 5-Cl-5-deaza-flavin with sodium sulfide. 1) 10-Ph-5-Cl-5-deazaflavin. 2) After adding 2

of 1M sodium sulfide in 0.1 M KPi, pH 7 at 25° C.

Fig. 2 : Reaction of 10-Ph-5-Cl-5-deaza-flavin with sodium azide. ( deaza- flavin; (

) 10-Ph-5-Cl-5-

) After reaction with sodium azide in 0.1 M KPi, pH 7 at 25° C.

Acknowledgements This research was supported by a grant from the U.S. Public Health Service GM-11106. References 1.Ghisla, S and Massey, V. 1986. Biochem. J. 1-12. 2.Yoneda, F and Sakuma, Y. 1981. Tetrahedron Letts. 22, 3977-3980. 3.Vargo,D and Jörns, M.S. 1979.J. Am. Chem. Soc. 7623-7626.

D-Lactate Dehydrogenase Model. Oxidation of a - H y d r o x y Acid by Functionalized Oxidation Active flavin Mimic in the Presence of Zn 2 + and Base in t-Butanol

Y. Yano, K. Mitsui, Y. Ohsawa, T. Nabeshima Department of Chemistry, Gunma University, Kiryu, Gunma 376, Japan

Introduction For design of artificial flavoenzymes, the followings would be of primary importance; (i) the catalytic group is highly active, (ii) the catalytic systems have a substrate-binding site with chirality , and (iii) functional groups are arranged in the proper position. Arrangement of functional groups at the catalytic site would be achieved by covalent and/or noncovalent functionalization.(l) We have successfully exploited oxidation-active flavin mimics by chemical modification of an isoalloxazine ring.(2) Among them, benzo-dipteridine (BDP), which shows ca. lO^'fold rate enhancement for the oxidations proceeding via a nucleophilic attack at C(4a)-position, is quite useful for model studies for flavin mediated oxidations.(3) Meanwhile D-lactate dehydrogenases from various bacterial and mammalian sources, which oxidize D-lactate to pyruvate, are the only flavoproteins to contain Zn^+, although the roles of Zn2+ are not clearly understood.(4) Recently we have found that a covalently functionalized BDP (6-bpy-BDP) is able to oxidize a-hydroxy acid to give the corresponding a—keto acid in the concurrent presence of

and amine base in t-BuOH, whereas 5-bpy-

BDP and Me-BDP are unable to oxidize it under the same conditions. (Scheme 1) Et

Scheme 1 It is helpful to summarize our previous data for this oxidation.(5) (i) The relative rates of PhCH(OH)COOH, PhCH(OH)COOMe, and MeCH(OH)COOH are 20 : 8 : 1. (ii) Deuterium kinetic isotope effect (kH/kD) for PhCD(OH)COOH is 7.2, (iii) a - C - H of

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

40 PhCH(OH)COOH does not exchange with solvent protons, (iv) the rate of the oxidation is not affected by 0 2 , and (v) Me-BDP oxidizes PhCH20" to give PhCHO in P I 1 C H 2 O H . Furthermore, the concentration effects of PhCH(OH)COOH and Z n 2 + on the rates suggest that the oxidation proceeds via a ternary complex of 6-bpy-BDP, Z n 2 + , and PhCH(OH)COO". From these data, the roles of Z n 2 + are considered to be; (i) Z n 2 + improves the oxidation-activity of 6-bpy-BDP due to interaction with C = 0 group of 6-bpyBDP, (ii) Z n 2 + acts as a substrate-binding site, and (iii) Z n 2 + activates the substrate by lowering pKa's of -OH and a - C - H hydrogens.(6) We proposed that the oxidation proceeds through nucleophilic attack of alkoxy anion at C(4a)-position to form C(4a)-adduct followed by base-catalyzed elimination to afford PhCOCOO" and 2e-reduced 6-bpy-BDP as shown in Scheme 2. In this paper, we describe oxidative decarboxylation of a-methyl mandelic acid by 6-bpyBDP and discrimination of the chiral substrate by using a chiral base.

Results and Discussion Reaction of PhCMefOfflCOOH with 6-bpy-BDP. To confirm the alkoxide mechanism, a-methyl mandelic acid which has no a-hydrogen atom was used as the substrate. If our proposed mechanism is correct, C(4a)-adduct would be observed because the successive elimination is unable to occur due to no a-hydrogen. It was found that 6-bpy-BDP reacts with PhCMe(OH)COOH to give 2e-reduced 6-bpy-BDP and PhCOMe in the presence of l,8-diazabicyclo[5.4.0]undec-7-ene (DBU) and N i 2 + in t-

41 BuOH.

Formation of PhCOMe

is accounted for by oxidative decarboxylation of

PhCMe(OH)COO"from the adduct (Scheme 3). The rate constants are shown in Table 1. The rate of the oxidative decarboxylation is much slower that of the elimination. Thus, the decarboxylation is considered to occur due to no a-hydrogen after formation of the adduct, implying the alkoxide mechanism. It is well known that decarboxylation of carboxylic acids bearing electron-withdrawing groups at a-carbon occurs under the similar conditions.(7) Table 1. Pseudo-first-order rate constants Acids PhCH(0H)C0 2 H PhCMe(OH)C0 2 H

k

obs (min"1) 2

1.6 x IO

kiel

190 2

8.4 x IO"

1.0

[6-bpy-BDP]=1.0x 10"5M, [Acid]=5.00 x lO^M [DBU]=1.50 x 10"3M, t-BuOH, N 2 , 25'C

PhCOMe +

2e-Reduced BDP

Scheme 3

Chiral recognition . Chiral recognition of the substrate was kinetically examined by using (+)- and (-)PhCH(OH)COOH with a chiral base ((-)-spalteine) The rate constants

were

spectrophotometrically determined by following the absorption increase of 2e-reduced 6-bpyBDP at 640 nm. The results are shown in Tables 2. The degree of chiral recognition is quite small. Design of the catalystic systems discriminating molecular chilarity is our future problem.

42 Table 2. Effect of (-)-spalteine Acids (+)-PhCH(0H)C0 2 H (-)-PhCH(0H)C0 2 H

IWmin1) 5.56 x 10"2 8.84 x 10"2

k.el 1.0 1.5

[6-bpy-BDP]=1.0 x 10 5 M, [Acid]= 5.00 x 10^M [(-)-spalteine]=5.00 x ÎO^M, [Zn 2+ ]=1.00 x lO^M

Conclusion This is the first example of D-lactate dehydrogenase model, although the oxidation mechanism is different from that of the enzymes.

The roles of

however, are

considered to be similar to those in enzymatic systems.

References

1. Yano. Y., N. Tamura. K. Mitsui, T. Nabeshima. 1989. Chem. Lett. 1655. 2. Yano. Y., M. Ohshima, I. Yatsu, R. E. Vasquez, A. Kitani, K. Sasaki. 1985. J. Chem. Soc. Perkin Trans. 2. 753; Yano. Y., H. Kamishima, S. Sutoh, K. Iizuka. 1986. J. Chem. Res.(S) 382. 3. Yano. Y., M. Ikuta, T. Yokoyama, K. Yoshida. 1987. J. Org. Chem. 52: 5606; Yano. Y., M. Nakazato, K. Iizuka, T. Hoshino, T. Tanaka, K. Koga, F. Yoneda. 1990. J. Chem. Soc. Perkin Trans. 2. 2179. 4. Gregolin. C., T. P. Singer. 1963. Biochem. Biophys. Acta 67: 201; Morpeth, F. F., V. Massey. 1982. Biochemistry 21: 1318; Folsen. S., V. Masey. 1979. Biochemistry 18: 4714; 1980. Ibid. 19: 3137. 5. Yano. Y., K. Mitsui, Y. Ohshima, T. Kobayashi, T. Nabeshima. J. Chem. Soc. Chem. Commun. (Submitted) 6. Gerlt. J. A., J. W. Kozarich, G. L. Kenyon, P. G. Gassman. 1991. J. Am. Chem. Soc. 113:9667. 7. Cram. D. J., P. Haberfield. 1961. J. Am. Chem. Soc. 83: 2354.

Mechanism of Flavin Reduction By Cu(I)-EDTA Complex

Benfang Lei and Shiao-Chun Tu Department of Biochemical and Biophysical Sciences, University of Houston Houston, Texas 77204-5934

Introduction Flavins, in free and enzyme-bound forms, have been extensively studied with respect to redox properties, and mechanisms have been formulated to account for hydride transfer, adduct formation, one and two electron reductions, and one electron superoxidation. Cu(I)EDTA complex has recently been reported to be an effective reductant for flavins (1). In order to understand the electron transfer between Cu ion and flavin, we have now studied the kinetic mechanism of the reduction of FMN by Cu(I)-EDTA using stopped-flow techniques. The formation of one molecule of FMNH 2 produces two molecules of Cu(II)EDTA. The process can be described as a second order reaction of Cu(I)-EDTA and FMN with a rate constant of 3.2 x 105 M _ 1 s _ 1 at 23°C, pH 7.0. A stepwise two one-electron reduction mechanism of FMN is proposed.

Results and Discussion Bleaching of FMN by Cu(I)-EDTA results in the formation of Cu(II)-EDTA which has an absorption peak at 730 nm. The ratio of Cu(II)-EDTA formed over FMN fully reduced was spectrophotometrically determined to be 1.9. Therefore, the two electrons needed to fully reduce one molecule of FMN are each from a Cu(I) ion. The reduction of FMN monitored by stopped-flow tracing of decrease in A 4 4 5 is a pseudofirst-order reaction in the presence of excess Cu(I)-EDTA (Figure 1). The pseudo-first-order rate constant is proportional to the concentration of Cu(I). Cu(I) ion in an aqueous solution undergoes a disproportionation reaction but this process can be retarded by the addition of acetonitrile. However, acetonitrile can form complexes with Cu(I) and such complexes can not reduce flavin. Therefore, increases in the concentration of acetonitrile dramatically

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

44 Fig. 1. Demonstration of pseudo-firstorder reduction of FMN by excess Cu(I)-EDTA. A solution containing 0.02 mM FMN, 150 mM EDTA and 0% acetonitrile, pH 7.0, was mixed with an equal volume of 5 mM Pi, pH 7.0, containing 0.86 mM Cu(I) and 20% (v/v) acetonitrile in a stopped-flow apparatus at 23°C. Solutions were deoxygenated by N 2 bubbling. FMN reduction was monitored by AA445 which was semilogarithmically plotted against time.

10-1

10-2

0.00

0.05

0.10 Time

0.15

0.20

(s)

decrease the reaction rate. To determine the second-order rate constant, EDTA titrations of the reaction were carried out at four levels of Cu(I). The double reciprocal plots of observed pseudo-first-order rate constants versus EDTA concentrations gave linear lines (Fig. 2A). 0.4 Q. FMNHj + Cu(II)-EDTA

> Dimer of semiquinone

(!) (2) (3)

Reaction 1 appears to be the rate-limiting step. In the presence of saturating EDTA and subsaturating acetonitrile, Cu(I) exists as Cu(I)-EDTA and EDTA-Cu(I)-An. The latter can not reduce FMN. Therefore, the rate expression corresponding to the reactions 1 and 2 or 1 and 3 is given by rate = k [FMN] [Cu(I)-EDTA] = k [FMN] [Cu(I)]t / {1 + K [An]}

(4)

l/*app = 1 / k + [ A n l ^

(5)

where K is the association constant of EDTA-Cu(I)-An from Cu(I)-EDTA and An. [Cu(I)] t is the total concentration of Cu(I). Eq. 5 forms the basis for the graphic analysis shown in Fig. 2C.

Acknowledgments This work was supported by grants GM25953 from NIH and E-1030 from The Robert A. Welch Foundation.

References 1.

Lei, B. and J. E. Becvar 1991. Photochem Photobiol. 54, 473-476.

2.

Beinert, H. 1956. J. Am. Chem. Soc. 78, 5323-5328.

SELF-ASSOCIATION OF FLAVIN: INTERACTION OF FMN WITH ALBUMIN

Tetsuo Ishida and Kihachiro Horiike Department of Biochemistry, Shiga University of Medical Science, Ohtsu 520-21, Japan

Introduction Ravin is one of self-associating ligands. We theoretically studied the general features of such ligand-binding systems (1): ligand self-association alone can cause non-hyperbolic binding curves or nonlinear Scatchard plots. In this study we investigated the interaction of FMN with serum albumin, and found an unusual binding behavior of FMN due to its indefinite selfassociation.

Results and Discussion Bovine and human monomelic mercaptalbumins (BS A and HSA) were purified from Fraction V albumin on a DEAE-Sepharose CL-6B column. FMN was purified on a reversed-phase column. Binding data were obtained in 0.138 M sodium phosphate (pH 5.8; ionic strength, 0.16) at 25°C. Under the conditions, albumin exists mainly in N-form (2). The fractional saturation value (r) was calculated with the following equation: r = ([L t ] - [Lf t ]) / [P t ], where [LJ and [ L f J are the total flavin and total free flavin concentrations, respectively, which are expressed in monomer units, and [PJ is the total albumin concentration. Since the initial slope of the binding curve (r vs. [ L f J plot) is nk\, where n is the number of binding sites on the protein, and

is the intrinsic binding constant of the ligand monomer

(1), we first determined the nk\ value by Hummel-Dreyer gel chromatography (3) on a short TSK-GEL G3000SW column (Tosoh; 7.5 x 75 mm) (4) with low [L ft ] (< 40 |IM). The r value increased in proportion to [ L f J in the initial parts of the binding curves. The nk\ value was found to be 761 ± 24 M"1 for BSA and 258 ± 54 M"1 for HSA, indicating that FMN monomer can bind more tightly to BSA than to HSA. Next we obtained the binding curves of FMN in a wide [ L f J range (0.2-5 mM) by the frontal gel chromatography (5) on a Bio-Gel P10 column. The binding curves leveled off at lower levels than saturation (Fig. 1). The

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

48 Scatchard plots were concave-upwards and the apparent r m a x was found to be - 0 . 7 for BSA and - 0 . 3 for HSA. This phenomenon could not be explained by a simple hyperbolic binding mechanism. Taking account of the self-association of FMN (6-8), we analyzed in detail the data for BSA on the basis of our theory (1). 1) Monomer-Dimer Equilibrium Model. From the results of Hummel-Dreyer's method, FMN monomer could bind to albumin, i.e. k\ * 0. A model with n= 1 and ¿2 = 0 (*2> the intrinsic binding constant of the dimer) fitted the data well. When the k\ value was taken as that determined experimentally, the model also fitted the data. However, the calculated association constants of FMN (/sTl2) with both models (432 M_1 and 235 M"1) were much larger than the reported values of 118 M_1 (7) and 140 M_1 (6). In the models in which k\ * 0 and ¿2 * 0, the ¿2 values were calculated to be negative. In the cases where n > 2, the parameters could not be uniquely determined. Hence, we excluded the above models. 2) Isodesmic Indefinite Self-Association Model. In this model, the equations describing the simplest case (« = 1, *i * 0,

= 0, and i > 2) are (1): 2*i [L f J

r = 1 + 2*i [Lft] + 2x-* 1 [L ft ] + (1 + 4AT*1[Lft])l/2

'"max

=

98.5%) is present as the

monomer in the low concentration range ([L t ] < 0.04 mM) in which the nk\ value was obtained by Hummel-Dreyer experiments. On the other hand, at concentrations of more than 0.2 mM, at which the frontal gel chromatographies were performed, oligomers exist in appreciable amounts (Fig. 2). In earlier studies on the interaction of flavin with albumin (10-13), neither the self-association of flavins nor the heterogeneity of albumin (2) was taken into consideration. In this study we prepared bovine monomeric mercaptalbumin and obtained binding data for a wide flavin

50 concentration range (0.01-5 mM). The binding curves leveled off at lower levels than saturation. The deviation of the estimated number of binding sites from an integer has been interpreted as being due to the heterogeneity of a protein and/or a ligand. Ligand selfassociation is one cause of ligand heterogeneity, as shown in our previous paper (1): a binding curve can approach a non-integer value of r m a x due to the effect of ligand selfassociation other than discrete self-association. We showed that the albumin-flavin binding system is a typical example in which ligand selfassociation plays an important role in the binding behavior of the ligand: the binding curve is non-hyperbolic, and the apparent cooperativity appears (1). There are many physiologically important ligands which self-associate (1 and references cited therein). The present study will provide basic information for studies on such ligand-binding systems.

References 1. Ishida, T„ K. Horiike, H. Tojo and M. Nozaki. 1988. J. Theor. Biol. 130, 49-66 2. Peters, T„ Jr. 1985. Adv. Protein Chem. 37, 161-245 3. Hummel, J. P. and W. J. Dreyer. 1962. Biochim. Biophys. Acta 63, 530-532 4. Tojo, H., K. Horiike, T. Ishida, T. Kobayashi, M. Nozaki and M. Okamoto. 1992. J. Chromatogr. 605, 205-213 5. Cooper, P. F. and G. C. Wood. 1968. J. Pharm. Pharmacol. 20, 150S-156S 6. Kharasch, E. D. and R. F. Novak. 1981. Arch. Biochem. Biophys. 212, 20-36 7. Grajek, H., R. Drabent, G. Zurkowska and C. Bojarski. 1984. Biochim. Biophys. Acta 801, 456-460 8. Medina de Gonzalez, M. J. and N. Langerman. 1977. Arch. Biochem. Biophys. 180, 75-81 9. He, X. M. and D. C. Carter. 1992. Nature 358, 209-215 10. Leviton, A. and M. J. Pallansch. 1960. J. Dairy Sci. 43, 1713-1724 11. Jusko, W. J. and G. Levy. 1969. J. Pharm. Sci. 58, 58-62 12. Singh, S. 1981. Indian J. Biochem. Biophys. 18, 236-237 13. Innis, W. S. A., D.B.McCormick and A. H. Merrill, Jr. 1985. Biochem. Med. 34, 151-165

BIOSYNTHESIS OF FLAVINS

STUDIES ON THE BIOSYNTHESIS OF FLAVINS. STRUCTURE AND MECHANISM OF 6,7-DIMETHYL-8-RIBITYLLUMAZINE SYNTHASE

A. Bacher 1 , K. Ritsert 2 , K. Kis 1 , K. Schmidt-Bäse1, R. Huber 2 , R. Ladenstein 3 , J. Scheuring1, S. Weinkauf 1 and M. Cushman 4 department of Chemistry, Technical University of Munich, Lichtenbergstraße 4, D-85747 Garching, Federal Republic of Germany 2

Max-Planck-Institut für Biochemie, D-82152 Martinsried, Federal Republic of Germany

3

Center for Structural Biochemistry, Karolinska Institute, S-14157 Huddinge, Sweden

4

Department of Medicinal Chemistry and Pharmacognosy, Purdue University, West Lafayette, Indiana 47907, USA

The lumazine synthase/riboflavin synthase complex of Bacillus subtilis is a large protein consisting of 60 (8 subunits and 3 a subunits (1, 2). The /3 subunits catalyze the condensation of 5-amino-6-ribitylamino-2,4(lH,3H)-pyrimidinedione (1) with 3,4-dihydroxy-2butanone 4-phosphate (2) yielding 6,7-dimethyl-8-ribityllumazine (3) (Fig. 1) (3, 4). The a subunits catalyze the subsequent dismutation of the lumazine 3 yielding riboflavin and the pyrimidine 1 (1). It should be noted that the pyrimidine 1 is both a substrate of the /3 subunits and also a product of the a subunits. As a consequence, one half of the pyrimidine 1 is regenerated in the second reaction step and is subsequently reutilized by the 0 subunits. The overall stoichiometry implicates the formation of one molecule of riboflavin from one molecule of the pyrimidine 1 and two molecules of the carbohydrate 2. The reaction mechanism of lumazine synthase had not been studied in detail up to now, since the second substrate for the reaction, the novel carbohydrate 2, was elucidated only recently. It has been known for some time that the 0 subunits of the enzyme complex form a capsid with icosahedral 532 symmetry (5, 6). The a subunit trimer occupies the central core of this capsid. The structure of the icosahedral capsid was determined to a resolution of 3.3 A by X-ray analysis of the native enzyme complex (6). No electron density was obtained for the enclosed a subunits which are probably disordered with respect to the crystal lattice.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

54

Results Detailed studies on the reaction catalyzed by the /? subunits required the synthetic preparation of the novel carbohydrate 2. We have therefore prepared both enantiomers of 2 by phosphorylation of L- and D-3,4-dihydroxy-2-butanone with dibenzylchlorophosphate.

0 gtp—

T r HN N 0 CH, H-9-OH H-C-OH H-C-OH CH,0©

CH, H-9-OH H-C-OH 4 H-C-OH CHO ,H Fig. 1: Biosynthesis of riboflavin and reactions catalyzed by the lumazine synthase/riboflavin synthase complex. A, 6,7-dimethyl-8-ribityllumazine synthase; B, riboflavin synthase.

Using the synthetic carbohydrate, it was shown that the o t ^ 6 0 complex catalyzes the formation of the lumazine 3 from 1 and L-3,4-dihydroxy-2-butanone 4-phosphate (2) with a velocity of v m a x = 12 jumol mg"1 h~>. The K M value for the carbohydrate substrate is about 130 /iM. The D-enantiomer of the carbohydrate can also serve as a substrate, but with a significantly reduced reaction velocity. The K ^ for the pyrimidine 1 has a value of

The native a^P^Q enzyme complex can be dissociated under mild conditions yielding tri-

55 mers of a subunits and large aggregates of 18 subunits with the shape of hollow spheres and a molecular weight of several million Da (5). The isolated subunits catalyze the enzymatic formation of the lumazine 3, and the catalytic rate (v m a x = 12 ftmol mg"1 h"1) is virtually identical with that of the native a 3 0 6 O complex. In order to study the regiospecificity of the enzyme reaction, [l-13C]-L-3,4-dihydroxy-2butanone 4-phosphate was prepared enzymatically from [l- 13 C]ribulose 5-phosphate by the action of 3,4-dihydroxy-2-butanone 4-phosphate synthase. The reaction product was used as a substrate for riboflavin synthase, and the resulting riboflavin was analyzed by NMR spectroscopy. The data show that the reaction is strictly regiospecific. This finding and other data not described in detail show that the enzyme does not serve as a diacetyl synthase. A hypothetical reaction mechanism based on these observation is shown in Fig. 2 (3). We propose that the initial reaction step is the formation of a Schiff base by reaction of the carbonyl groups of 2 with the 5-amino group of 1. This intermediate could then eliminate phosphate under formation of the enol structure 5 which could subsequently cyclize to yield the lumazine chromophore. ©o

HN i

n h

n h

r * jV — A -v- A o

0©

2

CH, H H-C-OH H-C-OH H-C-OH CH,0H

1

0

CH, H-C-OH H-C-OH 3 H-C-OH CH,OH

H0

if'

0

CH, H H-C-OH H-C-OH 4 H-C-OH CH 2 0H

1

ho'^y'

0

CH, H H-C-OH H-C-OH H-C-OH CHJOH

0

0

CH, H H-C-OH H-C-OH H-C-OH CH,OH

CH, H H-C-OH H-C-OH H-C-OH CH,OH

6

5

Fig. 2: Hypothetical reaction mechanism of 6,7-dimethyl-8-ribityllumazine synthase. Cushman and coworkers synthesized the trifluoromethyl derivative 7 (Fig. 3) which is a close analog of the hydrated intermediate 6 in the enzyme mechanism (7). The lumazine 7

56 has a chiral center at C-7 and forms two stable diastereomers designated epimer A and epimer B. The interaction of both epimers with the lumazine synthase/riboflavin synthase complex has been studied by

19

F NMR spectroscopy. The spectra (Fig. 3) showed remark-

able differences between the diastereomers with respect to the chemical shift of the enzyme bound species. In the case of epimer A, the chemical shift difference between the signals of the position 7 trifluoromethyl group in the free and enzyme-bound state is about 1 ppm. By contrast, the chemical shift of the position 7 trifluoromethyl group of epimer B is shifted downfield by 9 ppm in the bound state by comparison with the free state. These results imply that substituents on C-7 of the lumazine are very sensitive to the protein environment. Lumazine synthase shows little, if any stereoselectivity respect to the hydrated C-7 of 7, whereas riboflavin synthase has been shown earlier to bind epimer A with a very high degree of stereospecificity (7). Both epimers of 7 can eliminate the 7-CF3 group by formation of 6-trifluoromethyl-7-oxo8-ribityllumazine. Surprisingly, it was observed that this reaction was catalyzed by the lumazine synthase stereospecifically for epimer A but not for epimer B.

B 0 F J C N N O CH2 h H-C-OH H-C-OH H-C-OH CHjOH

7

15

10

11 1 1 ' 1 5 0 PPM

'I

-5

1111

I

1

-10

Fig. 3: a) 1 9 F NMR spectra of 7 in the presence of the lumazine synthase/riboflavin synthase complex, a, epimer A; b, epimer B; F, free ligand; B, bound ligand; x, impurities.

Earlier X-ray studies performed with the a ^ e o enzyme complex had revealed the structure of the capsid at a resolution of 3.3 A (6). Since the a subunit trimer present in the center

57 of this molecule does not follow the icosahedral 532 symmetry, its presence could be responsible for the relatively low resolution obtained in these experiments. We have therefore crystallized reconstituted hollow, icosahedral /36o capsids from isolated j3 subunits. The reaggregation requires the presence of an appropriate substrate analog such as 5-nitro6-ribitylamino-2,4(lH,3H)-pyrimidinedione (8, Fig. 8). The artefactual species could be crystallized in three different space groups (8). The monoclinic modification showed X-ray diffraction extending to a resolution of 2.4 A. The interpretation of the X-ray diffraction data was supported by the freeze-fracture electron microscopy which helped to establish the translational and rotational position of the molecules in the monoclinic cell (9). The monoclinic crystal structure was solved by molecular replacement followed by cyclic phase extension. The molecular model could be refined to a resolution of 2.4 A using the 30-fold averaged electron density. The general architecture of the capsid can be described briefly as follows. The 0 subunit consisting of 154 amino acid residues folds into an motif which is repeated four times. The central, parallel (3 sheet is flanked on both sides by 2 pairs of a helices (Fig. 4).

Fig. 4: Ribbon model of the main chain folding of the /3 subunit. The icosahedral capsid can be described as an assembly of 12 /3 subunit pentamers (Figs. 5, 6). Within the pentamer, the adjacent subunits form extensive contact areas. Moreover, the N-terminus of each subunit interacts with the adjacent subunit by acting as an additional strand to the central /3 sheet.

58 Fig. 5: Space-filling model of the icosahedral jSgo caP~ sid. The subunits of one penton are shaded. The viewing direction is parallel to one of the 3-fold symmetry axes. Selected amino acid residues are marked to indicate the sites where 5fold and 2-fold symmetry axes penetrate the molecular surface.

Fig. 6: C a model of the icosahedral (360 capsid. The model is cut open to show the internal cavity. The viewing direction is parallel to one of the 5-fold symmetry axes.

59 The active site of lumazine synthase is clearly defined in the X-ray structure by cocrystallization of the substrate analog, 5-nitro-6-ribitylamino-2,4(lH,3H)-pyrimidinedione (8). The compound is bound at the interface between adjacent subunits in the pentameric substructure described above. The 60 equivalent substrate binding sites are close to the inner surface of the capsid (6). The ribityl side chain is bound in an extended conformation. The aromatic ring of Phe 22 is closely adjacent and virtually parallel to the pyrimidine ring of the substrate analog (Fig. 7).

Fig. 7: Active site of lumazine synthase. The substrate analog 5-nitro-6-ribitylamino2,4(lH,3H)-pyrimidinedione (8) is shown at the center with the pyrimidine ring in an almost horizontal position. The phenyl ring of Phe 22 is shown on top of the pyrimidine ring.

A spherical volume of high electron density not pertaining to the peptide structure is located in close proximity of the pyrimidine substrate (Fig. 8). This electron density probably represents a fixed inorganic phosphate ion interacting with three highly conserved basic amino acid residues. It appears likely that this site can accomodate the phosphoric acid residue of the second enzyme substrate, 3,4-dihydroxy-2-butanone 4-phosphate (2, Fig. 1). Computer modelling showed that the distance between the phosphate binding site and the

60 pyrimidine substrate is appropriate to accomodate the hypothetical reaction pathway intermediate 4 (Figs. 2, 8). With the intermediate bound in this hypothetical conformation, the highly conserved His 88 residue may serve to abstract a proton from the intermediate, thus initiating the elimination of the phosphate residue which is conducive to the final steps of the reaction mechanism. His 88 has a high degree of mobility as shown by the temperature factor obtained in the X-ray structure analysis. The hypothetical conformation shown in Fig. 8 would satisfactorily explain that both stereoisomers of 3,4-dihydroxy-2-butanone 4phosphate can serve as substrates. These hypothesis can now be tested by site-directed mutagenesis.

Fig. 8: The active site of 6,7-dimethyl-8-ribityllumazine synthase. The electron density at the center of the image represents bound 5-nitro-2,4(lH,3H)-pyrimidinedione (8), a close structural analog of the pyrimidine 1. The hypothetical reaction pathway intermediate 5 (Fig. 2) was modelled with the carbohydrate phosphate side chain in an extended configuration. The phosphate residue of this hypothetical molecule was placed into the electron density corresponding to an inorganic phosphate ion in the experimentally determined electron density. His 88 shown at the bottom is in a favorable position to abstrac a proton from the Schiff base, thus initiating the elimination of phosphate.

61 The sequences for putative lumazine synthase genes of Escherichia coli and Photobacterium leiognathi have been reported recently (10, 11). Sequence comparison shows that virtually all amino acid residues in the neighbourhood of the bound ligand are highly conserved. The remarkable structure of the a ^ e o complex is critical for the interpretation of the unusual kinetic properties. It has been mentioned already that the lumazine 3 is the product of the jS subunit and the substrate of the a subunit, and that the pyrimidine 1 is both a product of the a subunits and a substrate of the 0 subunits (Fig. 1). In the native a ^ 6 0 complex, the central cavity seen in Fig. 6 accomodates a trimer of a subunits. Obviously, the substrate must enter into the icosahedral capsid in order to bind at the active sites of the /3 subunits as well as the a subunits. Moreover, the products must exit from the complex. This process is as yet not understood. Channels running along the 5-fold symmetry axes of the icosahedral capsid would be sufficient to allow the passage of the substrate 1 and 2. However, the width of these channels appears insufficient for the product, riboflavin. Dynamic properties of the large protein may be involved in product passage through the icosahedral capsid.

50

0

0

10

20

30

40

50

60

time

(min)

70

80

90

100

110

Fig. 9: Product formation from 5-amino-6-ribitylamino-2,4(lH,3H)-pyrimidinedione (1) and 3,4-dihydroxy-2-butanone 4-phosphate (2) by the native complex. O, riboflavin (4); • , transient 6,7-dimethyl-8-ribityllumazine (3).

62 Fig. 9 shows the formation of riboflavin from 1 and 2 by the a^ß^o complex. In the initial phase, the formation of riboflavin is accompanied by the transient formation of the lumazine intermediate 3. The velocity of riboflavin formation shows a marked decrease after the consumption of 1 and 2 (at about 10 min). Riboflavin formation from the transient 3 proceeds at an about 30-fold slower rate as compared to the initial reaction phase. These features can be explained by compartmentalization of the reaction inside the capsid. The intermediates 1 and 3 can diffuse from the active site of the a subunit to the active site of the ß subunit and vice versa. Alternatively, a fraction of the lumazine can leave the capsid and thus build up the transient lumazine concentration in the bulk solution. The specific architecture of the enzyme complex thus allows highly efficient substrate channeling with a substantial gain of catalytic performance. This work was supported by grants from the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. References 1.

Bacher, A., R. Baur, V. Eggers, H. Härders, M. K. Otto and H. Schnepple. 1980. J. Biol. Chem. 255, 632-637.

2.

Bacher, A., H. Schnepple, B. Mailänder, M. H. Otto and Y. Ben-Shaul. 1980. In: Flavins and Flavoproteins (Yagi K., T. Yamano, eds.), Japan Scientific Societies Press, Tokyo, p. 579.

3.

Volk, R. and A. Bacher. 1988. J. Amer. Chem. Soc. 110, 3651-3653.

4.

Ladenstein, R., B. Meyer, R. Huber, H. Labischinski, K. Bartels, H. D. Bartunik, L. Bachmann, H. C. Ludwig and A. Bacher. 1986. J. Mol. Biol. 187, 87-100.

5.

Bacher, A., H. C. Ludwig, H. Schnepple and Y. Ben-Shaul. 1986. J. Mol. Biol. 187, 75-86.

6.

Ladenstein, R., M. Schneider, R. Huber, H. D. Bartunik, K. Wilson, K. Schott and A. Bacher. 1988. J. Mol. Biol. 203, 1045-1070.

7.

Cushman, M., D. A. Patrick, A. Bacher and J. Scheuring. 1991. J. Org. Chem. 56, 4603-4608.

8.

Schott, K., R. Ladenstein, A. König and A. Bacher. 1990. J. Biol. Chem. 265, 12686-12689.

9.

Bacher, A., S. Weinkauf, L. Bachmann, K. Ritsert, W. Baumeister, R. Huber and R. Ladenstein. 1992. J. Mol. Biol. 225, 1065-1073.

10.

Taura, T., C. Ueguchi, K. Shiba and K. Ito. 1992. Molec. Gen. Genet. 234, 429432.

11.

Lee, L. Y. and E. A. Meighen. 1992. Biochem. Biophys. Res. Commun. 186, 690697.

BIOSYNTHESIS OF RIBOFLAVIN. CLONING, SEQUENCING, MAPPING, AND HYPEREXPRESSION OF THE GENES ribA CODING FOR GTP CYCLOHYDROLASE II AND ribC CODING FOR RIBOFLAVIN SYNTHASE OF ESCHERICHIA COLL

S. Eberhardt, G. Richter, H. Ritz, J. Brandt, A. Bacher Department of Chemistry, Technical University of Munich, Lichtenbergstraße 4, D-85747 Garching, Federal Republic of Germany

GTP cyclohydrolase II and riboflavin synthase catalyze the first and the last step respectively, in the pathway of riboflavin biosynthesis, respectively (Fig. 1). The reaction catalyzed by GTP cyclohydrolase involves the release of formate and pyrophosphate from GTP (1) and yields 2,5-diamino-6-ribosylamino-4(3H)-pyrimidinedione 5'-phosphate (2). Riboflavin synthase catalyzes the dismutation of 6,7-dimethyl-8-ribityllumazine (3) resulting in the formation of riboflavin (4) and 5-amino-6-ribitylamino-2,4(lH,3H)-pyrimidinedione.

"'"x'a"

H

0

NH

* - T^

CH, H-C-OH H-C-OH H-C-OH CH,OH

3

CH, H-C-OH H C-OH H-C-OH CH,OH

4

Fig. 1: Biosynthesis of riboflavin. The gene coding for GTP cyclohydrolase II of Escherichia coli was cloned on a 3 kb fragment from an EcoKl gene bank by a marker rescue strategy using a riboflavin mutant of E. coli. The gene and the surrounding region were sequenced. The gene was mapped to a position of 1356 kb on the physical map of the E. coli genome (1) where it is located between the aconitase gene (acn) and a gene coding for a membrane bound phosphatase (pgpB). The GTP cyclohydrolase II gene is identical with the gene ribA reported previously. The open reading frame codes for a predicted protein sequence of 196 amino acids with a molecular mass of 21.8 kDa. It shows homology to the 3'-part of the open reading frame II of the /ia-operon from Photobacterium leiognathi (2) and to the 3'-part of the central open rea-

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

64 ding frame of the Bacillus subtilis riboflavin biosynthesis operon (3) (Fig. 2). The protein from B. subtilis has been purified and was shown to catalyze the formation of 2 from GTP (4). ARRPQLE ARVPEL

I F A E K H G L K L G T I E I AKKHQLKM

I A D L I EYRTQQESH

I T I KDL

1ER

S E Y E L N T[Ë]Y G I F T L V T Y R D T

I Q Y R Y N L T T L V[E R

T[F|K v Y[G]Y T N E v O|G|K E|H V A | F | V |

M O L K]R

DJFJL M V[G|F(Ë]E L A T W H D H V A L | V |

G|E I |Q - A K A A T J L V R V H|V K P T L K|D

L 0 V G L S QW S -

G D|V"P F[G|E E I P V L V R V H S E C L T G D

F G I S H R C D C G P|Q L L[H 0 A A L1N|Q I A|A E G R G V L L Y I L I R Q E G RIG

G D IL— S O H T P V

LIÂIR

VHSECLTGD

- [ L :EA A|M Q R] I |Q T E|D —[G V L|V I I S|Q|Q E S P

:aalTO I AEE G R G[7|L

L F|S L R C D C G F|Q L E

P|N S P H S G

i v u aR NI I I C

L Y H R Q E G R N

1

K T | L F EJK LID M|Y A - K|E Q|

L G[SJQ I L A D L G V K K I [R L L

N|S N O G Y

2

1 G L I [N K LJK[A Y|KJL Q E Q G Y D T V E A N E A L G F L P[D]L R N|Y|G[T|G A 0 I L|R|O L G V R N M*K]L L

N NPIRIKII

3

1 G L L | N K [ I R|A Y A L Q[D]Q G Y D T V E A N

L G F A AIDIE R|D F T L C|A|D M F K L I L G V N E V|R L L

N N P K K V

G L E V V E|Y I Y D

1

R A L

2

A G L E[GJY G | J S [ Ï | S ] E R V P L

M E A K E H(N]K K|Y L|Q|T K|M N[K]L G H L L H F

3

E 1 L T E A GÎI N I I V E R V P L

V G R N P N|N]E H|Y L|D|T K|A E U M G H L L N K

Fig. 2: Alignment of the predicted amino acid sequence of GTP cyclohydrolase II from E. coli and homologues proteins from other microorganisms. 1, C-terminal part of the product of open reading frame II of the /wx-operon from P. leiognathv, 2, C-terminal part of open reading frame 3 of theriZ>-operonfrom B. subtilis; 3, GTP cyclohydrolase II from E. coli. The ribA gene was expressed under T5 promoter and lac operator control in plasmid pECH2. GTP cyclohydrolase II represents about 50 % of the protein in cell extracts of E. coli M15 harbouring pECH2. The protein was purified to apparent homogenity by a single column chromatography step on DEAE cellulose. The gene coding for riboflavin synthase (ribC) was cloned by screening a Sau3Al gene bank of E. coli in a strain lacking riboflavin synthase activity (E. coli strain BSV13 obtained from B. Bachmann) (5). The metabolic defect of this mutant was complemented by a plasmid carrying a 6 kb insert. The entire fragment was sequenced by primer walk strategy. Several potential open reading frames were found. One ORF of 639 basepairs shows homology to the riboflavin synthase gene of B. subtilis.

65 Genes, coding for cyclopropane fatty acid synthase (cfa) (6), for a putative DNA-binding regulatory protein and for a purine nucleotide synthesis repressor protein (purR) (7) are located downstream of the ribC gene. Restriction fragment anaysis mapped the gene to a position of 1757 kbp (36.2 min) on the physical map of the E. coli chromosome (8). This result does not correspond to earlier mappings at 50 to 70 minutes (5) and 56 to 59 minutes (9). The ribC gene codes for a protein of 213 amino acids with a molecular mass of 23.4 kDa. The predicted protein sequence shows approximately 25 % homology to the riboflavin synthases of Bacillus subtilis (3), Photobacterium leiognathi (2) and Saccharomyces cerevisiae (10) (Fig. 3). MF T G I V Q G T A K L V S I 0 R K P N F R T H V V E L P 0 H M L D G L E T gIAJSJVJA H N E S M K K A G H A M A L T I K C - S K I |L]E VH L G D S 1 AV N MF T G I E E TGT | U F T G I E S 1 GN I G A I I R H N E D L S I V V N T N N L D I S VN I G D S 1 AT N M F T G I E CM G T V L E N N P Y D D S E S G G Q G V S I T I G N A G S I 0 T CH V G D S 1 AV N

1 2 3 4

N G N H V S FD L U K T K N O F T V D VMP L P S GYT A D L S L NNDSFKVGISP

G DW VN|V]E R A A K F S 0 E 1 E TL R I TN L G D L K V ET V K A T S L N D L T K G S K V N L E R A MAANG R F E T Y K R T A F H S Y R 1 G 0 E V N L E@A ML P T T R L E T I K R S N V A S W 1 0G T Q V N L E R A V S Q D VR F

1 2 3 4

S E W N R Q I W F K V Q [ D ] S Q L 14 K Y I LYK K S|N]A V Y Y 0 L K M -|D]P S L T K T L V L K N G R A I N I W V A V — P V Q |L]K K Q L S E K E G 0 S I I F G F Q L R[D]Q E Y F|K]Y I V EK

1 2 3 4

l g k | k | k LjgIa r v n T e ] i I d I p q t q a v v d t v e r v l a a r e n a m n q p g t e a F S e|kJt[| cjs K V N I ECDM I G K Y M Y R F L H K A N E N K T Q Q T I T K A F L S E N G F LVNINIfflKKVN0E I DMMANYLEKL I K V D R Y E S E K T S N V S M D L E R Y G F MP L|K1K|I o ] ° E|VN I E|V|P|L T G K I I E K Q I L L T L E N Q I S K K D S T L N T M I S N I

GGH GGH GGH GGH

GC c l t v G 1C L T GV C L T GV C L T

t I e I V T|D F V&K L V T]E F

VAKILT V S G H V DG T[A E I T R I E E V S G H V D G V GIE V I E F K R v|Q1G H V D T V A N T V S R R P

G F[T|G|| D G I S L T|VG E 0 T P T R F C V H L|IP[EJT|L G S[IJT V]D G V S L T I|F C L T E D T VT I S L [ I P H T I GS V T l l D G I S L T I N a | v | y Q N V I K L T I vlp H T L G f | T | C | i o o | t | s l t i | i k | v ] d p l s q g g a f y i s m[7|k]h t | o

E R[T]T SET I A E TN DNV I

I S IEEKVRNYLN

Fig. 3: Alignment of the predicted peptide sequences of riboflavin synthases from different microorganisms. Identical residues are boxed. 1, Escherichia coli; 2, Bacillus subtilis", 3, Photobacterium leiognathi; 4, Saccharomyces cerevisiae. 39 Residues are identical in all four proteins. The yeast gene contains two inserts of 6 and 4 amino acid residues which are absent in the bacterial enzymes. The N-terminal motif MFTGI is strictly conserved. At the C-termini the sequences do not show homology and differ in length. All the amino acid sequences show the internal homology noted earlier for the riboflavin synthase of Bacillus subtilis. In the total alignment of all N-terminal and C-

66 terminal halfs, 8 amino acids are identical. Three of them form the motiv VNXE (marked by asterisks in Fig. 3), which earlier was proposed to code for the binding site for 6,7-dimethyl-8-ribityllumazine (11). The ribC gene has been overexpressed under T5 promoter and lac operator control to a riboflavin synthase level of approximatively 30 % of total cellular protein. The protein was purified to apparent homogeneity by ion exchange chromatography.

Acknowledgments We thank Drs. T. Werner and W. Gimbel for help with DNA sequencing. This work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie.

References 1.

Richter, G., H. Ritz, G. Katzenmeier, R. Volk, A. Kohnle, F. Lottspeich, D. Alendorf and A. Bacher. 1993. J. Bacteriol. 175, 4045-4051.

2.

Lee, C. Y. and E. A. Meighen. 1992. Biochem. Biophys. Res. Commun. 186, 690697.

3.

Perkins, J. B., J. G. Pero and A. Sloma. 1991. European Patent Application # 0405370.

4.

Boretskii, Y. R., Y. S. Skoblov, O. M. Khodova and P. M. Rabinovich. 1992. Biochemistry-Russia 57, 702-710.

5.

Bandrin, S. V., P. M. Rabinovich and A. I. Stepanov. 1983. Genetika 19, 14191425.

6.

Wang, A.-Y., D. W. Grogan and J. E. Cronan. 1992. Biochemistry 31, 1102011028.

7.

Kilstrup, M., L. M. Meng and J. Neuhard. 1989. J. Bacteriol. 171, 2124-2127.

8.

Kohara, Y., K. Okiyama and K. Isono. 1987. Cell 50, 495-508.

9.

Teslyar, E. and G. M. Shavlovski. 1983. Zitologia i Genetika 5, 54-56.

10.

Doiguar, F., N. Biteau, M. Crouzet and M. Aigle. 1993. Yeast 9, 189-199.

11.

Schott, K., J. Kellermann and A. Bacher. 1990. J. Biol. Chem. 265, 4204-4209.

BIOSYNTHESIS OF RIBOFLAVIN: ENZYMATIC FORMATION OF 6,7-DIMETHYL-8-RIBITYLLUMAZINE

IN Saccharomyces

cerevisiae

J.J. García-Ramírez, M.A. Santos, J.L. Revuelta Departamento de Microbiología y Genética, Universidad de Salamanca Edificio Interdepartamental, Avda. Campo Charro s/n, 37007 Salamanca, Spain

Introduction The immediate precursor of riboflavin, 6,7-dimethyl-8-ribityllumazine (DMRL), is synthesized from 5-amino-6-ribitylamino-2,4-(lH,3H)-pirimidinedione (ARAP) and ribulose-5-phosphate

in two enzymatic reactions. First, 3,4-dihydroxi-2-butanone-4-

phosphate (DHBP) is formed from ribulose-5-phosphate by elimination of carbon 4 in a reaction catalyzed by DHBP-synthase (1). Then, DMRL-synthase catalyses the condensation of one molecule of DHBP and one molecule of ARAP yielding one molecule of DMRL (2). Several mutants of Saccharomyces cerevisiae which are defective in riboflavin biosynthesis have been isolated (3,4). In particular, riboflavin auxotrophs defective in DHBP synthase activity, rib3 mutants, or DMRL-synthase activity, rib4 mutants, have been characterized (4). We have cloned the RIB3

and RIB4

genes by functional complementation of the

corresponding mutants. The sequence analysis of an r/W-complementing DNA fragment revealed the presence of a single open reading frame of 624 bp(4). This open reading frame could potentially encode a protein of 208 amino acids with a predicted molecular weight of 22400 Da, a value that is consistent with what has been observed for purified Candida guilliermondii

DHBP-synthase (5). Similarly, a single open reading frame of 507 bp was

found within a 2.1 kb Bgltt-Hpal fragment which complements an rib4 mutation. The RIB4 open reading frame predicts the synthesis of a protein of 169 amino acids (molecular mass = 18600 Da.) which is highly homologous to the Bacillus subtilis DMRL-synthase (6). Both these genes are expressed at very low level so the purification of DHBP synthase or DMRLsynthase from a wild-type strain proved to be difficult. As an aid to our efforts to study the molecular and kinetic properties of these enzymes, the coding sequences of both genes have been cloned into constitutive expression vectors using an expression cassette polymerase chain reaction technique. Here we report the overexpression, purification and molecular characterization of DHBP synthase and DMRL-synthase from S. cerevisiae.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

68 Results and Discussion The construct pJR414 was designed to allow overexpression of DHBP synthase yeast (Fig. 1). A 700 bp long DNA fragment containing the RIB3 coding and terminator sequences was fused to the PGK (3-phosphoglycerate kinase) promoter sequences using a polymerase chain reaction based technique and properly designed primers. The pPGK-RIB3 gene fusion was then cloned into the episomic vector YEp352 and the resulting expression vector transformed into the null allele rib3 mutant strain AJ71. An abundant 24 kDa protein was detected in SDS-PAGE of crude cell extracts of AJ71/pJR414 cells which was absent in extracts of AJ71 or AJ71/YEp352 cells. Western blot analysis using polyclonal antibodies specific to DHBP synthase revealed that the 24 kDa protein reacts specifically with the antibodies and, therefore, corresponds to overexpressed DHBP synthase.

Figure 1. Schematic maps of DHBP-synthase, pJR414, and DMRL-synthase, pJR627, expression vectors. Purification was carried out in six steps (Table 1). AJ71/pJR414 was cultured in uracildeficient minimal medium with 2% glucose. Cells were collected by centrifugation, diluted in a 1: 2.5 (mass/volume) ratio in 20 mM-Tris-HCl buffer, pH 7.5 and glass beads were added. The cell suspension was then disintegrated into a Bead Beater (Biospec Products) and the crude extract centrifuged for 20 min at 35,000 x g. Thereafter the supernatant was ultracentrifuged for 60 min at 100.000 x g. After a selective precipitation step with 37.5 % methyl alcohol the membrane-free supernatant was centrifuged for 20 min at 40,000x g. and the supernatant containing DHBP synthase activity was recovered, liophilized, resuspended in a small volume of 20 mM Tris-HCl buffer, pH 7.5 (starting buffer), and extensively dialysed against the same buffer. The dialysed supernatant was then applied onto a prepacked cation-exchange FPLC column (MonoQ, Pharmacia LKB Biotechnology Inc.), previously equilibrated with starting buffer. The column was washed with starting buffer for 5 min (5 ml). Protein was eluted from the column at a flow rate of 1 ml/min with an increasing salt

69 gradient (25 min, 0-500 mM Na CI) Fractions reacting with specific antibodies were pooled and concentrated by liophylization. Finally, DHBP-synthase was purified to apparent homogeneity after passage through two prepacked gel filtration FPLC columns, SuperDex 200 and SuperDex 75 (Pharmacia LKB Biotechnology Inc.) using 20 mM Tris-HCl, 90 mM NaCl, pH 7.5 as elution buffer. Table 1. Purification of DHBP-Synthase from a S. cerevisiae Overproducing the Enzyme. Procedure Cell extract Clarified extract Methyl alcohol MonoQ SuperDex 200 SuperDex75

Volume (ml) 12 10 5 3 1 1

Activity (nmol DMRL/h) 8344 2388 2303 747 381 143

Protein (mg) 186 52 28 2 5 x 10-1 62 x 10-3

Strain (AJ71/pJR414)

Specific activity (nmol DMRIVmg h) 45 46 84 415 762 2293

A molecular mass of approximately 24 kDa was calculated for the purified enzyme by SDSPAGE and by gel filtration chromatography. These data indicate that DHBP-synthase is a monomer. To obtain large amounts of yeast DMRL synthase the expression vector pJR627 was constructed. In this YEp352-based vector the coding region of RIB4 is placed under the control of the strong constitutive promoter of the TEF1 (transcription elongation factor la) gene (Figure 1). As expected, pJR627 specifically directs the synthesis of a single prominent 18 kDa protein in the transformant cells. This protein, which amounted to only 0.001% of the total protein in wild-type cells, accounted for about 2% of the total protein in the transform ants. DMRL-synthase was purified from the overexpressing strain AJ106/pJR627 by the following procedure: the complete cell extract,in 20 mM Tris-HCL buffer, pH 7.5, was clarified by ultracentrifugation at 100,000 xg during 60 minutes. After a selective precipitation with 30% methyl alcohol, the extract was applied to a cation-exchange FPLC column ,MonoQ, previously equilibrated with 20 mM Tris, pH 7.5. The elution of the protein was done with a linear salt gradient (0-500 mM NaCl), monitoring the DMRL synthase activity. Positive fractions, eluting at 300 mM NaCl, were pooled, concentrated and applied to a gel filtration FPLC column,Superdex200, using a 20 mM Tris-HCl, 90 mM NaCl buffer, pH 7.5. Fractions with pure DMRL synthase were selected.

70 SDS-PAGE analyses of the purified enzyme revealed a single silver-stained band with a molecular mass of approximately 18 kDa. Gel filtration chromatography of purified DMRL synthase resulted in a single activity peak. When the elution position was correlated with molecular mass, a value of approximately 95 kDa was obtained. These data indicate that DMRL-synthase is a pentamer consisting of five identical subunits. This organization resembles that of heavy riboflavin synthase of B. subtilis. This enzyme is organized in an icosahedral capsid-like structure. During the folding of the icosahedral capsid a pentamer is the most probable intermediate because it exhibits the most extended interfaces (7). Table 2. Purification of DMRL-Synthase from a S. cerevisiae Overproducing the Enzyme. Procedure Cell extract Clarified extract Methyl alcohol MonoQ SuperDex 200

Volume (ml)

Activity (nmol DMRL/h)

12 10 5 3 1

18699 7993 3260 928 752

Protein (mg) 276 89 31 12 x 10-1 93 x 10-3

Strain (AJ106/pJR627)

Specific activity (nmol DMRL/mg h) 68 90 117 774 8089

Acknowledgements This research was supported by BASF Aktiengesellschaft and the Comision Interministerial de Ciencia y Tecnología of Spain (grant BI092-0036).

References 1. Volk, R. and A. Bacher. 1990. J. Biol. Chem. 265:19479. 3. Oltmanns, O., A, Bacher, F. Lingens and F.K. Zimmermann. 1969. Mol. Gen. Genet 105:306. 4. García-Ramírez, J.J., M.A. Santos, M.and J.L. Revuelta. 1993. (in preparation) 5. Volk, R. and A. Bacher. 1990. J. Biol. Chem. 265:19479-19485. 6. Mironov, V.N., M.L. Chikindas, A.S. Kraev, A.I. Stepanov and K.G. Skryabin. 1989. Mol. Biol. (Moskow) 312:237. 7. Ladenstein, R., M. Schneider, R. Huber, H.D. Bartunik, K. Wilson, K. Schott and A. Bacher. 1988. J. Mol. Biol. 203:1045-1070.

ELECTRON MICROSCOPIC STUDIES ON THE LUMAZINE SYNTHASE/ RIBOFLAVIN SYNTHASE COMPLEX OF BACILLUS SUBTILIS

S. Weinkauf, J. Brandt and A. Bacher Department of Chemistry, Technical University of Munich, Lichtenbergstraße 4, D-85747 Garching, Federal Republic of Germany

The lumazine synthase/riboflavin synthase of Bacillus subtilis is a Afunctional enzyme complex which catalyzes the final reactions in the biosynthesis of riboflavin (1). The protein consists of a trimer of a subunits which is enclosed by a spherical capsid of 60 (3 subunits arranged in icosahedral symmetry (1). Artefactual hollow /3go capsids can be obtained by ligand-driven renaturation of dissociated /3 subunits (1) and have been shown to be catalytically active. The native a ^ 6 0 complex and the reconstituted capsids have been crystallized in several modifications (2,3) which were studied by X-ray crystallography and by electron microscopy. In an earlier study, the application of freeze-etching to distinguish between different packing models of the hexagonal crystal modification of the «3^60 species facilitated the structure analysis by X-ray crystallography considerably (2). Since then, freeze-etching electron microscopy has been applied extensively to hydrated crystals of a^fao as well as 060 molecules. The surface topography was obtained by heavy-metal shadowing (Fig. 1). On shadowed crystal planes, the molecules appear spherical. The lattice constants obtained from the optical diffractograms agree well with X-ray crystallography.

;

^

v

v

,

- ^ v r . ..

vyr^sv^^

/

, - v TV y ^ r .• , 1 1 ' 1 ^ V » ) 1 1 V | M

11 IV.; a value too high for the

Q,

R-CH 2 -NH 2 ~ ^ R - C H r N H ;

r

^

R-CH-NH 2

'LJ

R-CH=NH 2 +

Scheme 2. Proposed mechanism for MAO-catalyzed amine oxidation (8).

covalent flavin coenzyme to be considered as a possible oxidizing agent. Model system SET amine oxidations show p values of ~-l (9). In the case of MAO B, the large kinetic isotope effects observed show that C-H bond cleavage is rate-limiting in enzyme reduction. Therefore, according to the second step of the mechanism depicted in Scheme 2, the H + abstraction step should exhibit a positive p value such as observed in the C-H cleavage step in the plasma amine oxidase mechanism (3). If the reverse commitment were larger than the forward committment in the mechanism shown in Scheme 2, it is conceivable that the cancelling effects of the negative p value (for the SET reaction) with the positive p for the H + abstraction step could lead to the observed small p value (Fig. 2B). The magnitude of the kinetic isotope effects even with substrate analogues that vary in pre-isotopic equilibria over 1-2 orders of magnitude (Fig.lA & B) demonstrates that the forward commitment for MAO B catalysis is much larger than the reverse

114 commitment. This conclusion is also supported by the H/T and D/T kinetic isotope effect data in an accompanying paper in this symposium (10). These considerations and the failure to observe any spectral evidence for the intermediacy of either neutral or anionic flavin radicals in stopped flow reductive half-reaction experiments with any of the substrate analogues tested further argues against the validity of the mechanism depicted in Scheme 2 for MAO B catalysis. As an alternative, we feel the available data support a H- abstraction as the most likely mechanism of C-H bond cleavage step in MAO B catalysis. Model studies of such reactions show little dependence of rate on 0 (11) as observed here with MAO B. Model H- abstraction reactions from the a-carbon of amines have demonstrated a pronounced stereoelectronic effect (12). These studies show the a-C-H bond is broken most readily when that bond is eclipsed with the lone pair orbital on the amino nitrogen. Thus, one explanation for the differential sensitivity of k3 values to meta and para substituents (Fig. 2A) is that meta substitution of the benzyl ring orients the substrate in the active site of MAO B to maximize eclipsing of the pro-R-a-C-H bond (13) with the amine lone pair orbital with the result that all analogues reduce the flavin coenzyme with similar rates. In the case of para-substituted analogues, the slower rates observed for flavin reduction also show a marked dependence on steric parameters which (Fig. 2B) could reflect altered geometries of the pro-R-a-C-H bond with the amino nitrogen orbital. The correlation of rate with E s is suggested to reflect the substituent effect on orientation of the pro-R-a-CH bond at angles differing from coplanarity from the amino nitrogen lone pair orbital which would result in a modulation of the rate of a C-H bond cleavage. It remains for future work to determine the identity of the H- abstracting species in MAO B catalysis and the events involved in electron transfer to the 8a-S-cysteinylFAD to form the flavin hydroquinone and the protonated imine product (14). The a—carbon-based substrate radical formed on H- abstraction would be expected to readily transfer an electron to the flavin coenzyme. Flavin radical intermediates have, to date, not been observed in anaerobic stopped flow experiments. This could be the result of very fast electron transfer processes subsequent to the C-H bond cleavage step.

115 Acknowledgements The authors wish to thank Dr. J. Klinman for valuable discussions. This work was supported by NIH Grant GM-2933 and by funds from Emory University.

References

1. Klinman, J.P. 1976. Biochemistry 15, 2018-2026 2. Miller, S.M. and Klinman, J.P. 1985. Biochemistry 24,2114-2127 3. Hartmann, C. and Klinman, J.P. 1991. Biochemistry 30, 4605-4611 4. Bondi, A. 1964. J. Phys. Chem. 68,441-451 5. Husain, M„ Edmondson, D.E., and Singer, T.P. 1982. Biochemistry 21, 595-600 6. Klinman, J.P. and Matthews, R.G. 1985. J. Am. Chem. Soc. 107, 1058-1060 7. Hansch, C. and Leo, A. 1979. Substituent Constants for Correlation Analysis in Chemistry and Biology, J. Wiley & Sons, New York 8. Silverman, R. 1991. Biochem. Soc. Trans. 19,201-206 9. Hull, L.A., Davis, G.T., Rosenblatt, D.H., and Mann, C.K. 1969. J. Phys. Chem.73, 2142-2146 10. Jonsson, T., Edmondson, D.E., and Klinman, J.P. These proceedings. 11. Pryor, W.A., Lin, T.H., Stanley, J.P., and Henderson, R.W. 1973. J. Am. Chem. Soc. 95, 6993-6998 12. Griller, D„ Howard, J.A., Marriott, P.R., and Scaiano, J.C. 1981. J. Am. Chem. Soc. 103, 619-623 13. Yu, P.H., Bailey, B.A., Durden, D.A. and Boulton, A.A. 1986. Biochem. Pharmac. 35, 1027-1036. 14. Edmondson, D.E., Bhattacharrya, A.K., and Walker, M.C. 1992. Biochemistry 32, 5196-5202.

Experimental Probes of Hydrogen Tunneling in Bovine Liver Monoamine Oxidase B

Thoriakur Jonsson and Judith P. Klinman University of California, Berkeley, Berkeley, CA 94720, U.S.A. Dale E. Edmondson Emory University School of Medicine, Atlanta, GA 30322, U.S.A.

Introduction Quantum mechanical tunneling has been well characterized for enzymatic reactions involving electron transfer (1,2). Whether the same phenomenon is important for hydrogen transfer reactions has until recently been largely unexplored. However, during the last several years clear evidence has emerged for the phenomenon of hydrogen tunneling in the reactions catalyzed by yeast alcohol dehydrogenase (3,4), horse liver alcohol dehydrogenase (5) and bovine serum amine oxidase (4,6). These reactions involve hydride transfer to a nicotinamide cofactor (alcohol dehydrogenases) and proton transfer to an active site base (bovine serum amine oxidase). This report extends previous studies to include bovine liver monoamine oxidase B, a flavin-dependent enzyme. The temperature dependence of competitive k^/kx and ko/kx isotope effects using the substrate p-MeObenzylamine reveals both deuterium and protium tunneling, as judged by values for isotopic Arrhenius pre-factors (AH/AT and AD/AT) which fall well below limits expected in the absence of hydrogen tunneling. As described earlier (4,6), an unambiguous demonstration of tunneling through temperature dependent isotope effects generally requires that the hydrogen transfer step be fully rate limiting across the experimental temperature range, in order to rule out a change in rate determining step arising from a temperature dependent commitment. In the case of the monoamine oxidase B reaction, we have not been able to eliminate the possibility of a small, temperature independent commitment; however, as discussed, the effect of such a commitment on the magnitude and interpretation of AH/AT and AD/AT is expected to be quite minor.

Results and discussion Table 1 summarizes isotope effects measured for p-MeO-benzylamine oxidation at pH 7.5. At 25°C, the primary kH/kT isotope effect is 21.4±0.8, which is comparable to a kn/kD isotope effect of 8.4 [from the Schwain-Schaad (7) relationship, (kn/kD)^- 4 ^ = kH/kT]This value is in good agreement with a previously determined value for kn/kD using stopped flow kinetics (8). Analogous to the original Schwain-Schaad expression, multiple isotope effects can be related by the expression (kD/kx)3-26 = kn/kx- As first pointed out by Saunders (9), the exponent relating kn/kx to ko/kx is expected to be especially sensitive to tunneling, with exponential values greater than 3.26 characteristic of this phenomenon.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

118 Subsequently, Cha et al. (3) demonstrated that the exponent will fall below 3.26 when the hydrogen abstraction step is only partially rate limiting. In the monoamine oxidase B reaction, exponents relating observed primary and secondary isotope effects at pH 7.5 and 25°C are somewhat reduced from 3.26 to 3.08 and 2.14, respectively (Table 1), indicating the possibility of a commitment. Table 1. Observed Isotope Effects and Exponential Relationships for p-MeO-Benzylamine Oxidation Catalyzed by Monoamine Oxidase B. a l°exp. b 2°exp.b Temp. l°kH/k T l°kD/k T 20kD/kT 2°kH/kT 2.0 13.03±0.51 2.70710.040 1.115±0.012 1,095±0.007 2.5710.05 1.2010.15 1.119±0.034 2.703±0.102 3.08±0.12 2.1410.36 25.0 21.43±0.81 1.272±0.030 43.2 16.87±0.35 1.078±0.015 2.464±0.069 1.233±0.010 3.13±0.10 2.7910.53 Experiments performed in 0.10M HEPES, pH 7.5,5mg/ml reduced Triton X-100. ''exponent = ln(kH/kT)/ln(kiykT) When the temperature was lowered to 2°C, observed exponents were lowered even further (to 2.57 and 1.20) and the observed kn/kx isotope effect decreased to a value below that at 25°C. Since isotope effects are expected to increase with decreasing temperature, this is a clear indication of an increase in kinetic complexity through a temperature dependent commitment. To search for conditions that might give observed isotope effects closer to intrinsic values, D kc a t and D (kc a t/KM) were measured at pH 6.0,7.5 and 9.0 (Table 2). Table 2. Steady-state Isotope Effects on p-MeO-Benzylamine Oxidation as a Function of pH. a pH 9.0 pH 6.0 pH 7.5 6.010.3 7.511.3 5.910.3 Dkcat D 8.911.6 4.811.0 11.113.9 (kc a t /K M ) Experiments performed in 0.10M phosphate (pH 6.0), 0.10M HEPES (pH 7.5) or 0.10M borate (pH 9.0), at 25°C. It can be seen that there is a general trend where observed isotope effects increase as the pH is lowered. This suggests that oberved isotope effects are closer to intrinsic values at pH 6.0 than 7.5. Additionally, k c a t/KM decreases approximately 20-fold as the pH is lowered from 7.5 to 6.0 (data not shown), conditions expected to reduce the contribution of an external commitment to the overall rate. Encouraged by these findings, we pursued the measurement of k f j / k j and k o / k j isotope effects at pH 6.1. Table 3 summarizes observed isotope effects and exponents in the 10-43°C range. Experiments were not pursued below 10°C, due to an untenably slow reaction with the deuterated substrate. At 25°C, both isotope effects and exponents are found to be very similar to those measured at pH 7.5, indicating that if commitments are present they are not altered by the large change in rate that accompanies the reduction in pH. Of particular significance is the observation that observed exponents do not change systematically in the 10-43 °C range. This shows that there are no temperature dependent changes in the level of expression of intrinsic isotope effects and hence, that a commitment, if present, would have to be independent of temperature as well as pH. The fact that the primary exponent is unchanged,

119 Table 3. Temperature Dependence of Observed Isotope Effects and Exponents in the 1043°C Range at pH 6.1. a Temp. l°kH/kT l°kn/kT 2°kH/kT 2°kn/kT l°exp. 2°exp. 10.0 28.89±1.10 2.896±0.132 1.388±0.030 1.149±0.019 3.16±0.14 2.32±0.31 15.0 27.59±0.70 2.892±0.054 1.132±0.018 3.12±0.05 2.47±0.31 1.359±0.016 25.0 21.98±1.16 1.131±0.012 3.20±0.12 2.690±0.086 1.339±0.037 2.37±0.30 30.0 21.49±0.95 2.631±0.044 1.127±0.010 3.17±0.07 1.341±0.011 2.45±0.19 35.0 18.52±0.99 2.569±0.070 1.129±0.032 3.09±0.11 2.41±0.57 1.340±0.013 40.0 17.42±0.40 2.509±0.067 1.109±0.042 3.11±0.09 2.43±0.91 1.286±0.028 43.0 17.02±0.52 2.507*0.133 1.263±0.029 1.118±0.015 3.08±0.18 2.09*0.33 a All experiments performed in 0.10M phosphate, 5mg/ml reduced Triton X-100. despite a combined 260-fold change in rate (20-fold due to a change in pH from 7.5 to 6.1 and appproximately 13-fold due to change in temperature when going from 10°C to 43°C), supports the view that the H-transfer rate is fully expressed at pH 6.1. In this context we were surprised to find that the exponents relating secondary isotope effects all fall well below 3.26. Analogous to earlier studies with bovine plasma amine oxidase (6), the small magnitude of exponents relating secondary isotope effects rules out significant coupling of motion between primary and secondary positions of substrate. Although we do not yet have a satisfactory explanation for exponents below 3.26, there are two possibilities under consideration. The first is that a small temperature independent commitment is present under all conditions of measurement. Since secondary isotope effects, by nature of their smaller size, are more sensitive to commitments than primary effects, this could explain the greater reduction in the exponent relating secondary isotope effects. As the commitment does not appear to change with pH at 25°C nor with temperature at pH 6.1, it would have to be an internal commitment related to the hydrogen transfer process. A second, more speculative explanation relates to the nature of the Htransfer step itself. As described by Truhlar and co-workers (12), tunneling leads to corner cutting across the potential energy surface, with H and D crossing the barrier at different positions. In the event of a large degree of tunneling (see below), the position of barrier crossing for H vs D will differ significandy. Since secondary isotope effects reflect changes in force constant between the ground state and point of barrier penetration, this could give rise to "different" secondary isotope effects for D transfer ( k o / k r ) than H transfer (kn/kx). Under these conditions, the exponential relationship between k o / k r and k n / k r may be difficult to predict. In Figure 1 we show the temperature dependence of observed isotope effects at pH 6.1. Extrapolating the linear least squares fit to infinite temperature gives primary AH/AT and AD/AT values of 0.13+0.03 and 0.52±0.05, respectively. These values are both well below the lower limits of 0.6 and 0.9 expected in the absence of tunneling contributions (10). In light of the possibility of a temperature independent commitment, the data were re-analyzed on the assumption of a small commitment which would restore the secondary exponent to 3.26. Using a Cf value of 0.5 changed the data in Figure 1 very little, yielding values for AH/AT and AD/AT of 0.18 and 0.53 respectively. It should be noted that the presence of this commitment has a far greater effect on the individual isotope effect values, elevating

120

Figure 1. Temperature dependence of observed isotope effects at pH 6.1, L=H or D. (A) Observed primary kn/kx (o) and k c / k j (A) isotope effects. (B) Observed secondary k n / k j (o) and k c / k r (A) isotope effects. intrinsic isotope effects above measured values in Table 3, e.g., to a primary kn/kx of 32.5 and a secondary k n / k r of 1.51 at 25°C. Both of these appear outside the semi-classical range. As first pointed out by Grant and KlinmaiA the temperature dependence of kp/kx can provide an excellent control in situations where small commitments may arise in measured kn/kx parameters. Since the commitment contributing to k o / k j is reduced by the size of the primary k n / k o isotope effect (approximately 10-fold with monoamine oxidase B), there are no ambiguities with respect to the AD/AT value. In the present study AQ/AT is well below the limit of 0.9 expected in the absence of tunneling. It can be seen that the data presented clearly demonstrate that both protium and deuterium tunneling contribute to the monoamine oxidase B reaction. The temperature dependence of observed secondary isotope effects (Figure IB) does not give AH/AT or AD/AT values significantly below 1, the values being 0.77±0.15 and 0.92±0.04, respectively. While the small deviations from unity could be due to coupled motion between the hydrogen being transferred and the secondary hydrogen, the magnitude of exponents argues against this being the case. The large secondary isotope effects observed (Table 3) are close to equilibrium values seen for sp3 to sp^ rehybridizations. Thus, it is possible that the oxidation of substrate occurs in two steps, with an equilibrium secondary isotope effect on the first step and a large primary isotope effect on the second step. However, there is as yet no evidence for chemical intermediates in the monoamine oxidase B reaction in the turnover of benzylamines (8). As a result of substituent effects on benzylamine oxidation, which indicate no electronic effects of para and mera-substituted benzylamine analogues on hydrogen transfer (8, 11), the most straightforward mechanism which can be written for this enzyme involves a simple hydrogen atom abstraction process accompanied by significant tunneling.

Acknowledgements We acknowledge the assistance of Laurene Kelly in the purification of monoamine oxidase B. This work was supported by grants from the National Science Foundation (DMB 8911632 to J. P. K.) and National Institute of Health (GM 29433 to D. E. E.).

121 References 1. Axup, A. W„ Albin, M„ Mayo, S. L„ Crutchley, R. J. & Gray, H. B. (1988) J. Am. Chem. Soc. 110, 435 and references therein. 2. Boxer, S. G. (1990) Annu. Rev. Biophys. Biophys. Chem. 19,100. 3. Cha, Y. Murray, C. J. & Klinman, J. P. (1989) Science 243,1325. 4. Rucker, J., Cha, Y„ Jonsson, T., Grant, K. L. & Klinman, J. P. (1992) Biochemistry 31, 11489. 5. Bahnson, B„ Park, D-H., Kim, K„ Plapp, B. V. & Klinman, J. P. (1993) Biochemistry 32,5503. 6. Grant, K. L. & Klinman, J. P. (1989) Biochemistry 28, 6597. 7. Swain, C. G., Stivers, E. C., Reuwer, J. F. & Schaad, L. J. (1958) J. Am Chem. Soc. 80, 5885. 8. Walker, M. C. (1987) Ph. D. thesis, Emory University. 9. Saunders, W. H. (1985) J. Am. Chem. Soc. 107,164. 10. Schneider, M. E. & Stern, H. J. (1972) J. Am Chem. Soc. 94, 164. 11. Walker, M. C. & Edmondson, D. E., in this volume. 12. Truhlar, D. G. & Gordon, M. S. (1990) Science 249,491.

Stereospecificity and Deuterium Isotope Effect in the Oxidative Deamination Catalyzed by Flavine and Non-flavine Amine Oxidases P. H. Yu and B.A. Davis Neuropsychiatrie Research Unit, University of Saskatchewan, Saskatchewan, Canada, S7N 0W0

Saskatoon,

Introduction Two types of amine oxidases, namely, monoamine oxidase (MAO) and semicarbazide-sensitive amine oxidase (SSAO) are well known to be involved in the deamination of monoamines. MAO is a flavine-containing enzyme located in the outer membrane of mitochondria (13), while SSAO is a copper enzyme probably possessing 6-hydroxydopa as cofactor (7). During enzymatic oxidative deamination of primary amines, one hydrogen atom at the a-carbon position adjacent to the amino group is abstracted. Studies on the deamination of stereospecific deuterium substituted amines not only provide information about the participation of a carbon-hydrogen bond breaking step, but also reveal the stereospecificity during such oxidative reactions. In the present study R(a- 2 Hi)and S(a- 2 Hi) enantiomers of dopamine and benzylamine were synthesized by enzymatic procedures, and the stereospecificity and the deuterium isotope effect with respect to different amine oxidases were analyzed. Experimentals Mono-deuterated S(a- 2 Hi)-dopamine was synthesized by decarboxylation of L-(a- 2 Hi)-3,4-dihydroxy-phenylalanine (L-DOPA) in water and R(a- 2 Hi)dopamine by decarboxylation of unsubstituted L-DOPA in 2 H 2 0 (15). Stereospecific (a- 2 Hi) benzylamine enantiomers are synthesized chemically by means of alcohol dehydrogenase (16). MAO-A and MAO-B were prepared from mitochondria of different liver tissues. SSAO was partially purified from human umbilical artery and rat aorta. Diamine oxidase was prepared from hog kidney and pea seedling. MAO activity towards dopamine was measured by a HPLCelectrochemical detection procedure and benzylamine by a fluorometric method by measuring the production of hydrogen peroxide (13).

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

124 Results and Discussion When stereospecifically deuterated amine enantiomers R - ( a - 2 H i ) - and S(a- 2 Hi)-benzylamine and dopamine are incubated with different amine oxidases, the

deuterium

atom

may

be

either

retained

or

eliminated

to

form

monodeuterated or non-deuterated benzaldehyde products respectively.

These

two benzaldehydes are readily separated from one another and identifiable by HPLC. They have also been also identified by mass-spectrometry (15). As can be seen in Table 1, three types of stereospecific abstraction of a proton from the a - c a r b o n of benzylamine during deamination observed.

have

been

[1] Pro-R proton was removed from the a-carbon of benzylamine and

dopamine, for instance, when the dopamine was incubated with MAO-A or Table 1. Stereospecific abstraction of hydrogen from a-carbon of monoamines during oxidative deamination catalyzed by different amines oxidases. Enzyme

Dopamine

Benzylamine

Reference

Type I Rat liver MAO-A

R

(15)

Human placenta MAO-A

R

(15)

Rat liver MAO-B

R

R

(15,16)

Human platelet MAO-B

R

R

(15,16)

Bovine liver MAO-B

R

R

(10,16)

Hog kidney MAO-B

R

R

(10,16)

Rat aorta SSAO

S

S

(14,16)

Hog kidney diamine oxidase

S

S

(14,16)

Pea seedling diamine oxidase

S

S

(2, 3,14,16)

Type II

Type III Bovine plasma amine oxidase Human umbilical artery SSAO

S+R

S S+R

R: abstraction of pro-R hydrogen; S: abstraction of pro-S hydrogen. *P.H. Yu, D.M. Zuo and B.A. Davis (unpublished)

(4,10,11,14)

•

125 MAO-B, or benzylamine with MAO-B from various sources. [2] Pro-S proton was removed from the a-carbon in reactions catabolized by rat aorta SSAO or diamine oxidase from hog kidney and pea seedling. [3] No stereospecificity was involved in benzylamine deamination catalyzed by bovine serum amine oxidase (BSAO) and SSAO from human umbilical arteries or serum. It is interesting that both MAO-A and MAO-B are bound to the mitochondrial outer membrane (6), and both types of enzyme possess a flavine prosthetic functional group (9).

Although MAO-A and MAO-B are different

isozymes with different primary structures (1), the primary structures adjacent to the flavine binding sites of both MAO-A and MAO-B have been found to be identical

(12).

It is reasonable

to assume

that they

have

a

similar

stereoconfiguration in binding the substrate, and so only the pro-R proton is removed during oxidation. Different diamine oxidases and SSAOs are known to be soluble or plasma membrane bound enzymes and require C u + + as cofactor, but the prosthetic group is less understood.

The tertiary structure of these enzymes

at the active center is obviously different from that of MAO, which facilitates the elimination of the pro-R proton. The prosthetic group for bovine serum amine oxidase

(BSAO) has now identified as 6-hydroxydopa

(7) rather as a

pyridoxal(PLP)- (5) or pyrroloquinoline quinone (PQQ ) (8) group. It is not yet known whether or not other diamine oxidases or SSAOs possess the same cofactor as BSAO. Enzymes in the type III oxidize (Table 1) either R-(a- 2 Hi)- or S( a - 2H 1 )-benzylamine, and break either the C-H or C- 2 H bond with similar preference. It is unclear why the BSAO and human umbilical artery SSAO do not exhibit stereospecificity in their deamination of dopamine, although the enzymes may

possess

the same prosthetic group as other SSAOs.

The lack of

stereospecificity with respect to BSAO has been previously observed (10). The kinetic deuterium isotope effect in the above reactions varied greatly. Both VH/VD and (V/K)H/(V/K)D values were compared. The deuterium isotope effect with respect to benzylamine, as is the case for reactions catabolized by MAOB

and

rat

aorta

SSAO

((V/K)

H

/(V/K)

d

values are 4 . 0 2 ± 0 . 1

and

2.51±0.05

respectively), indicates that the proton abstraction is probably the rate-limiting step in the deamination. The low isotope effect in the systems in which dopamine is catabolized by diamine oxidases from hog kidney and pea seedling ( ( V / K ) H / ( V / K ) d values are 1.55±0.06 and 1.68±0.06) suggests that cleavage of the C-H bond at the a-carbon position is faster than subsequent steps of the reaction and thus it is unlikely that this bond break is involved in the rate-limiting step.

126 There was no correlation with the three stereospecific types of amines oxidases with respect to deuterium isotpe effects.

The

mechanisms

of

oxidative

deamination catalyzed by the different enzymes appear to be f u n d a m e n t l y different. SSAOs from both human artery tissue and serum do not exhibit a significant amout of deuterium isotope effect or stereospecificity.

They are

distinctly different from BSAO. This observation also suggests that the cleavage of the hydrogen atom on the a-carbon is not the rate-limiting step in the deamination of benzylamine catalyzed by human SSAO (Yu et al, unpublished data). Acknowledgements This research was supported by Saskatchewan Health, the Medical Research Council, Deprenyl Research and the Parkinson Foundation of Canada. References (1)

Bach, A.W.J., Lan, N.C., Johnson, D.L., Abell, C. W., Bembenek, M. E., Kwan, S. W., Seeberg, P.H., Shih, J. C. 1988. Proc. Natl. Acad. Sci. USA. 85:4934. (2) Battersby, A.R., Staunton, J., Summer, M.C. 1976. J. Chem. Soc. Perkin 1:1052. (3) Battersby, A.R., Staunton, J., Summer, M.C., Southgate, R. 1979a. J. Chem. Soc. Perkin /:45. (4) Battersby, A.R., Buckley, D.G., Staunton, J., William, P.J. 1979b. J. Chem. Soc. Perkin 1:2250. (5) Buffoni, F. 1966. Pharmacol. Rev. 25:1163. (6) Greenawalt, J.W. 1972. Biochem. Pharmacol. 5:207. (7) Janes, S.M., Mu, D., Weimer D, Smith, J.A., Kaur, S., Maltby, D., Burlingame, A.L., Klinman, J.P. 1990. Science 248: 981. (8) Lobenstein-Verbeek, C.L., Jongejan, J.A., Frank, J., Duine, J.A. 1984. FEBS Lett. 270:305. (9) Nara, S., Igave, I., Gomes, B., Yasonobu, K. 1966. Biochem. Biophys. Res. Comm. 23:324. (10) Summers, M.C. Markovic, R., Klinman, J.P. 1979. Biochem. 28:1969. (11) Suva, R.H., Abeles, R.H. 1978. Biochemistry 27:3538. (12) Yu, P.H. 1981. Can. J. Biochem. 59:30. (13) Yu, P.H. 1986. In, Neuromethods V: Neurotransmitter Enzymes (eds. A.A. Boulton, G.B. Baker and P.H. Yu) Humana Press, Clifton, New Jersey, p. 235. (14) Yu, P.H. 1988. Biochem. Cell Biol. 66: 853. (15) Yu, P.H., Bailey, B.A., Durden, D.A. and Boulton, A.A. 1986. Biochem. Pharmacol. 35:1027. (16) Yu, P.H., Davis, B.A. 1988. Int. J. Biochem. 20:1197.

CHARACTERIZATION OF L-ASPARTATE OXIDASE OVEREXPRESSED COLI

IN E.

A. Negri, M. Mortarino, T. Simonie, G. Tedeschi, S. Ronchi Istituto di Fisiologia Veterinaria e Biochimica and C.I.S.M.I., Università di Milano, Milano, Italy H.G. Gassen Institut fur Biochemie, Technische Hoschule, Darmstadt, Germany

Introduction L-aspartate

oxidase

(LASPO)

is

involved

biosynthesis of pyridine nucleotide organisms

in

the

coenzymes

in

de

novo

several

(l) . This flavoprotein catalyzes the oxidation

L-aspartate

to

iminoaspartate

dihydroxyacetone

phosphate

intermediate

this

condensation

in

is carried

which to

is then

give

biosynthetic

condensed

quinolinate, pathway.

out by a second

a

The

of

with key

latter

enzyme, protein

A.

LASPO and protein A are sometimes referred as subunits of the quinolinate

synthetase

complex,

although

at the moment

the

existence of such complex has not been experimentally proved. Gene complementation studies suggest that LASPO

is

involved

in quinolinate

and

aerobic

organisms

bioyinthesis

both

in

anaerobic

(1) . This implies that the enzyme must be capable

of using electron acceptors different from 0 2 . This, in turn, poses the problem whether 0 2 itself is the real oxidant also in aerobic conditions in vivo. The genes

for

cloned from acids

(M r

LASPO

E.coli

(nadB) and protein A

(nadA) have

been

(2) . LASPO is a polypetide of 540 amino

60306) containing

1 mol of non-covalently

bound

FAD per mol protein. Protein A is a polypeptide of 280 amino

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

128

acids (Mr 31555) . Protein A has also been cloned from S. typhimurium, together with a gene (nadR) involved in the regulation of the expression of nadA and nadB genes in this organism (3). In contrast with the detailed knowledge of the genetics of NAD biosynthesis, less informations are available about the proteins involved in such pathways. In particular, the minute quantity of enzyme available from the natural source allowed only a preliminary characterization of E. coli LASPO, conducted using an enzyme preparation at an early stage of purification (4) . Recently, however, the nadB gene product (LASPO) has been overexpressed in E. coli under the control of the heat inducible Apl promoter (5) , supplying a large amount of easily purifiable enzyme. A preliminary report on the properties of the overexpressed enzyme is presented in this paper.

Results Isolation of Intact L-Aspartate Oxidase Overexpressed in E. coli BL 21 LASPO, isolated from NK6042S, the bacterial strain-originally used in Ref. 5, risulted partially proteolyzed at Argl27. The resulting fragments, which were resistant to further proteolysis, could not be separated from the intact enzyme. In order to overcome this problem we identified the protease responsible for the cleavage (ompT) and used as the overexpression host an E. coli strain lacking this protease (BL21) transformed with plasmids pQAg213 and pRK248, carrying the nadB gene and the the thermosensitive repressor cl 857, respectively (5). The yield of enzyme purified using BL21 was comparable with the one previously reported (5) but the specific activity was about doubled.

129 FAD Binding to Apo-L-Aspartate Oxidase Since

FAD

loss

was

observed

during

the

last

purification

step, it w a s necessary to determine the strength of binding to the enzyme prior to any further study.

The

coenzyme

K d could

be measured

coenzyme

spectrophotometric by

following

the

quencing of the protein fluorescence upon binding of FAD. Fig. 1

shows

a

kinetic

analysis

of

the

process.

The

following

values were obtained from a second order analysis of the data of Fig. min-1

1: k o n =

1.53 x 10 4 M _ 1

and K d of 6.6 x 10~ 7 M

min-1,

kQff

=

(in 50 mM Hepes,

x

10~ 2

pH 8.0,

20 %

1.01

glycerol, 10 °C). The K d value was in good agreement with the K^j of 7.3 x 1 0 - 7 M determined at 25 °C and the value of 5.6 x 10~7

M

obtained by titration

amounts observed similar

of

of

apoprotein

chromatographically

for

the

dissociation

to the one observed

pure of

FAD

(10

from

for other amino

such as pig kidney D-amino acid oxidase of FAD to apo-LASPO

FAD

with

increasing

°C) . T h e

LASPO acid

is

Kd

thus

oxidases,

(6). However, binding

is a simple second order process and

is

not followed by the slow conformational change of the protein observed in the case of other flavoenzymes

Fig.

1.

Time

dependence

of

the

fluorescence upon binding to ( • ) 5, ( • )

10, and (A.) 20 jUM FAD.

(6).

decrease 0.5, ( • )

in

apo-protein

1, ( A ) 2.5,

(•)

130 Primary Structure Similarity between L-Aspartate Oxidase and Other Flavoproteins The amino acid sequence of E. coli LASPO, deduced from the gene

(5),

shows

no

extended

similarity

flavooxidases, except for those regions (1-37

with

other

and 365-375 of

LASPO) which are known to be involved in FAD binding and are also present in D-amino acid oxidase and D-aspartate oxidase (see

Faotto,

L.

et

al. ,

this

Volume

and

Ref.

7).

Surprisingly, Fig 2 shows that LASPO presents a high degree of similarity

with the

flavoprotein

subunits

of

succinate

dehydrogenase (SDH) (8) and fumarate reductase (FRD) (9). The latter

are

multi-subunit

membrane-bound

proteins

(10)

containing one mol of covalently bound FAD (at position His45 in SDH and His44 in FRD). The similarity extends to the whole molecule except for the C-terminal portion which,

however,

might be involved in subunit-subunit interactions in SDH and FRD. A high degree of identity is also present at the level of

region

227-253

of

FRD which

has

been

shown

by

site-

directed mutagenesis to comprise two residues, namely His232 and Arg248, directly involved in catalysis

(11). In view of

these similarities the possibility of association of LASPO with some membrane element cannot be ruled out, despite the fact

that

catalytic

the

enzyme

activity

appears

is not

to

be

increased

cytosolic

and

by the presence

its of

detergents as observed, for example, in the case of pyruvate oxidase and malate oxidase (12, 13). Site Directed Mutagenesis of L-Aspartate Oxidase Site directed mutagenesis Ser45 with Arg and

of LASPO at positions Glu43 and

His, respectively,

was

carried

out

in

order to verify the effect of the presence of amino acids identical to those known to be involved in coenzyme binding in FRD and SDH. A 1.2 kb EcoRI fragment of E.coli nadB gene

131 LASPO FRDa SDHa

MNTLPEHSCDVLIXGSGAAGLSLALRLADQHQ VXVLSKGPVTEGSTF Q-TFQA DLAIVGAGGAGLRAAIAAAQANPNAKIALISKVYPKRSHTV M-KLPVREFDAWIGAGGAGIARALQISQSGQTC--ALLSKVFPTRSHTV : ::: *..::*.*.**:. *: .:. : . .:** : .*

47 46 47

LASPO FRDa SDHa

YAQGGIAAVFDET---DSIDSHVEDTLIAGAGICDRHAVEFVASNARSCVQ AAEGG-SAAVAQDH-DSFEYHFHDTVAGGDWLCEQDWDYFVHHCPTEMT SAQGGITVALGNTHEDNWEWHMYDTVKGSDYIGDQDAIEYMCKTGPEAIL *:**:.: .: *: . *. **. .: ::: :::.. ..

95 94 97

LASPO FRDa SDHa

WLIDQGVLFD THIQPNGEESYHLTREGGHSHRRILHAADATGRE QLELWGCPWSRRPDGSVNVRRFGGM KXERTWFAADKTGFH ELEHMGLPFSRLDDGRIYQRPFGGQSKNF GGEQAARTAAAADRTGHA

139 134 14 4

LASPO FRDa SDAa

VETTLVSKALNHPNIRVLERTNAVDLIVSDKIGLPGTRRVVGAWVWNRNK MLHTLFQTSLQFPQIQRFDEHFVLDILVD-DGHVRGLVAM NMME LLHTLYQQNLK-HHTTIFSEWYALDLVKNQDGAWGCTAL CIET

189 177 187

LASPO FRDa SDHa

ETVETCHAKAWLATGGASKVYQYTTNPDISSGDGIAMAWRAGCRVANLE GTLVQIRANAWMATGGAGRVYRYNTNGGIVTGDGMGMALSHGVPLRDME GEWYFKARATVLATGGAGRIYQSTTNAHINTGDGVGMAIRAGVPVQDME

239 227 237

LASPO FRDa SDHa

FNQFHPTALYHPQARNFLLTEALRGEGAYLKRPDGTRFMPDFDERGE FVQYHPTGL PGSGILMTEGCRGEGGILVNKNGYRYLQDYGMGPETPL MWQFHPTGI AGAGVLVTEGCRGEGGYLLNKHGERFMERYA

286 274 277

LASPO FRDa SDHa

LAPRDIVARAIDHEMKRLGADC MFLDISHKPADF GEPKNKYMELGPRDKVSQAFWHEWRK-GNTISTPRGDWYLDLRHLGEKK —PNAK—DLAGRDWARSIMIEIRE-GRGCDGPWGPHAKLKLDHLGKEV

320 323 322

LASPO FRDa SDHa

IRQHFPMIYEKLLGL-GIDLTQEPVPIVPAAHYTCGGV MVD LHERLPFICELAKAYVGVDPVKEPIPVRPTAHYTMGGIET LESRLPGILELSRTFAHVDPVKEPIPVIPTCHYMMGGIPTKVTGQALTVN

360 363 372

LASPO FRDa SDHa

DHG-RTDVEGLYAIGEVSYTGLHGANRMASNSLLECLVYGWSAAEDITRR DQNCETRIKGLFAVGECSSVGLHGANRLGSNSLAELWFGRLAGEQATER EKGEDVWPGLFAVGEIACVSVHGANRLGGNSLLDLWFGRAAGLHLQES

409 413 422

LASPO FRDa SDHa

MPYAHDISTLPPWDESRVENPDERWIQHN WHELRLFMWD AATAGNGNEAAI—EAQAAGVEQRLKDLVNQDGGENWAKIRDEMGLAMEE IAEQGALRDAS ESDVEASLDRLNRWNNNRNGEDPVAIRKALQECMQH *:::...*. * ::

449 461 469

LASPO FRDa SDHa

YVGIVRTTKRLERALRRITMLQQE IDEYYAHFRVSDNL—LELRN GCGIYRTPELMQKTIDKLAELQERFKRVRITDTSSVFN-TDLLYTIELGH NFSVFREGDAMAKGLEQLKVIRERLKNARLDDTSSEFN-TQRVECLELDN ::*: :. ::. . * .:: :**:

492 510 518

LASPO FRDa SDHa

LVQVAELIVRCAMMRKESRGLH— FTLDYPE LLTHS GLNVAECMAHSAMARKESRGAHQRLDEGCTERDDVNFLKHTLAFRDADGT LMETAYATAVSANFRTESRGAHSRFD—FPDRDDENWLCHSLYLPESESM

526 560 566

LASPO FRDa SDHa

GPSI LSPGNH-Y INR TRLEYS-QVKI-TTLPPAKRVYGGEADAADKAEAANKKEKANG TRRSVNMEPKLRPAFPPKIRTY

*

:

.

:

*

.:

***

**

540 601 588

Fig. 2. A l i g n m e n t of the amino acid sequence of LASPO with t h o s e of the flavoprotein suburiit of E. coli FRD and SDH.

132

was used as DNA template for the in vitro synthesis of the mutated strands primed by synthetic oligonucleotides. The resulting genes, reconstituted in the expression system, produced two single mutated proteins (E43R and S45H) with an yield similar to that obtained for the wild type enzyme. Preliminary results show that both mutated enzymes are expressed as holoproteins but bind FAD less tightly than the wild type, the coenzyme being completely lost during the DEAE purification step (while almost no loss of FAD for the wild type enzyme is observed at this stage of purification) . In addition, both mutated enzymes appear to be catalitically inactive also in the presence of added FAD. These results indicate that both mutations introduced a very pronounced effect on the coenzyme properties, confirming that this region is indeed involved in FAD binding also in LASPO. As expected, the introduction of the single mutation S45H was in itself not sufficient to give covalent incorporation of FAD in LASPO. Further studies, and the obtainement of the double mutant E43R, S45H are needed to clarify the effects exerted by mutations on the interactions between FAD and apo-LASPO. Binding of Ligands to L-Aspartate Oxidase LASPO is extremely specific as concerns substrate specificity, being active only towards L-aspartate. On the contrary, many dicarboxylic compounds can act as inhibitors. Among them, it was particularly interesting to study the interaction of LASPO with succinate, fumarate and oxaloacetate. Fig. 3 shows the absorption spectrum of the LASPO-succinate complex (Kd = 2.4 x 10 -4 M) . The spectral perturbation induced by the ligand was very small, with no indication of the resolution of the 450 nm band often observed in the case of other flavoenzymes upon binding of ligands to the active site. Qualitatively similar results were obtained with fumarate (Kd = 1.9 x 10~ 4 M) and

133

Fig. 3. Visible absorption spectrum of LASPO before ( ) and after ( ) addition of 1 m M succinate in 50 m M Hepes, 20 % glycerol, p H 8.0, 25 °C. Inset: differential absorption spectrum observed during the titration of LASPO w i t h different amounts of succinate.

F i g . 4. V i s i b l e a b s o r p t i o n s p e c t r a o b s e r v e d a t t h e t i m e indicated during reduction of LASPO in the presence of 300 ,MM x a n t h i n e , 3 nM x a n t h i n e oxidase and 2.5 f*. M benzylviologen under anaerobiosis. Conditions as in Fig. 3.

134

oxaloacetate (Kd = 6.0 x 10 -5 M). Thus, in agreement with the primary structure similarity outlined above, LASPO binds succinate and fumarate with affinity similar to the one shown by FRD and SDH (10). Redox Forms of L-Aspartate Oxidase A feature common to flavooxidases is the stabilization of the anion semiquinone during anaerobic reduction experiments in the presence of EDTA and flavins or xanthine/xanthine oxidase systems as source of reduction equivalents (14, 15). No semiquinone formation was observed upon reduction of LASPO in the absence of ligands (data not shown). On the contrary, the anion semiquinone is observed when succinate is present (Fig. 4) •

Reaction of L-Aspartate Oxidase with Sulfite Another distinctive property of flavooxidases is the stabilization of an adduct between the N(5) position of the coenzyme and sulfite (16). LASPO forms a sulfite adduct with a low Kd (9.7 x 10 -6 M) , indicating a normal "oxidase-type" reactivity (not shown). However, while attainement of equilibrium after each addition of sulfite during titration experiments in the case of other oxidases is a fast process, in the case of LASPO it is a multi-phasic process which is complete in about 3 0 min at 25 °C.

Discussion The properties of E. coli LASPO reported in this paper differ from those usually observed for other amino acid oxidases as concerns the primary structure, the spectral perturbation upon binding of ligands, the semiquinone stabilization by

135 free holoenzyme and the kinetics of sulfite adduct formation. These

results

are

in

agreement

with

the

very

different

biological role of LASPO, an enzyme involved in an anabolic pathway, as compared to the other amino acid oxidases, which are involved properties

in catabolic pathways. of

LASPO,

protein A, will primary

also

allow to address the

structure

functional

conducted

Further

homology

similarities

with

beyond

in

studies

the

question

SDH

the

and

simple

on

the

presence

of

whether

the

FRD need

implies for

a

structure designed to bind dicarboxylic molecules.

Acknowledgments This work was from

the

supported

C.N.R.

by grants from

Target

Project

the M.U.R.S.T. "Biotechnology

Bioinstrumentation"

References 1. Foster, J. W., A. G. Moat. 1980. Microbiol. Rev. 44, 83-105. 2. Flachmann, R., N. Kunz, J. Seifert, M. Gutlich, F.-J. Wientjes, A. Laufer, and H. G. Gassen. 1988. Eur. J. Biochem. 175, 221-228. 3. Foster, J. W. , Y. K. Park, T. Penfound, T. Fenger, M. Spector. 1990. J. Bacteriol. 172, 4187-4196. 4. Nasu, S., F. D. Wicks, R. H. Gholson. 1982. J. Biol. Chem. 257, 626-632. 5. Seifert, J., N. Kunz, R. Flachmann, A. Laufer, K.-D. Jany, H. G. Gassen. 1990. Biol. Chem. Hoppe-Seyler 371, 239-248. 6. Massey, V. , B. Curti. 1966. J. Biol. Chem. 241, 34173423 . 7. Negri, A., F. Ceciliani, G. Tedeschi, T. Simonie, S.

and and

136 R o n c h i . 1992. J. B i o l . C h e m . 267,

11865-11871.

8. W o o d , D . , M . G. D a r l i s o n , R. J. W i l d e , J. R. G u e s t . B i o c h e m . J. 222, 519-534. 9. C o l e , S. T. 1982. Eur. J. B i o c h e m . 122,

1984.

479-484.

10. A c k r e l l , B . A . C . , M. K. J o h n s o n , R. P. G u n s a l u s , G. C e c c h i n i . 1992. 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 F l a v o e n z y m e s , V o l . 3 (F. M u l l e r e d . ) , C R C P r e s s , B o c a R a t o n , FL, pp, 2 2 9 - 2 9 7 . 11. S c h r o d e r , I., R. P. G u n s a l u s , B. A . C. A c k r e l l , C o c h r a n , G. C e c c h i n i . 1991. J. B i o l . C h e m . 266, 13579. 12. R u s s e l , P., H. L. S c h r ö c k , R. B. G e n n i s . C h e m . 252, 7883-7887. 13. N a r i n d r a s o r a s a k , B i o l . C h e m . 254,

1977. J.

S., A. H. G o l d i e , B. D. S a n w a l . 1540-1545.

14. M a s s e y , V. , P. H e m m e r i c h . 246-257.

B. 13572Biol. 1979. J.

1980. B i o c h e m . Soc. T r a n s .

15. M a s s e y , V . 1991. In: F l a v i n s a n d F l a v o p r o t e i n s 1990 C u r t i , S. R o n c h i , a n d G. Z a n e t t i e d s . ) . W a l t e r d e G r u y t e r B e r l i n , N e w Y o r k . pp. 5 9 - 6 6 .

8, (B.

16. M a s s e y , V. , F. M u l l e r , R. F e l d b e r g , M. S c h u m a n , P.A. S u l l i v a n , L. G. H o w e l l , S. G. M a y h e w , R. G. M a t t h e w s , G. P. F o u s t . 1969. J. B i o l . C h e m . 244, 3999-4006.

IDENTIFICATION OF ANOTHER SUBUNIT IN L-PHENYLALANINE OXIDASE FROM PSEUDOMONAS SP. P-501 AND THE PRIMARY STRUCTURE OF THE SUBUNIT

Etsuko B. Mukouyama, Haruo Suzuki, Toshihide Sasaki Department

of

Biophysical

Chemistry, Kitasato

University

School

of

Medicine, Sagamihara, Kanagawa 228 Hirokazu Koyama R & D

Division, Kikkoman Corporation, 339 Nöda, Chiba 278, JAPAN

Introduction L-Phenylalanine oxidative

oxidase from Pseudomonas sp. P-501 catalyzes

deamination

alanine(l-3).

The

and

enzyme

oxygenative

decarboxylation

has been known to be a

dimer

both

of

L-phenyl-

of

identical

subunit(Mr 68,000) and contains 2 mol of FAD per mol of enzyme(2). During the structural studies on the enzyme, a possible presence of the another subunit was indicated. of

Here we describe the

identification

the another subunit, the subunit composition of the enzyme, and

its

primary structure.

Results and Discussion When

the highly purified enzyme was analyzed by

HPLC,

two peaks were observed(Fig. 1).

areas

was

materials lane

estimated to be 1:6.4. were

reversed-phase

The ratio of peak 1 to

The molecular masses of

estimated to be 8.2kDa and

4 and 5) by SDS/PAGE(4).

a

68kDa,

peak

these

2

peak

respectively(Fig.2,

From the densitometric analysis

of

the

staining of SDS/PAGE, the ratio of 8.2kDa and 68kDa bands was calculated to

be 1:6.5.

This agreed with the value obtained by the HPLC

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

analysis

138 of the enzyme.

From the ratio and the molecular masses of two

species,

the ratio of small to large molecular species was estimated to be Considering the method of Mr determination, the small(a ) and

1.28.

large(/3 )

molecular species must be present 1:1 in the enzyme.

Fig. l(left)

Reversed-phase HPLC profile of L-phenylalanine oxidase.

The absorbance was monitored at 210nm and acetonitrile(AcCN) gradient is indicated(

).

Fig. 2(right)

SDS/PAGE analysis of L-phenylalanine oxidase and the peak

materials

from

the

Coomassie

brilliant blue.

peptide standard.

reversed-phase HPLC.

The

gel

was

stained

Lane 1, the protein standard.

Lane

2,

with the

Lane 3, L-phenylalanine oxidase, Lane 4 and 5, peak 1

and peak 2 materials in Fig.l, respectively. Koyama reported that the enzyme was dissociated into monomer in the presence

of 0.2M Na2S0 4 , so we tested if a also dissociates

enzyme.

In the absence of 0.2M ^ 2 8 0 4 , the enzyme was

than the

eluted

aldolase(Mr 158,000) and no other peak was detected. presence

detectable concentrated SDS/PAGE.

with

about

Mr 65,000.

The eluates around

Centricon-30(Amicon

Co.,

Ltd.),

The data showed the presence of a and 0

and

the

earlier

However,

of 0.2M Na 2 S0 4 and 0.1% SDS, the enzyme showed

peak

from

in

only

one

Mr 65,000

was

analyzed

molecular

by

species

139 in the eluates.

This suggests the strong interaction between these

two

molecular species. To confirm this, we performed isoelectric focusing of the enzyme in the presence of 8M urea.

Only one band with

was

The fluorescence

observed

presence treatment was

at the pi value of 4.9.

of

FAD.

species

protein in the

with the microwave irradiation, then the SDS/PAGE of the

gel

inactivated

Both the a and

were observed, so even in the 8M urea, the a

dissociated from the enzyme protein. as

was

the the

as the second dimension.

gel

indicated by

performed

The

fluorescence

0

species was

not

From these results, we conclude a

the enzyme subunit, and the enzyme was suggested to be

two pairs of an a 0

molecular

present

as

dimer.

As the start to know the relationship between the structure and the function both the

of enzyme, we analyzed the N-terminal amino acid sequences

subunits. 58th

amino

determined amino

The sequence of the a subunit has been acid residues, whereas the fi subunit

presumably

acid.

because of the heterogeneity of

after

to

could

be

the

and

the

resulting

peptide

N-terminal

fragments

These results were summarized in Fig.3.

consists of 92 amino acid residues, and contains the

the

not

or were

The C-terminal Gly was determined by the amino acid analysis

hydrazinolysis.

subunit for

determined

The a subunit was digested with chymotrypsin, trypsin

asparaginylendopeptidase, sequenced.

of

The

a

candidate

common sequence characteristic of the AMP binding in

the

FAD

binding domain(5), Gly-Gly-Gly-Ala-Gly-Gly.

1 10 20 30 40 50 60 KKIATTVGEARLSGINYRHPOSALVSYPVAAAAP LGRL PA6NYRI A IVGGGAGGIAALYE 61 70 80 90 92 LGRLAATLPAGSGIDVQIYEADPDSFLHDRPG Fig. 3 for

The primary structure of the a

amino acid

is used.

subunit.

One letter code

The sequences were determined from

the

proteolytic peptide fragments. Using the isolated a subunit, we examined whether the a

interacts

with FAD by measuring the visible absorption spectrum in 50 mM potassium phosphate buffer at pH 7.0 and at room temperature. a

The addition of the

at the molar ratio of 1:1 did not change the spectrum of FAD.

140 We also tested if the antibody against enzyme

activity.

anti-a

antisera were obtained.

both the isolated incubated by

a

has any effect on the the

BALB/C mice were immunized with the

a

a

subunit and the enzyme by ELISA.

measuring

the

oxygen uptake using Phe or Met

indicate that the though

the is

a

as

to

assayed

substrate. These

The

results

subunit is not bound to FAD in the enzyme molecule,

subunit contains the sequence of the AMP another

and

The antisera were

with the enzyme, then the activity of the enzyme was

antisera did not have any effect on the enzyme activity.

There

subunit

The antisera were shown to be bound

possibility

that the

isolated

conformation different from the native one, thus the

binding a a

domains.

subunit

has

subunit did

a not

interact with FAD and the antisera had no effect on the enzyme activity. The possibility can be clarified by preparing the intact subunit.

Acknowledgments

We are indebted to Professor Emeritus S. Horie and Dr. Y. KawamuraKonishi for their valuable discussion.

References

1.

Koyama.H. 1982. J.Biochem. 92, 1235-1240

2.

Koyama.H. 1983. J.Biochem. 93, 1313-1319

3.

Koyama.H.

4.

Hashimoto,F., Horigome,T., Kanbayashi,M., Yoshida.K.,

1984. J.Biochem. 96, 421-427

&Sugano,H.

1983. Anal.Biochem. 129, 192-199

5.

Hanukoglu.I. & Gutfinger.T. 1989. Eur.J.Biochem. 1J0, 479-484

Kinetic Isotope Effects on Reductive Half-Reactions of Flavoprotein Oxidases P. F. Fitzpatrick1-2, J. M. Denu1, J. J. Emanuele1, and V. Menon2 Departments of 1 Biochemistry and Biophysics and 2Chemistry, Texas A&M University, College Station, TX 77843-2128 USA

Introduction Kinetic isotope effects provide an extremely powerful probe of enzyme mechanisms. Detailed examination of the quantitative effects of replacement of multiple atoms in a substrate with heavier isotopes can provide information about structures of intermediates and transition states which are not available by any other approach. In addition, the structural change in the substrate upon isotopic substitution is so minor that it is commonly assumed that no mechanistic changes occur. This need not be the case with approaches involving substrate analogs which may utilize alternate reaction pathways. However, kinetic isotope effects measured on kinetic parameters seldom reflect direcdy the effect on a single chemical step. Rather, the intrinsic effect is masked by slower nonchemical steps. Thus, to determine the effects of isotopic substitution upon individual chemical steps, it is helpful to find conditions where the chemical step of interest is rate-limiting. We have been utilizing a variety of deuterium kinetic isotope effects to examine the chemical mechanisms of several FAD-containing oxidases. These are D-amino acid oxidase (DAAO), possibly the most studied of this group of enzymes, tryptophan monooxygenase (TMO), an oxidasedecarboxylase, and alcohol oxidase (AO). DAAO is typically assumed to utilize a carbanion mechanism (1), while AO has been proposed to utilize a radical (2). TMO catalyzes the same chemistry as DAAO, the oxidation of an amino acid, but also catalyzes an oxidative decarboxylation

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

142

(3). Our goals are to determine the structures of intermediates and transition states in the reactions catalyzed by these enzymes.

Results and Discussion D-Amino acid oxidase Scheme 1 shows the most accepted mechanism for the reductive half reaction of DAAO and most other flavoprotein oxidases (1). Generation of a carbanion by removal of the substrate a-proton is followed by attack of the carbanion at the flavin N(5) position. The adduct formed subsequently breaks down to generate reduced flavin and oxidized substrate. However, a direct hydride transfer mechanism is suggested by the transfer of the substrate a-proton to the flavin when 5-deazaFAD is used in place of the native FAD in DAAO (4). In addition, Miura and Miyake (5) have proposed a third mechanism in which removal of the substrate a-proton is concerted with transfer of a hydride from the substrate amino group to the flavin. Secondary kinetic isotope effects provide a straightforward means of distinguishing among these possibilities. The mechanism of Scheme 1 predicts that a primary deuterium kinetic isotope effect will be seen upon carbanion formation and a secondary effect will occur when the adduct breaks down in a subsequent step. In contrast, for any mechanism involving direct hydride transfer, the primary and secondary kinetic isotope effects will occur on the same step. Consequently, our approach has been to establish conditions where cleavage of the substrate CH bond is fully rate-limiting and to

Scheme 1

143

measure secondary deuterium isotope effects under these conditions. D-Alanine is the best characterized substrate for DAAO. However, at pH 9, the pH optimum, only very small isotope effects are seen on the V/Kaia value when a-[2H]-D-alanine is used as a substrate. This is because the rate-limiting step under these conditions is the initial encounter of the amino acid with the enzyme (6). Significant isotope effects can be measured if the pH is decreased, until a limiting value of about 5.5 is reached below pH 4 (Figure 1). To determine if this is indeed the value of the intrinsic deuterium kinetic isotope effect on cleavage of the amino acid CH bond, the primary isotope effect was determined with a-[3H]-D-alanine. At pH 4, this value is 12.6, consistent with a deuterium effect of 5.8 (7). The agreement between the measured deuterium effect and that predicted from the tritium effect (Table 1) is sufficiently good to conclude that CH bond cleavage is fully rate-limiting with D-alanine as substrate at pH 4. Consequently, ft-secondary effects were measured with this substrate at low pH. ftSecondary kinetic isotope effects result from changes in hyperconjuga-

5

-

3

-

7

11

PH Figure 1. Effect of pH on the primary deuterium kinetic isotope effects with D-alanine ( • ) and glycine (O) as substrates for DAAO.

144

Table 1. Intrinsic Kinetic Isotope Effects for DAAO substrate deuterium effect tritium effect alanine primary 5.8 12.6 ^-secondary 1.0 glycine primary 6.4 3.6 a-secondary 1.0 serine primary 4.5 8.6 tion between the ft-hydrogens and the carbon involved in bond cleavage (8). Hyperconjugation is unlikely if a carbanion is involved. However, for a hydride transfer mechanism significant ^-secondary effects can frequently be measured. For example, a value of 1.2 has been measured when ft,fi,JH2H]lactate is used as a substrate for lactate dehydrogenase (9), a reasonable model for the DAAO reaction. With DAAO, ft-deuterium isotope effects of 1.03 and 0.99 were measured at pH 4.5 and 5, respectively. Neither is significantly different from one, and both are significantly less than 1.2, consistent with the predictions if the mechanism of Scheme 1 is correct. Glycine is the only amino acid for which it is possible to determine an a-secondary kinetic isotope effect as a direct measure of the degree of rehybridization at the a-carbon of the amino acid. With DAAO the primary deuterium kinetic isotope effect on the V/K g i y value increases at high pH, reaching a limiting value of about 3.4 at pH 11 and above (Figure 1). The difference in this pattern from that seen with D-alanine is due to CH bond cleavage being reversible unless a group on the protein with a pKa value of 8.5 is deprotonated (10,11). Both the primary and secondary tritium kinetic isotope effects on the V/K g i y value were measured in a single experiment. The secondary effect was determined from the tritium content of the glyoxylate product, while the primary effect was determined from the tritium content of the water. At pH 10.8, the primary tritium effect was 6.4, consistent with a deuterium effect of 3.6. This is close to the value measured directly and confirms

145

that CH bond cleavage is rate-limiting with glycine at high pH. The asecondary tritium effect at pH 10.8 was 1.025±0.018. The calculated equilibrium secondary tritium effect is 1.5 (12), setting an upper limit on the rehybridization of the glycine a-carbon bond during CH bond cleavage of 8%. When a-[2H]-D-serine is used as substrate for DAAO the °V/Kser value of 4.5 is pH-independent. The deuterium isotope effect on the limiting rate of reduction measured in the stopped flow is 4.7. The primary tritium isotope effect on the V/K ser value is 8.6, consistent with a deuterium effect of 4.5. This establishes that CH bond cleavage is ratelimiting with D-serine at all pH values. It was therefore possible to measure the solvent kinetic isotope effect on CH bond cleavage. Concerted mechanisms predict that there will be a significant solvent effect on the CH bond cleavage step. In contrast, the measured value was not significantly different from one. With all three substrates for DAAO which we have examined in detail, it has proved possible to establish conditions where CH bond cleavage is fully rate-limiting. The secondary and solvent isotope effects measured under these conditions are fully consistent with the carbanion mechanism of Scheme 1 and provide strong evidence against mechanisms involving hydride transfer for transfer of the hydride equivalent from amino acid to flavin in DAAO. Tryptophan monooxygenase The effects of pH and isotopic substitution on V/K values for the reductive half reaction were determined with several amino acids as substrates for TMO. The V/K values for tryptophan and methionine decreased at high and low pH (Figure 2), consistent with a requirement that a group with an apparent pKa value of 5.1 be unprotonated and a group with a pKa value of 9.9 be protonated for activity. The pKa values of tryptophan was not seen in the profile, suggesting that the

146

protonation state of the substrate amino group is not critical for productive binding to TMO. Only small ( 10

8

10

12

pH Figure 2. Effect of pH on the V/K values for tryptophan (O) and methionine (•) with TMO.

147

effects of pH on the °V/K and V/K me t values Alanine is a very slow substrate for TMO, with a K m value of about 0.5 M. Fortunately, it is not necessary to saturate the enzyme with substrate to determine V/K values. Both primary and secondary kinetic isotope effects were measured with deuterated alanine at pH 8.3. The Q(V/K)aia value was 5.3, very close to the intrinsic value obtained with DAAO and consistent with fully rate-limiting CH bond cleavage with this substrate. The ^-secondary kinetic isotope effect was measured with [p,p,p-2H3]alanine; the value was 1.0, consistent with a carbanion intermediate. Indole acetamide is a competitive inhibitor versus amino acid substrates for TMO. Tight binding requires that a group with a pK a value of 6.0 be unprotonated and a group with a pK a value of 10.4 be protonated. These are the same groups that are seen in the V/K profiles; the pKa of 6 is decreased to 5 because of the stickiness of the substrates. The temperature dependence of this pKa value was determined. From the results an enthalpy of ionization of 6 kcal/mol could be calculated 6

5

E

4

> °

3

2

1 4

6

pH

8

10

Figure 3. Effect of pH on the primary deuterium kinetic isotope effect with methionine as a substrate for TMO.

148

for the amino acid residue involved. This value is consistent with a histidine as the active site base in TMO. Alcohol oxidase Sherry and Abeles (2) have proposed a mechanism for AO in which removal of the substrate hydroxyl proton to form the alkoxide precedes homolytic cleavage of the CH bond. We have determined primary and solvent kinetic isotope effects on V/KEtOH values for alcohol oxidase in order to establish whether the OH and CH bonds are cleaved in the same step. With several substrates, the pH dependency of the V / K R O H value showed that a group with a pK a value of 8.3 must be deprotonated. Therefore, the kinetic measurements were done at pH 9. As the value of the primary deuterium isotope effect increases, the value of the solvent effect decreases (Table 2). The most straightforward explanation for this result is that two different steps are being affected. In contrast, a mechanism in which OH bond cleavage is synchronous with hydride transfer would result in parallel changes in the values of the two isotope effects.

Table 2. Isotope Effects for Alcohol Oxidase substrate ethanol BrEtOH MeOH

PH 9.0 6.8 9.0 9.0

DV/KH20 1.54 1.62 4.74 6.06

D20V/K 1.94

°V/KD20 1.51

1.18 1.08

To examine the order of the two steps, additional isotope effects were measured with ethanol as substrate. The DV/K.ETOH value is pH independent, even at pH values well below 8.3. Therefore, either ethanol is not a sticky substrate and there is a significant internal commitment or the incorrectly protonated form of the enzyme will not bind the alcohol. There is no change in the W/KEtOH value in D2O. The simplest explanation for such a result is that the mechanism is concerted, but that possibility is ruled out by the results with the other substrates. If the pri-

149

m a r y deuterium effect is a kinetic effect on a separate step f r o m the solvent sensitive step, the CV/KBDH value must decrease in D2O as the solvent-step b e c o m e s slower. The d a t a can be reconciled if the ^ / K R O H value is an equilibrium isotope effect. In other words, cleavage of the CH bond is relatively rapid and freely reversible and is followed by slow cleavage of the OH bond. With 2-Br-ethanol a n d methanol, CH bond cleavage becomes increasingly rate-limiting and irreversible, so that little or no solvent effect is seen. While these results do not address the chemical nature of the CH bond cleavage, secondary isotope effects as described above for DAAO and TMO should d o so.

Conclusion Kinetic isotope effects have been used as mechanistic probes of several flavoprotein oxidases. Conditions have been established utilizing slow substrates or pH extremes which allow intrinsic kinetic isotope effects to be expressed on steady state kinetic parameters. Under these conditions, secondary and solvent isotope effects are not consistent with hydride transfer mechanisms for DAAO or TMO. Similarly, a combination of primary and solvent isotope effects do not support concerted OH and CH bond cleavage with AO. These approaches should be readily applicable to study of other flavoprotein oxidases.

Acknowledgments This work was supported in part by the National Science Foundation. PFF is an Established Investigator of the American Heart Association. References 1. Ghisla, S. 1982. In: Flavins and Flavoproteins (Massey, V. and C. H. Williams, Jr., eds.). Elsevier, New York. pp. 133-142

150

2. Sherry, B. and R. H. Abeles. 1985. Biochemistry 24, 2594-2605 3. Kosuge, T. 1970. Methods Enzymol. 1 7A, 446-449 4. Hersh, L. B. and M. S. Jörns. 1975. J. Biol. Chem. 250, 8728-8734 5. Miura, R. and Y. Miyake. 1988. Bioorganic Chem. 16, 97-110 6. Porter, D. J. T., J. G. Voet, and H. J. Bright. 1977. J. Biol. Chem. 252, 4464-4473 7. Swain, C. G., E. C. Stivers, J. F. Reuwerjr., and L. J. Schaad. 1958. J. Am. Chem. Soc. 80, 5885-5893 8. Cleland, W. W. 1987. In: Isotopes In Organic Chemistry, vol. 7 (Elsevier Inc., New York. pp. 61-113 9. Cook, P. F., N. J. Oppenheimer, and W. W. Cleland. 1981. Biochemistry 20, 1817-1825 10. Massey, V. and Nishino, T. 1980. In: Flavins and Flavoproteins (Yagi, K. and T. Yamano, eds.). University Park Press, Baltimore, pp. 1-11

11. Denu, J. M. and P. F. Fitzpatrick. 1992. Biochemistry 31, 8207-8215 12. Cleland, W. W. 1980. Methods Enzymol. 64, 104-125

Effects

of Ligands on the R e a c t i v i t i e s of Reduced and Semiquinoid Forms of

D-Amino Acid Oxidase

¥. Nishina,

K. Sato,

and K. Shiga

Department of P h y s i o l o g y , Kuaamoto U n i v e r s i t y School of Kuaaaoto, Kuaaaoto 860, Japan

Medicine,

Honjo,

Introduction Purple

interaediates

are

produced

in the r e a c t i o n of pig kidney D-aaino

a c i d o x i d a s e (DAO) w i t h neutral aaino a c i d s . provided

a p i e c e of e v i d e n c e t h a t the i n t e r a e d i a t e

DAO with the 2 - i a i n o a c i d in the produced

from

substrate

cationic

s p e c t r a (2).

fora

D-aaino a c i d (1).

p i c o l i n a t e and n i c o t i n a t e bind t o reduced

have

i s a complex of reduced

at

the

iaino

Previously,

DAO,

Here we report t h a t t r i g o n e l l i n e

and s e a i q u i o n i d DAOs, the

Resonance Raaan s t u d i e s

nitrogen,

we reported t h a t

producing

charge-transfer

i s a good l i g a n d f o r reduced

in a d d i t i o n to f o r o x i d i z e d DAO (3),

and a l s o r e p o r t

e f f e c t s of l i g a n d s on the r e a c t i v i t i e s of reduced and s e a i q u i n o i d DAOs

with O2 and other e l e c t r o n

acceptors.

R e s u l t s and D i s c u s s i o n The r e d u c t i o n of DAO by D - a r g i n i n e in the presence

of

f e r e n t f r o a the case of the absence of t r i g o n e l l i n e , s p e c t r u a having a t a i l

at long wavelength;

of reduced DAO-nicotinate coaplex (2). binds

to

reduced

DAO,

a

produces an a b s o r p t i o n

indicates

that

charge-transfer

trigonelline spectrua.

s p e c t r o p h o t o a e t r i c t i t r a t i o n with t r i g o n e l l i n e of reduced DAO D-arginine

(data

are

not

shown)

reduced DAO in a 1:1 molar r a t i o , nicotinate constant, nicotinate

(2).

The

indicated

that

s i a i l a r to the

relationship

Kd, and pH i s shown in Fig.

dif-

the spectrum i s s i a i l a r to t h a t

This

accoapanying

trigonelline,

between

the

The

prepared

by

t r i g o n e l l i n e binds t o

case

of

apparent

picolinate

or

dissociation

1; the pH p r o f i l e s f o r p i c o l i n a t e and

(4) are a l s o shown f o r comparison.

The p a t t e r n of the p l o t f o r

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

152

COO"

COO"

•tpcoo H N Fig.1: pH Effects on apparent dissociation constant of reduced DAO with trigonelline (T), picolinate nicotinate trigonelline shows that two ionizable groups participate i.e.,

one

region.

with

pK

in

the

acidic

(P),and

(N) at 25°C in

the

binding,

region and the other in the alkaline

The two ionizable groups associated with the binding of

trigonel-

line are probably on the apoprotein moiety, since the reduced flavin in DAO is

anionic and trigonelline is zwitterionic in the pH region studied.

pH profile for trigonelline is quite different from and nicotinate.

those

for

The

picolinate

Trigonelline is in the cationic form at the N atom even in

the high pH region studied, while picolinate and nicotinate have a dissociable proton at each N atom (pK values are 5.37 and 4.73, are neutral at the N atom in the high pH region.

respectively) and

Therefore, the pH effects

support that only the protonated cationic form at the N atom of

picolinate

or nicotinate can bind to reduced DAO.

Absorbance

traces

after

mixing

of

the

reduced DAO-solution containing

trigonelline and the benzoate-solution containing O2 were obtained by means of the stopped-flow method. The time course of the absorbance change at 560 nm within the charge-transfer absorption band was identical 462

nm

within

the

characteristic

kobs,

by

constant

O2.

The

at

these

observed

in the absence and the presence of trigonelline, as a

function of the concentration of O2, were obtained (Fig. rate

that

large band of oxidized flavin;

decays correspond to the process of the reoxidation rate constant,

with

2).

The observed

is linearly related to the concentration of O2.

were analyzed by the method reported (5). obtained are summarized in Table I.

The second order rate

These data constants

The value for free reduced DAO and the

value for the purple complex with D-alanine are consistent with those

153 T

150 -100

w XI o

50 »Li!

0,

0

200 [023.

A00

I

0

I

I

200

400

[02] ,

MM

p M

Fig.2: Observed reaction rate constants between reduced DAO and

as a

function of O2 concentration.

reported previously

(5).

(trigonelline, picolinate, free reduced DAO. DCIP,

The or

reactivity nicotinate)

of

reduced

with

DAO-ligand

complex

O2 is higher than that of

The reduction rate of ferricytochrome

c,

ferricyanide,

or methylene blue by reduced DAO was decreased by trigonelline.

The

reactivity of purple intermediate with O2 is higher than that of free fully reduced DAO (5), Therefore,

the

while the reactivity with cytochrome c reactivities

of

the

reduced

is

DAO-ligand

reverse

electron acceptors are similar to those of purple intermediates,

Table 1

Reaction Rate Constants Ligand

1 2 3 4 5 6 7 8

ee

(6).

complexes with suggesting

(k) of Reduced DAO with O2 k at pH 8. 3 1.5 x 10^

Ratio M ls

1

1.0 8. 7 10.7 20.6 14.7 12.0 14.7 22.7

2 ~ 5 , purple intermediates of DAO with D-amino acids (D-alanine, D-proline, D-pipecolic acid, and N-methylalanine); 7, at pH 7.0.

154 t h a t t h e complexes a r e good models f o r t h e e l u c i d a t i o n of t h e n a t u r e of purple

intermediate.

In r e d u c t i o n of DAO by d i t h i o n i t e reduced

to

s e n i q u i n o i d form,

in t h e p r e s e n c e of t r i g o n e l l i n e ,

DAO was

b u t was n o t r e d u c e d up t o r e d u c e d f o r m : t h e

s e n i q u i n o i d D A O - t r i g o n e l l i n e complex was h a r d l y r e d u c e d by d i t h i o n i t e . complex depressed

of

1:1

with

respect

to

the

enzyme.

t h e r e a c t i v i t y of s e n i q u i n o i d DAO w i t h C^,

of r e d u c e d DAO,

Trigonelline

contrary to the case

and d i d n o t a f f e c t t h e r e a c t i v i t y of s e n i q u i n o i d DAO

electron

acceptors,

methylene

blue.

such a s f e r r ¡ c y t o c h r o m e c,

ferricyanide,

f o r t h e l i g a n d t o have a c a t i o n i c group. of

a

l i g a n d or a s u b s t r a t e

The e n t r a n c e of

intermediate

and

is essen-

a

positive

in t h e v i c i n i t y of

may c o n t r i b u t e t o t h e m o d u l a t i o n of t h e r e a c t i v i t y w i t h e l e c t r o n

with

DCIP,

Nhen a l i g a n d b i n d s t o r e d u c e d DAO or a n i o n i c s e m i q u i n o i d DAO, i t charge

The

has a b r o a d s p e c t r u m in t h e l o n g - w a v e l e n g t h r e g i o n and t h e b i n d i n g

has a s t o i c h i o m e t r y

tial

the

flavin

acceptors.

Rerferences 1. N i s h i n a , Y. , K. S h i g a , M. Miura, H. Tojo, M. Ohta, Y. Miyake, T. Yanano, and H. W a t a r i . 1983. J. Biochem. 94. 1979-1990 2. N i s h i n a ,

Y., H. T o j o and K. S h i g a .

1986. J. Biochem. 99.,

3. N i s h i n a ,

Y., K. S a t o and K. S h i g a .

1990. J. Biochem.

4. N i s h i n a , 327-332

Y., H. Tojo, H. U s h i j i m a and K. Shiga.

5. P o r t e r , D . J . T . , 4464-4473 6. T a k a i , 1222

J.G.

Voet and H. J .

A., N. O h i s h i and K. Yagi,

Bright.

107,

673-680 726-731

1987. J. Biochem.

1977. J. B i o l .

102.

Chem. 252,

K. 1974. Anales de Quimica 70,

1219-

Thermodynamic Study of FAD Binding in D-Amino Acid Oxidase

F. Tanaka

Mie Nusing College, 100 Torii-cho, Tsu 514, Japan

Introduction D-amino acid oxidase from hog kidney is a flavoenzyme containing one mole of FAD per monomer

(Mw

400001.

The

enzyme is in an equilibrium state among monomer, dimer and higher origomers, depending on its concentration (11.

It is

found that the enzyme displays a positive cooperativity in the binding of the coenzyme, which is induced by drastic difference in the equilibrium constants of FAD dissociation from the enzyme species (2).

In the present work thermodynamic prope-

rty of the dissociation-equilibrium constant of FAD was investigated by changing both the enzyme concentration and solution temperature, by means of a steady-state fluorometry.

Results and Discussion Determination of Apparent Equilibrium Constant of FAD Dissociation Relative intensities (R^) of fluorescence of the enzyme solution to that of free FAD are shown in Figure 1. value of R.| increased as the concentration was diluted.

The It

also increased both when temperature was lowered from room tempetarure, and when it was elevated.

Fluorescence anisot-

ropy (A) of the enzyme solution at various levels of the enzyme and temperatures are shown in Figure 2. as the concentration was lowered.

It decreased

The anisotropy decreases

when FAD dissociates from the apoenzyme, since a rotational

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

156

1.

tr

0.

5 10 50 100 [Holoenzyne] (jM)

5 10 [Holoenzyne] ((M) Figure 1

Relative Intensity

Figure 2

Anisotropy

motion of FAD becomes faster upon the dissociation.

Relative

intensities of the fluorescence of FAD bound to the enzyme to free FAD in the enzyme solution were obtained by the following equation. R 2 = (A - A f ) / (Ab - A)

(1 )

where A^ and A^ are fluorescence anisotropies of free FAD and FAD bound to the enzyme. 0.4,

according

The value of A^ was assumed to be

to the reasons

described

elsewhere

(2,3).

With R^ and R2 apparent dissociation constants of equilibrium between FAD and the apoenzyme were determined by Eq (2). K = R 1 2 [F] 0 / (1 + R2)(1 + R 2 - Ri)

(2)

where [F] Q is total concentration of FAD, which is equal to the concentration of protein, [P]Q. Dependences of K on Temperature and the Enzyme Concentration van't Hoff plots of K at pH 8.3 are shown in Figure 3. The value of K increased at all temperatures as the concentration was diluted.

K increased both when temperature was

elevated from room temperature, and when it was lowered.

The

157 10-=

2

3. 3

3. 2

3. 5

3. 4 1000/T

3. 6

(K"')

Figure 3 v a n ' t H o f f P l o t of K C o n c e n t r a t i o n s (yM) : a 1.0, b 1.8, c 3.0, d 5.0, e 8.3, f 14, g 23, h 38, i 64, j 106 in 0 . 0 1 7 M p y r o p h o s p h a t e at pH 8.3.

values

of K w e r e

represented

with two

straight

l a r b e h a v i o r of K w a s a l s o s h o w n a t p H 7.0. py

change,

Table 1. the

two

AH°,

depended

AH° were

lines

- 1 8 - -21

intersect.

(kcal/mol)

creased yM,

as

from

the

(kcal/mol)

1.5 ( k c a l / m o l )

in the

temperatures yM.

The rather

at

at

a b o v e Tc.

a temperature concentration

further

106

yM

diluted.

t o 15

3.0

present

unusual.

yM

range

from

-5.4 ( k c a l / m o l ) a t 106

at temperatures above

results It is

on

oxidase

and

FAD

the

suggested

display

of

from

It

106 yM

inat 1

were

-20

to 23 y M

b e l o w Tc, a n d i n c r e a s e d to -12 ( k c a l / m o l ) from

in

which

(kcal/mol)

at 3

Tc.

tempetature-dependence that elementary

different

at

t o 5.9 is

equilibrium

c o n s t a n t s b e t w e e n t h e v a r i o u s a s s o c i a t e d s p e c i e s of acid

at

range

A t p H 7.0 t h e v a l u e s

concentration

It i n c r e a s e d

(kcal/mol)

enthal-

as s h o w n

b e l o w Tc, and i n c r e a s e d to -15

concentration

at temperatures

Apparent

A t p H 8.3 t h e v a l u e s

in the

1 0 6 t o 8.3 y M a t t e m p e r a t u r e s (kcal/mol)

Simi-

on the concentration,

In the T a b l e Tc r e p r e s e n t s straight

lines.

thermodynamic

D-amino proper-

158 Table 1 Constant conc. (UM) 106 64 38 23 14 8.3 5.0 3.0 1.8 1.0

Apparent of FAD

Enthalpy

pHa

A H 0 (k c a l / m o l ) T < Tc T > Tc

Tc (°C)

8.3 8.3 8. 3 8.3 8.3 8.3 8. 3 8.3 8.3 8.3

23 23 23 23 23 23 23 24 25 26

-18 -20 -21 -20 -18 -18 -17 -15 -14 -15

Change

of

Dissociation-Equilibrium

P

1 ,. 5 3., 6 4.,0 6.,6 6..6 8.,0 8., 7 8., 7 10 1 5

Hb

Tc (°C)

7.0 7.0 7.0 7.0 7.0 7.0 7.0 7.0

A H 0 (k c a l / m o l ) T < Tc T > Tc

1 7 1 7 1 7 1 7 18 19 19 19

-20 -20 -20 -20 -1 5 -1 5 -14 -12

-5.4 -2.9 -1.8 -1.8 1. 1 3.2 3. 1 5.9

a 0.015 M pyrophosphate buffer, b 0.1 M p h o s p h a t e b u f f e r .

ties.

At

relates

the

to a

present

transition

stage,

temperature

originally

proposed

parameters

of the r e s p e c t i v e

obtained

in

order

to

it

by

Massey

et

is of

not the

al.

the

whether

enzyme

(4).

equilibrium

elucidate

clear

confomation

Thermodynamic

constant

temperature

should

be

dependence

of

K.

References

1.

Tojo,

2.

T a n a k a , F , a n d K. Y a g i . 1 9 7 9 . B i o c h e m i s t r y ,

3.

T a n a k a , F., N . T a m a i ,

and

H., K. H o r i i k e ,

Yoshihara. 4.

K. S h i g a , Y. N i s h i n a , H.

T. Y a m a n o . 1 9 8 5 . J. B i o l . C h e m .

Massey, Chem.

1989.

V., B. C u r t i ,

247:2347.

Watari,

260:12615. 75:1531.

I. Y a m a z a k i , N . N a k a s h i m a ,

Biophys. and

J.

a n d K.

56:901.

H. G a n t h e r .

Tc

1 9 6 6 . J.

Biol.

Structure of Human D - Amino Acid Oxidase Gene

Kiyoshi Fukui Department of Biochemistry, National Cardiovascular Center Research Institute, 5 - 7 - 1 , Fujishirodai, Suita, Osaka 565, JAPAN

Introduction Since the initial characterization of D - Amino- acid oxidase (EC 1.4.3.3, DAO), a flavoenzyme with FAD as its prosthetic group catalyzing the oxidative deamination of a wide range of D - amino acid, extensive studies on the enzymatic characteristics of DAO have been carried out and well documented. However, little is known about the regulation of DAO expression, especially the mechanism underlying its induction or suppression at the molecular level, which appears to provide the molecular basis of the physiological function of DAO. In this study, in order to examine the structural organization of the human gene and the regulation of its expression in a search for some clues as to the physiological function of DAO, the isolation of the human DAO gene and the determination of its structure have been carried out. In addition, polymorphic microsatellite DNA in the human DAO gene is also described. Results and Discussion The genomic clones covering the entire sequence of the human gene encoding DAO have been isolated from human placental genomic libraries using a previously cloned cDNA for human DAO (1) as a probe. The human genome contains a single copy of the DAO gene, comprising 11 exons and spanning 20 kb. The presence of a 5 ' - noncoding exon (exon U) at about 5 kb upstream of the translation initiation codon was demonstrated, and a putative hydrophobic FAD - binding region of the enzyme is encoded by exon 1 in the human genome. Multiple initiation sites for human DAO transcription were identified by primer extension analysis, which is consistent with commom structural properties in the 5'-flanking region of the

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

160

••••••••••

DAO

[-247]

— GGATCCTCTCCAGCTGGCCAGAGAGACAGACCTTCTT

ACTH

[ -1971-568]

— GAGCAGGCCCCAGGGGAGCAGTGCAACTCACCTTCAC

••••• GTGCTCATCAACCCTCCAAGAATGCCTGCCCTCCCTCCTTCCCCCAAGGCCTGTCCACAG * W W * W*W* * * * * ** ***** W W W W W W * w WW W ACCCACAAGACGGCTCC-TGACTTCTGCTCCCTCCTCCCCTCCCCAAAG-TGGAACAGA-

ii

vwwvw GGGCTTGAGATCAGCCAGAAAAGTCA-GGCA-ACTTTTCAGGGACT * * * w w w w w w * ** ** * *** ** WW

GGGAGC-GAGG * * * www*

GAGAATATGATTCCCCACGACTTCCACAJCACAGTTTCCAAACAATGGGGAAATCGGAGG

oooo

oooooooo

TCT-CCCG GCCGGGCCTGGGTCCAGTCTCTGTGGGCAGTGCAGTGCCGAGCCCC w w wwww * *** * * * * * w ww w **** w w w w w w w CCTCCCCGTGTGCAGACGGTGATATTTACCGCCAATGCGAACCJAGGCAGATGCCAGCCCC ACCCCTCAAGC-CGTGCCCTGTCCATAGCTCCAGACTTTGACCC— [ - 1 ] * W W WW WW W W WW WW w * * ** AGCACGCACGCAGGTAACTTCACCCTCGCCTCAACGACCTCAGA—{+61/-311 ]

Figure 1. Alignment of nucleotide sequences within 5 ' - flanking regions; asterisks indicate nucleotide identical in both DAO and ACTH sequences; open triangle, closed and open circles indicate cAMP responsive element - like sequence, glucocorticoid responsive element and 1 2 - bp (G+C)- rich region, respectively; arrowhead indicate alternate transcription initiation site of ACTH gene. Negative numbers indicate the distance upstream of the transcription initiation site. DAO gene, namely the absence of well - defined TATA and CAAT boxes. When the 5 ' - flanking region of the human DAO gene was compared with the nucleotide sequence data bases to see whether or not it shared common regulatory sequences, homology between the human DAO and human corticotropin

/3 - lipoprotein

precursor (2) genes was found to be significant immediately upstream of the transcription initiation site as shown in Figure 1 (51.4 % over 247 bp). In addition, a sequence including 7 nucleotides in common with 1 2 - nucleotide consensus glucocorticoid responsive element (3), 1 2 - bp (G+C) - rich region, and a sequence resembling that of a canonical cAMP - responsive element were also found in the 5 ' - flanking region (Figure 1). Moreover, two sequences , (CA) 20 and (CA)„, of alternating pyrimidine and purine nucleotides, which have the potential to form Z -

161 Caucasian

Black

Oriental

PCR-Amplified Fragment

Figure 2. Autoradiograph of DNA sequencing gel analyzing P C R - amplified fragments derived from 21 different human genomic DNA templates, Caucasian, B l a c k and Oriental, using primers surrounding C A repeats in the first intron of the human DAO gene. D N A (4), are present in the first intron.

This repeat was reported to show the

genetic polymorphism due to the variation in the length of the repeating unit among individuals (5). Caucasian,

Therefore, 21 independent human genomic DNAs, comprising

Black

and

Oriental

populations,

were

heterogeneity in the sequences around the C A repeat

analyzed

to

search

for

microsatellite DNA.

As

shown in Figure 2, P C R amplified fragments derived from the human DAO genes of different individuals exhibited significant polymorphism in the length when analyzed on denaturing polyacrylamide gel electrophoresis.

There are three other

potential

first

sequences

for

genetic

polymorphism

in

the

intron,

namely

pentanucleotide A l ' I ' l T and 2 poly T tract. The human D A O turned out to be an

162 abundant source of genetic markers. The human DAO gene locus was assigned to chromosome 12 by screening a panel of Chinese hamster and human somatic cell hybrids with unique sequence primers for PCR analysis. Whether or not the DAO gene is linked to any genetic diseases would be interesting to determine in view of the unknown physiological function of this enzyme.

Acknowledgements This research was supported in part by a Grant- in- Aid for Scientific Research (05670159) from the Ministry of Education, Science and Culture of Japan, Research Grant 91A1202 for Aging and Health from the Ministry of Health and Welfare of Japan and a grant from Yamanouchi Foundation for Research on Metabolic Disorders.

References 1. Momoi, K., Fukui, K., Watanabe, F., Miyake, Y. 1988. FEBS Lett. 238: 180 2. Cochet, M., Chang, A. C. Y., Cohen, S. N. 1982. Nature. 297: 335 3. Beato, M. 1989. Cell 56: 335 4. Nordheim, A., Rich, A. 1983. Proc. Natl. Acad. Sci. U. S. A. 80: 1821 5. Weber, J. L., May, P.E. 1989. Am. J. Hum. Genet. 44: 388

AMINO ACID SEQUENCE OF D-AMINO ACID OXIDASE FROM THE YEAST Rhodotorula gracilis

Ludovica Faotto*, Loredano Pollegioni*, Fabrizio Ceciliani*, Giovanni Gadda^, Severino Ronchi* and Mirella S.Pilone*. •Institute of Veterinary Physiology & Biochemistry and C.I.S.M.I. and ^Department of General Physiology & Biochemistry, University of Milano, Milano, Italy.

Introduction

R. gracilis D-amino acid oxidase (DAAO,1.4.3 .3) is a flavoprotein of major interest in basic research and in biotechnology (1). The enzyme possesses a tight binding with FAD (Kd = 2.0xl0"8 M) (2) and a high catalytic efficiency (3). Determination of the sequence of this flavoprotein has thus been addressed to obtain oligonucleotides for the screening of cDNA library for R. gracilis. The cloning of the enzyme gene is in fact a major issue in order to obtain overexpression of the protein as well as to perform site-directed mutagenesis studies. This report presents the experimental work that allowed characterization of fragments derived from trypsin digestion and their alignment by sequence similarity with other flavooxidases.

Results and Discussion Determination of the primary structure has been approached through sequence analysis of the intact protein and of tryptic fragments after reduction and carboxymethylation. N-terminal sequence analysis carried out on native R. gracilis DAAO brought to the identification of a single amino acid derivative through 30 degradation cycles giving the following sequence: MHSQKRVWLGSGVIGLSSALILARQGYSV Separation of the tryptic peptides was accomplished by HPLC chromatography as previously reported (4). Amino acid composition and sequence analysis of the material present in the different peaks allowed us to identify 18 tryptic peptides out of the 30 expected on the basis of the amino acid composition of the protein (5). Their primary structure is reported in Table 1.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

164 MHSQK RWVLGSGVIGLSSALILAR KGYSVHILAR DLPEDVSSQTFASPWAGANWTPFMT... KWVELVPTGXAMXL.. FAQNEDGLLGHWYK DITPNYRPLPSSEIPPGAIGVTYDTLSVXAPK TVTSLEQAFDGADLWNATGLGAK SIAGIDDQAAEPIR GQTVLVK CTXDSSXPASPAYIIPRPGGEVIQGGTYGVGDWDLSVNPPET... LDPTISSDGTIEGIEVLR HNVGLRPAR EK EVTLVHAYGFSSAGYQQSWGAAEDVAQLVDEAFQ...

Table 1. Amino acid sequence of the tryptic peptides from R.gracilis D-amino acid oxidase

The reported sequences account for 270 amino acid residues corresponding to about 77% of the total number. N-terminus of the protein presents a GXGXXG motif common to the flavin binding site. This particular region can fold into a PaP unit for the binding of ADP moiety of FAD as predicted by Wierenga et al. (6). It is interesting to note that Asp37 (the numbering is always referred to pig enzyme sequence) is highly conserved in all DAAOs and D-aspartate oxidase and it could play a role similar to Glu50 in glutathione reductase (7). Tryptic peptide alignment has been performed by homology comparison with other DAAOs, expecially those from microorganism source. From this comparison a highly conserved region (171-205) has been identified. The role of the carboxyl group of Asp157 in glycollate oxidase, Asp180 in lactate oxidase and Asp282 in flavocytochrome

165

..*.*.****..**. p 1 h r m f t rg

.

*

.

*..

MR VWIGAGVIGLSTALCIHERYHSVLQPLDVKVYADRFTPFTTTDVAAGL MR VWIGAGVIGLSTALCIHERYHSVLQPLHIKVYADRFTPLTTTDVAAGL MR VWIGAGVIGLSTALCIHELYHSALQPLDMTIYADRFTPLTNTDVAAGL MR VAVIGAGVIGLSTALCIHERYHPT-QPLHMKIYADRFTPFTTSDVAAGL MS—NTIVWGAGVIGLTSALLLSKNKGNKITWAKHMPGD-YDVEYASPFAGAN MA—K-IWIGAGVAGLTTALQLL-RKGHEVTIVSEFTPGD-LSIGYTSPWAGAN MHSQKRVWLGSGVIGLSSAL-ILARKGYSVHILARDLPEDVSSQTFASPWAGAN

p 171-205 h r m f t rg

....*..*..

.

*

****..*

ARGGADVIINCTGVWAGVLQ—PDPLLQPGRGQIIKV AREGADVIVNCTGVWAGALQ—RDPLLQPGRGQIMKV AGGGVDVIVNCTGVWASALQ—PDPLLQPGRGQIIKV AR-GVDVIINCTGVWAGALQ—ADASLQPGRGQIIQV AGKTPNIIVNATGLGSYKLGGVEDKTMAPARGQIVW SGSRPDVIVNCSGLFARFLGGVEDKKMYPIRGQWLV AFDGADLWNATGLGAKSIAGIDDQAAEPIRGQTVLV *. .

**

.

. . *

p 226-256 h r m f t rg

SPYIIP—GLQAVTLGGTFQVGNWNEINNIQDH SPYIIP—GTQTVTLGGIFQLGNWSELMNIQDH SPYIIP—GVHAVTLGGIFQMGNWS EGNSTDDH SPYIIP—GSKTVTLGGIFQLGNWSGLNSVRDH VMYLMQRAAGGGTILGGTYDVGNWESQPDPNIA ALYIMTRF-DGTSIIGGCFQPNNWSSEPDPSLT PAYIIPR-PGGEVIQGGTYGVGDWDLSVNPPET * ** *

p 278-286 h r m f t rg

EYTGFRPVR EATGFRPVR EWTGFRPVR ELTGFRPVR HAVGMRPWR ECVGHRPGR HNVGLRPAR

p 302-330 h r m f t rg

..*.**..*

**.

NTEVIHNYGHGGYGLTIHWGCALEVAKLF NTEVIHNYGHGGYGLTIHWGCALEAAKLF KTEVIHNYGHGGYGLTIHWGCALEAAKLF SAEVIHNYGHGGYGLTIHWGCAMEAANLF ETWIVHNYGHSGWGYQGSYGCAENWQLV VGFWHNYGAAGAGYQSSYGMADEAVSYV EVTLVHAYGFSSAGYQQSWGAAEDVAQLV

Figure 1. Homology comparison between D-amino acid oxidases from various sources (4). p: pig; h: human; r: rabbit; m: mouse; f: Fusarium solani, t: Trigonopsis variabilis, Rhodotorula gracilis.

* identity

• similarity

rg:

166 b2 is to stabilize the positively charged group of hystidine during catalysis (8): a similar role might be proposed for Asp192. The 226-256 region contained the conserved Tyr228, followed by a sequence GGTYGVGDW in R.gracilis D-amino acid oxidase (see Figure 1), which can be identified as a second FAD binding region as proposed by Eggink et al. Asp246 could be hydrogen bonded with the O3 group of the ribityl portion of the flavin moiety. A third homologous region is centered around Arg283 which is present in all DAAOs: the role of this residue is discussed in an accompanying paper (see Gadda et al., this Symposium). Finally, a high similarity is evident in the region 302-330 (see Fig. 1), encompassing His307 which is considered to be the active site base responsable for the abstraction of the a hydrogen as a proton from the substrate. It can be concluded that R.gracilis D-amino acid oxidase maintains the general structural features of flavin oxidases. However, some amino acid residues considered mechanistically important in all so far studied oxidases, are present in the R.gracilis enzyme, whereas other residues for which a role in catalysis has been previously discussed (Tyr55, Met 110 , Lys204, His217 and Tyr224) are not conserved.

Acknowledgements We thank Dr. A. Negri for critical comments. This work was supported by grants from M.U.R.S.T. and C.N.R. Target project "Biotechnology and Bioinstrumentation".

References 1. Pilone, M.S., L. Pollegioni and S.Butö. 1992. Biotech.Appl.Biochem. 16, 252-262 2. Casalin, P., L. Pollegioni, B. Curti and M.S, Pilone. 1991. Eur.J.Biochem. 197, 513-517 3. Pollegioni, L., B. Langkau, W. Tischer, S. Ghisla and M.S. Pilone. (1993) J.Biol.Chem. (in press) 4. Negri, A., F. Ceciliani, G. Tedeschi, T. Simonie and S. Ronchi. 1991. J. Biol. Chem. 267, 11865-11871 5. Pilone Simonetta, M., L. Pollegioni, P. Casalin, B. Curti and S. Ronchi. 1989. Eur.J.Biochem. 180, 199-204 6. Wierenga, R.K., P. Terpstra and W.G.J. Hoi. 1986. J. Mol. Biol. 187, 101-107 7. Karplus, P.A. and G.E. Schulz. 1987. J.Mol.Biol. 195, 701-707 8. Ghisla,S. and V. Massey. 1991. In: Chemistry and Biochemistry ofFlavoenzymes, Vol. 2 (F. Müller, ed.). CRC press, Boca Raton, pp. 243-289

CHEMICAL MODIFICATION OF ARGEVINE GROUPS IN D-AMINO ACID OXIDASE FROM Rhodotorula gracilis Involvement in catalysis and assignment in the sequence

G. Gadda*, A. Negri* and M.S. Pilone* 'Department of General Physiology & Biochemistry, "institute of Veterinary Physiology & Biochemistry, University of Milano, Milano, Italy

Introduction D-amino acid oxidase (DAAO, 1.4.3.3) from the yeast R. gracilis is a flavoenzyme which performs the oxidative deamination of D-amino acids with an unusually high catalytic efficiency (1). This DAAO shares with the class of flavoprotein oxidases several properties attributed to the existence of a positively charged group near the flavin locus N(l)-C(2)=0 exerting an inductive effect. This residue has been demonstrated by crystallographic studies to be a lysine in flavocytochrome b2 and glycolate oxidase (2,3) and a histidine in glucose oxidase (4). In pig kidney DAAO it has been inferred that this residue is arginine not identified in the protein sequence (5,6). Lacking the 3-D structure of the DAAO and as a prerequisite to sitedirected mutagenesis studies, we investigated a possible role of arginines using the chemical modification approach. We here report the first instance in which direct location of the basic residue interacting with the FAD N(l)-C(2)=0 locus in different flavooxidases has been identified in the amino acid sequence by chemical modification.

Results and Discussion Treatment of the holo-DAAO with phenylglyoxal (PGO), an arginine-specific reagent, in borate buffer pH 8.0

at 25°C caused a time-dependent loss of activity (Fig.l). The

inactivation followed pseudo-first-order kinetics with a biphasic pattern. Both phases were dependent on reagent concentration; the fast phase was completed within 1 minute and its extent was linearly dependent on PGO concentration. The slow phase was linear up to

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

168

Time

(hours)

Fig.l. Time-course of inactivation of holo-D-amino acid oxidase withphenylglyoxal. 3.2 JIM DAAO was incubated with PGO in 150 raM borate pH 8.0, 10% glycerol, 5 mM 2mercaptoethanol and 2 mM EDTA at 25°C in the dark. [PGO] was 2.32 mM; (A.) 3.10 mM; ( • ) 4.64 mM; (O); 6.29 mM and (O) 10.03 mM. Inset: replot of the apparent first order slow inactivation rate versus phenylglyoxal concentration. complete enzyme inactivation. The second-order rate constants of inactivation for the slow phase was 1.55 M-'min-', while that for the fast phase was not determined due to the very high rate of inactivation at 25°C. The reaction order with respect to [PGO] was 1. PGO modification of DAAO was irreversible; no change in enzyme specific activity was observed after removal of excess of reagent or after stopping the reaction with L-arginine. Due to the instability of the apoprotein in borate buffer at 25°C, the kinetics of inactivation by PGO was investigated in phosphate buffer pH 8.0 at 25°C. Apoenzyme inactivation by PGO was a timedependent process having the same biphasic pattern observed in the holo form. The only significant difference between the apo and holo enzyme was that the slow phase inactivation by PGO was faster for the apo form. The second-order inactivation rate constants for the slow phase were 18.0 M^min"1 and 8.3 M"1min"1 for apoprotein and holoprotein respectively. The presence of high concentrations of benzoate, a competitive inhibitor of holo-DAAO, during holoenzyme incubation with PGO strongly protected the enzyme in the second phase of inactivation, without affecting the extent of the phases (Fig.2). A plot of the reciprocal of the inactivation rates versus benzoate concentrations in the Kd range gave a straight line, indicating competitive inhibition of PGO by benzoate. This result suggests that PGO acts at the active site of DAAO and, moreover, that the fast and slow reacting residues probably have

169

Time (hours) Fig.2. Phenylglyoxal ¡motivation of D-amino acid oxidase in the presence and absence of benzoate. 3.2 jxM DAAO was incubated with 10.03 mM PGO in 150 mM borate pH 8.0, 10% glycerol, 5 mM 2-mercaptoethanol and 2 mM EDTA at 25°C in the dark, (.A.) without benzoate; ( • ) 20 mM benzoate. different mechanistic roles in the enzyme active site. The enzyme modified during fast phase inactivation with PGO could not be recovered by gel filtration, whereas the modified holoenzyme isolated during slow phase inactivation was stable, soluble and maintained its characteristic absorption spectrum with peaks at 460, 380 and 274 nm. Steady-state measurements of KM and VMAX on 90% inactivated DAAO showed that partially modified forms of the enzyme were not present. The modified enzyme could not be reduced by D-alanine under the anaerobiosis. The inactivated enzyme was not able to produce the N(5) adduct with sulfite, suggesting that PGO acted on the protonated group near the flavin N(l)-C(2)=0. PGO-apoprotein was prepared and the modified apoenzyme retained the ability to reconstitute the holo form in the presence of exogenous FAD. Reconstitution experiments using 8-SH-FAD are under investigation. The results point to the presence of an arginine at the flavin locus N(l)-C(2)=0, now covalently modified by PGO. This result is different from that obtained with pig kidney DAAO, where PGO modified one essential arginine involved in the binding of FAD ribityl moiety (7). Amino acid analyses and [14C]-PGO incorporation indicated that only one arginine was modified during the fast phase and that, when the enzyme was completely inactivated, a total of 4 arginines were modified. To identify the residue(s) modified by PGO, DAAO was reacted with [14C]-PGO (1.91 nCi/nmol) both in the presence and absence of high benzoate concentration. Samples were digested with trypsin. The peptides were separated by HPLC on an Aquapore RP-300 (Brownlee) column. Comparison of the two chromatograms and

170 evaluation of [ 14 C]-PGO incorporation showed the presence of a radiolabeled peak in the sample incubated without benzoate. This peptide was missing in the tryptic map when benzoate was present in the incubation. The peptide was 9 residues in length, and the determined sequence was: HNVGLRPAR * PGO-modified Comparison of the amino acid sequence of the arginine-containing peptide with the primary structure of pig kidney DAAO allows us to conclude that the PGO modified arginine corresponds to Arg 283 in pig kidney DAAO (see Faotto et al., this Symposium). This work gives evidence that, in the R.gracilis DAAO, the positively charged group near the flavin N(l)-C(2)=0 is an arginine residue; we have identified this residue in the partially available amino acid sequence of the enzyme. The importance of this arginine emerges from comparison of the known sequences of DAAO from different sources, this residue is conserved in all proteins and it belongs to a peptide sequence which shares a high degree of similarity with the other known sequences.

Acknoledgements This work was supported by grants from Italian M.U.R.S.T. and from the C.N.R. Target Project "Biotechnology and Bioinstrumentation".

References 1. Pollegioni, L., B. Langkau, W. Tischer, S. Ghisla and M.S. Pilone. 1993. J.Biol.Chem. (in press) 2. Xia, Z. and F.S. Mathews. 1990. J.Mol.Biol. 212, 837-863 3. Lindvist, Y. 1989. J.Mol.Biol. 209, 151-161 4. Hecht, H.J., H.M. Kalisz, J. Hendle, R.D. Schmid and D. Shomburg. 1993. J.Mol Biol. 229, 153-172 5. Ferti, C., B. Curti, M.S. Pilone, S. Ronchi, M. Galliano and L. Minchiotti. 1981. Eur.J. Biochem. 119, 553-557 6. Fitzpatrick, P.F. and V. Massey. 1983. J.Biol.Chem. 258, 9700-9705 7. Vanoni, M.A., M. Pilone Simonetta, B. Curti, A. Negri and S. Ronchi. 1987. Eur.J.Biochem. 167, 261-267

FUNCTIONAL AND STRUCTURAL ASPECTS OF D-AMINO ACID OXIDASE FROM Rhodotorula gracilis PROBED BY LIMITED PROTEOLYSIS

Loredano Pollegioni*, Fabrizio Ceciliani^, Bruno Curti*, Severino Ronchi# and Mirella S. Pilone* ""Department of General Physiology and Biochemistry and ^Institute of Veterinary Physiology and Biochemistry and C.I.S.M.I., University of Milano, Milano, Italy

Introduction D-amino acid oxidase (DAAO, EC 1.4.3.3) from the yeast Rhodotorula gracilis is present in the peroxisomes under induction conditions (1). The purified enzyme in its native holoenzyme form is a homodimer of 79 kDa, while the apoprotein exists as a 39 kDa monomer. No active DAAO monomelic form has so far been detected due to the rapid shift to a dimeric aggregation state when the apoprotein is reconstituted with coenzyme FAD (2). In this paper we report on the use of limited proteolysis performed on yeast apo- and holo-D-amino acid oxidase to investigate the structural requirements for catalytic activity of the enzyme, binding of the coenzyme FAD and protein monomer-monomer interactions.

Results Correlations between activity and proteolytic process were studied for DAAO, DAAObenzoate complex and apo-DAAO in the presence of 10% (w/w) trypsin. The kinetics of inactivation was in all cases a biphasic process, the rate and the extent of inactivation being different for the three enzyme species (Fig. 1). The presence of benzoate strongly protected the holoenzyme. The correlation between time course of inactivation and proteolytic pattern (as indicated by densitometric analysis of SDS-PAGE slabs) revealed the presence of a biphasic proteolysis process: the first rapid phase was concomitant with progressive digestion of the 39 kDa protein (the intact enzyme monomer) and the appearance of a protein band with a molecular weight of 35.7±0.7 kDa (see inset Fig.l). The inactivation rate of the second slower phase corresponds to degradation of the 36 kDa fragment.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

172 Structural studies were performed under conditions in which the 36 kDa fragment accumulated (holoenzyme-benzoate complex treated with 3% (w/w) trypsin at 25°C for 45 min). This fragment, isolated by gel filtration under native conditions, showed the typical flavoprotein spectrum and an apparent molecular weight of about 36 kDa. On the basis of the value of 79 kDa for the native dimeric enzyme, the truncated enzyme was thus present in a monomelic aggregation state. The N-terminal sequence of the 36 kDa fragment was identical to that of the native protein, indicating that the 3 kDa peptide(s) was splitted from the Cterminus region of the enzyme.

The 36 kDa truncated enzyme was a fully active form,

indicating that the C-terminus portion is not involved in catalysis. The apoprotein prepared from the 36 kDa holo-fragment was fully reconstitutable in the presence of a large excess of FAD. The equilibrium binding of FAD to apoprotein was measured by the quenching of both FAD and protein fluorescence: a Kd of 1.2xl0"8 M was determined, a value quite similar to that of the native enzyme (2.0xl0-8 M) (2).

Figure 1. Proteolytic inactivation by 10% trypsin (w/w) at 25°C of R.gracilis D-amino acid oxidase. ( • ) Holoenzyme; (A) holoenzyme-benzoate complex; (®) apoprotein. Inset: time courses of proteolytic cleavage of native holoenzyme-benzoate complex (39 kDa) DAAO ( a ) and of formation of the 36 kDa truncated form ( # ) followed by densitometry analysis of SDS-PAGE slabs.

173 The purified 36 kDa fragment showed the typical flavoprotein spectrum with absorbance maxima at 455, 378 and 274 nm (Figure 2). A flavin absorption coefficient of 10,140 M-'cnv1 was determined for the truncated enzyme form. When benzoate and anthranilate were bound to the truncated forms of R. gracilis DAAO they produced typical resolved spectra (Figure 2): Kd values, determined by absorption-difference spectroscopy, were similar to those previously determined for the native enzyme (3). The specific activity of the 36 kDa holo-enzyme with D-alanine and D-proline as substrates was 90% of the native enzyme, with no significant changes in the corresponding K,,, values. The truncated form did not require exogenous FAD for maximum activity pointing to the existence of an intact functional domain with full FAD binding capacity. However, when the thermodynamic parameters for the thermoinactivation process were determined, significantly lower thermostability was observed for the 36 kDa truncated form compared to the native enzyme.

.10

a> o c to

°

o CO

.05

jO «
2 + H 2 Q2 In addition, 2-nitropropane, nitroethane, 3-nitro-2-pentanol, and nitrocyclohexane are good substrates. The enzyme loosely binds with an oxidized form of FAD, and easily releases this bound FAD from protein moiety to form an apoenzyme during purification. The apoenzyme still contains a tightly bound chromophore absorbing at 340 nm. We here describes characterization of this 340-nm chromophore.

Results and Discussion The absorption spectrum of the purified enzyme does not show maxima at 380 and 450 nm, which are characteristic of an oxidized form of FAD (Fig. 1 (A)). The purified enzyme was inactive in the absence of FAD, showing that it is an apo form. The absorption spectrum shows a shoulder at 340 nm (Fig. 1 (A)). This chromophore is bound with the protein moiety so tightly that it was not released during the purification of the enzyme and by dialysis against 1 M KBr. We denatured the enzyme with 8 M urea in

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

196

1.5

0

240 300

500

Wavelength (nm)

700

Fig. 1. Absorption spectra of nitroalkane oxidase before and after urea treatment. (A) The enzyme (1.6mg/ml enzyme, 0.1 M Tris-HCl (pH 8.0)). (B) The enzyme after 8 M urea treatment (1.6 mg/ml enzyme, 0.1 M Tris-HCl (pH 8.0), 8 M urea). (C) The high molecular weight fraction of the urea denatured enzyme. (D) The low molecular weight fraction of the urea denatrued enzyme.

order to dissociate the chromophore from the protein moiety, and found that the absorption at 340 nm decreased, and the absorption maxima appeared at 380 and 450 nm (Fig. 1 (B)). Thereafter, we subjected this enzyme solution to ultrafiltration with Amicon Centricon-10 to obtain high molecular weight and low molecular weight fractions. The high molecular weight fraction did not show an absorption shoulder at 340 nm; it is a simple protein (Fig. 1 (C)). The low molecular weight fraction showed absorption maxima at 260, 380, and 450 nm (Fig. 1 (D)). This spectrum is closely similar to those of oxidized FAD, FMN, and riboflavin. We compared properties of the chromophore in the low molecular weight fraction with those of oxidized FAD, FMN, and riboflavin. The fluorescence spectrum of the chromophore was indistinguishable from those of oxidized FAD, FMN, and riboflavin: the excitation spectra showed maxima at 380, 450, and 470 nm (emission: 530 nm), and the emission spectra showed a maximum at 530 nm (excitation: 450 nm). We subjected these compounds to reverse phase high-performance liquid chromatography (column: Cosmosil

197 5 C j g - A R (4.6 x 150 mm); solvent: methanol + 10 mM NaH2PC>4 (pH 5.5) (35:65)). The chromophore was identical to oxidized FAD, but not to oxidized FMN and

riboflavin.

Therefore, we examined the coenzyme activity of the chromophore with apo-D-amino acid oxidase, which requires FAD as a coenzyme. The chromophore certainly showed a coenzyme activity. The amount of FAD in the low molecular weight fraction determined with apo-D-amino acid oxidase was identical to that of FAD determined by fluorometric analysis: 13 nmol of FAD was recovered from 10 nmol of tetramer enzyme. From these results, we concluded that the chromophore released from the enzyme is an oxidized form of FAD. Judging by the absorption spectrum of the enzyme (Fig. 1 (A)), the bound chromophore is not the oxidized form of FAD. However, the oxidized FAD was obtained from the enzyme not only by 8 M urea treatment as described above, but also by 7 M guanidine-HCl

0.4

0.4

-

Omin i Anaerobic 25min i Aerobic 30min

D o §

XI 11 0.2

0.2

o c/j


0.2 6.5 x 1.9 x 1.1 x 4.6 x

5.6 x 10-s 6.4 x 10-4 9.3 x 10-4 3.2 x 10-8 4.2 x 10-6 3.6 x 10-8 7.4x10-8

-

10 -5

10-5 10-3 10-2 10-5

Substitution of the residues which are postulated to interact with the substrate led to lowered turnover rates (Table 1). However, the binding of substrate is similar to that found in wild type enzyme or tighter. It seems that these amino acid residues contribute to catalysis mainly by

206 stabilizing the transition state of the reaction. This is derived from the dissociation constants measured with oxalate (Table 2). Oxalate is postulated to mimic the transition state of the reduction step in lactate monooxygenase (8,22). Fig. 3 shows that a linear relationship occurs between the tightness of binding of oxalate and the rate of reduction. All of the mutant enzymes which show a decreased rate of reduction, with the exception of H290Q, bind oxalate less tightly.

The linear relationship is also an indication that the structure of the mutant enzymes has not been significantly altered by the mutation. Thus the measured properties can indeed be attributed to the amino acid replacements. Even K266M, for which only a weak binding of both L-lactate and D-lactate has been determined, lies within experimental error of the established plot. The only mutant which does not fit the relationship is H290Q, underscoring its functional difference. The active site base has been removed and the residual activity must proceed by an altered mechanism. It should be emphasized, however, that flavin reduction by substrate with this enzyme is seven to eight orders of magnitude slower than with wild type enzyme, illustrating the importance of histidine 290 to catalysis.

For two of the mutations, Y44F and R293K, the reduction step is no longer rate limiting (Table 1). This implies that additional steps during turnover have been affected such as the release of pyruvate. Both show a substantial percentage of the reaction proceeding according to the

207 uncoupled pathway, again indicating a rapid release of pyruvate compared to the reduction of oxygen by the reduced enzyme-pyruvate complex. Reoxidation in the absence of pyruvate, however, has not been affected (Table 1). Y152F shows a rate of reoxidation similar to that found in wild type enzyme. It could also be shown that the presence of pyruvate enhances the rate of reoxidation of Y152F to 106 M-1 s-1 (wild type enzyme: 1.8 x 106 M-1 s-1). In keeping with this, turnover is only about 20-30 % uncoupled. This may explain why the rate of flavin reduction of Y152F is 70 fold slower than that in Y44F, yet kcat is four fold faster than with Y44F. C203A behaves similarly to the wild type enzyme in all the properties examined. The chemical modification of C203 by FDNB presumably inactivated the enzyme by blocking access to the active site. The small differences measured in rate of reduction and binding of oxalate fit well the linear relationship that describes the active site mutations (Fig. 3). The cysteine to alanine substitution probably causes a slightly altered conformation that results in a two fold less active enzyme. Again the decreased efficiency is reflected in the capacity to stabilize the transition state.

C287A has not yet been thoroughly characterized. In all the properties studied so far, however, it is even more like the wild type enzyme than C203A.

The spectral properties of R293K imply a unique protein-flavin interaction (Fig. 4). R293K shows a peak at 340 nm with a shoulder at 360 nm, and the ratio Azso to A456 is 18.0 in 10 mM ImCI, pH 7.0. The extinction coefficient of 6700 M-1 cm-1 is unusually low, as determined by denaturation of the enzyme with SDS. This indicates that, as isolated, the enzyme may contain part of the flavin in a derivatized form. Covalent adducts of the N(5) position in lactate monooxygenase exhibit an absorbance peak typically around 350 nm (23), with an extinction coefficient in the order of 7000 M-1 cm-1. Treatment with methyl methanethiolsulfonate (MMTS) led to spectral changes suggesting that a protein thiol may be involved. Upon reacting the mutant enzyme with MMTS, the absorbance at 456 nm increased 1.4 fold (Fig. 5), which was correlated with an activity increase of the same magnitude. The spectrum at the end of the reaction was similar to that of underivatized flavin with a peak at 370 nm. Moreover, the ratio of A280 to A456 was decreased to 9.1, indicating that the equilibrium had been displaced by MMTS reacting with the free cysteine thiol. Thus, the model assumes a covalent bond between a cysteine residue and the flavin N(5) position. The bound flavin which causes the peak at 340 nm is postulated to be in equilibrium with underivatized flavin on the enzyme. Freshly purified enzyme contains 20 - 30 % of the covalent adduct, which

208 does not absorb in the 450 nm region. Upon treatment with SDS, the flavin is released and the adduct immediately displaced.

H,0,

?

o

N-^N^O

Reduction

NH

/ Protein

O

/

H

Protein - S ^ max" 340 nm

»C

? O N-^N O O

NH

Protein - S X

m a x = 456 nm

Protein -S~ + C H 3 - S - S - C H 3

-

P r o t e i n - S - S - C H 3 + CH3S02"

O MMTS

Fig. 4. Postulated Formation of Covalent Adduct with R293K.

It appears that the equilibrium is also displaced upon reduction of the enzyme. When oxygen is admitted to reduced enzyme, reoxidation occurs in two phases. The first phase leads to an increase in absorbance at 456 nm which is about 1.4 times that of the initial enzyme. The second phase then returns the enzyme to the starting absorbance and is attributed to the re-establishment of the original equilibrium. Importantly, the spectrum of the initial reoxidation intermediate does not show the peak at 340 nm, but resembles the MMTS-treated enzyme.

Turnover also increases the absorbance of the enzyme, which seems to conflict with return of the adduct after reoxidation of the enzyme. However, the products of catalysis by R293K are 60-80 % H2O2 and pyruvate (Table 1), and it was shown that treatment with H2O2 will also lead to an increase in absorbance at 456 nm. Again the equilibrium is displaced, presumably irreversibly, due to oxidation of the postulated protein thiol. Further evidence needs to be collected; however, the data obtained so far are consistent with the model proposed.

209

c

0.1

re .o

o tn

-O


A(OH) # CoA + NAD + + H 2 0

(I)

A(OH) # CoA + NADH + H + —> A(OH)H 2 CoA + NAD +

(II)

where ACoA is the physiological substrate 2-aminobenzoyl-CoA, A(OH)#CoA is a nonaromatic intermediate and A(OH)H 2 CoA is 2-amino-5-oxocyclohex-l-enecarboxyl-CoA. Reaction (I) is a typical monooxygenation while (II) is a hydrogenation (reduction). Recendy we have shown that ACMR binds 0.5 equivalents ACoA per enzyme flavin and that the kinetics of flavin reduction with NADH and flavin reoxidation with either oxygen or Nethylmaleimide (NEM) are strongly biphasic [3]. NEM serves as an artificial electron acceptor [1] for ACMR and is considered to be a substrate analog for the hydrogenation reaction (equation II). In this report we discuss the relationship between these half-site reactivities and the two flavin cofactors involved in catalysis.

Results Resolution of purified ACMR by chromatography on Mono Q Purified ACMR can be resolved preparatively into three major (1, 2, 3) and two minor species (la, 2a) by ionic exchange chromatography on Mono Q (Fig. 1). The three main species differ markedly in their specific activities and in their ratios of absorption A280/450, which reflects the proportion of holo- to apo-protein, i.e. the FAD content. This ratio increases from species 1 to 3 suggesting a decrease of FAD content. The relative FAD contents are approximately (in %): Untreated ACMR = 100; fraction 1 - 78; fraction 2 = 72; fraction 3 - 52. Most importantly, the distribution of products depends on whether ACoA is turned over by ACMR

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

260 fractions 1, 2 or 3. Upon separation from NAD + /NADH and enzyme, product analysis was carried out spectrophotometrically as shown in Fig. 2. From the results it is apparent that turnover of ACoA using untreated ACMR yields predominantly A(OH)H2CoA (IV), while with fraction 3 the content of A(OH)CoA (III) is = 60%. This suggest that all three species 1, 2, and 3 are fully competent in monooxygenation, but vary in their capacity to hydrogenate. The relative differences between these activity profiles can readily be explained by selective loss of FAD at the active site which catalyzes hydrogenation. — < — 1 — >

1

2

0.3

1a o

s0-2 s c CO -Q

1

ri \

0.75

0.5

3 /

O

eo

2 c m TJ CO

20

II

40 Time (min)

.max = 365 nm

Scheme 3. Mode of interactions of the substrate or analog thioester carbonyl group with the protein. In this specific case of p-aminobenzoyl-CoA the free electron pair of the p-amino group is delocalized towards the thioester carbonyl group as a consequence of the two hydrogen bridges formed by the ribityl-2'OH and by the Glu376 backbone N-H. The interactions which bring about these effects have been identified and consist of two hydrogen bridges pointing towards the thioester carbonyl as shown in Schemes 2, 3 and 4. One involves the riboflavin 2'-OH function. This interaction has been suggested from the three-dimensional structure [17,18]. In order to prove it we have synthesized 2-deoxy-FAD (Scheme 4) and used it to reconstitute pig kidney MCADH from apo enzyme. The artificial 2deoxy-FAD-MCADH binds octanoyl-CoA, albeit weaker as compared to wtMCADH. It is essentially devoid of activity, the flavin being reduced at a rate = 10^ times smaller than that of wtMCADH. The extent of reduction is also much smaller, =50% of the total flavin, compared to wtMCADH (> 95%) and indicating a modification of the redox potentials involved in the couples (Scheme 1,1).

CH 2 —O-P-AMP

H-COH

2'-deoxy-FAD

h-C OH l H C-H

I CH 2

3

2 — "deoxy"

, H-C

f

Y

Y

T

H

2

291 „„

G,u376

—

^ A l C


3 in 2 N nitric acid set at 21.60 ppm which is the chemical shift value for the ^ N H 4 resonance from liquid ammonia (2).

Results

The

13

C-NMR spectra of MCAD reconstituted with l 3 C-enriched FAD at 2-, 4-, 4a- and

lOa-positions are shown in Fig. 1. The signal for each labeled atom was observed as a broad but distinct peak. In Fig. 2 are shown the 15 N -NMR spectra of MCAD reconstituted with 15N- enr i c hed FAD at 1-, 3-, and 5-positions. The signals for the labeled atoms were observed as indicated in the figure. The 13^- and 15n_NMR spectra of MCAD in the reduced state reconstituted with 13^- and ^ N - F A D were likewise measured. The reduced form of the enzyme was obtained by anaerobic addition of small molar excess of sodium dithionite. The chemical shift values of the NMR signals of bound FAD of oxidized and reduced MCAD are summarized in Table 1 in comparison with those of free FMN in the oxidized and reduced forms in aqueous solutions and with those of tetraacetylriboflavin in a nonpolar medium. Comparison of these values for MCAD with those for the reference species revealed that a weak hydrogen bond exists at C(2)=0 and that strong hydrogen bonds are formed at N(l), N(3)-H, C(4)=0, and N(5). It is to be noted that the resonance positions of both 4a-

and 10a- 13 C are at lower fields than the corresponding carbons for any other

flavoprotein examined thus far (3), indicating, the unique environment around these positions of flavin in MCAD. From comparison of the chemical shift values for reduced MCAD with

295

Fig. 1 (Upper). !3C-NMR spectra of MCAD reconstituted with 13C-enriched FAD in the oxidized state. The arrow heads indicate the positions of the corresponding l^C-resonances. Fig. 2 (Lower). 15 N-NMR spectra of MCAD reconstituted with 15N-enriched FAD in the oxidized state. The arrow heads indicate the positions of the corresponding resonances.

296 those of reduced FMN in the neutral and anionic forms (Table 1), we conclude that the reduced flavin in reduced MCAD is in the anionic form. Among the chemical shift values for reduced MCAD, that of 4 a - ^ C is particularly downfield-shifted from that of anionic reduced FMN, implying that the electron density at this position is low. This is consistent with the low reactivity of reduced MCAD toward molecular oxygen. In the l^C- and 15N-NMR spectra of the complex of MCAD with a substrate analog, acetoacetyl-CoA, the 5- 15 N resonance was specifically upfield shifted from that of free oxidized MCAD, while other resonances were not appreciably shifted (Fig. 3 and Table 1). This suggests that this substrate analog, hence a substrate, induces a specific change in the electronic state at N(5) of FAD in the substrate (analog)-MCAD complex.

Discussion

Accumulated investigations on 13C- and 15 N -NMR of flavoproteins as well as on free flavins under various conditions have demonstrated that the isotopically enriched atoms of the flavin nucleus are excellent probes for the electronic states and hydrogen-bonding network associated with flavin bound to flavoproteins (4). The hydrogen bonding network is one of the most influential factors in fine-tuning the flavin reactivity, as demonstrated by molecular orbital calculations (5-7). Close inspection of the chemical shift values of N(l), C(2), N(3), C(4), and N(5) for oxidized MCAD with other forms of flavin (Table 1) suggests there exist strong hydrogen bonds at N(l), N(3), C(4)=0, and N(5) and a weak one at C(2)=0. It is noted that the chemical shift value of this carbon is the highest field among the flavoproteins with ^ C - N M R data available to date (3), indicating the hydrogen bond at this position is weak. In contrast, the chemical shift value for C(4) is at the lowest field among the flavoproteins examined to date by 1 ^C-NMR (3). This suggests that the hydrogen bond at C(4)=0 is particularly strong.

This hydrogen-bonding network

corresponds to a model where hydrogen bonds are formed at N(l), N(3)-H, C(4)=0, and N(5) but not at C(2)=0. The ab initio calculation of this hydrogen bonding network lowers the energy of the LUMO of the oxidized flavin from that of flavin without hydrogen bonds and increases the AO coefficient at N(5) in LUMO (7). This hydrogen bonding network, therefore, tunes the flavin reactivity in such a way that electron flow from the substrate to flavin at the N(5) position is facilitated.

297 Table 1.

13

C- and

15

N-Chemical Shift of Enriched FAD Bound to MCAD Positions C(2)

C(4)

C(4a)

C(10a)

N(l)

N(3)

N(5)

MCAD (oxidized)

159.5

166.8

141.1

155.5

183.6

161.1

334.7

MCAD (reduced)

160.3

160.6

106.1

155.7

175.1

151.4

63.9

a

MCAD-AAC >

160.1

167.0

140.3

159.4

325.2

161.4b)

165.3W

137.8b)

156.0 153.7b)

184.0

FMN

190.8°)

160.5°)

334.7°)

TARFd)

156.8

161.4

137.2

150.7

199.9

159.8

344.3

e

FMNH2 )

152.7

158.8

104.7

145.9

128.0

149.7

58.0

FMNH" f )

159.8

159.3

103.0

157.1

182.6

149.3

57.7

TARFH28'

152.2

158.6

106.8

138.7

116.7

145.8

60.4

a) The complex of MCAD with acetoacetyl-CoA. b) Values extrapolated to infinite dilution (16). c) Vervoort et al. (17). d) Teraacetylriboflavin in chloroform (17). e) Neutral form of reduced FMN (17). f) Anionic (at N-l) form of reduced FMN (17). g) Reduced teraacetylriboflavin in chloroform (17).

r

MCAD-AcAcCoA

ll

H

/

C

o ^

[1-

A




low

tion

/ C

^ C

/

II

III

assigned

4: Structure

the

band

to

of

but

[ 3 - ^ C ] - l a b e l i n g of C(3)=0

was s c a r c e l y

frequency,

Therefore,

the bond o r d e r

double

bond s t r e t c h i n g mode o f low f r e q u e n c y

in the enzyme-bound a c e t o a c e t y l - C o A . e f f e c t near the

labelings. Judging from

lowered from t h a t o f

the resonance s t r u c t u r e

c o u l d be an e l e c t r o s t a t i c

acetoacetyl-CoA.

a f f e c t e d with other

is

\ C O A

acetoacetyl-CoA

The 1478-cm - ^ band s h i f t e d t o a f a i r l y

C] -labeling,

band

c

I

the 1478-cm~^ band i s a s s o c i a t e d w i t h the C ( l ) - 0 m o i e t y .

nature.

3

H

acetoacetyl-CoA. 13

o

H

band s h i f t e d t o 1616 cm~^ upon we

C

o

H

Fig.

Thus,

A

o"

I

(Fig.

ligand,

e.g..,

Thus the

a double bond

4 ) has a l a r g e

T h i s s u p p o r t s the

with

contribu-

idea that

there

by placement o f

a

306 suitable positive charge or a positive end of a dipole adjacent to the enolate

oxygen (7).

As both bands of oxidized flavin and the enolate for« of

acetoacetyl-CoA were resonance-enhanced in the spectra excited at 632.8 nm, the complex is concluded to be oxidized

enzyme

and

the

the

enolate

charge-transfer for«

complex

between

of acetoacetyl-CoA.

the

The highest

frequency band assigned to a C(3)=0 stretching node of acetoacetyl-CoA

was

observed

and

at

different

positions for the three enzymes (SCAD,

MCAD,

butyryl-CoA dehydrogenase), as mentioned above. This may depend on the difference in the enzyme-ligand interaction and may be

related

to

the

sub-

strate specificity.

Purple complex When the enzyme reacts with a substrate,

acyl-CoA,

enzyme with the product enoyl-CoA is produced. called

the

oxidase.

non-labeled

spectrum is similar to octanoyl-CoA butyryl-CoA.

The intermediate complex is

purple complex after the famous purple complex of D-amino acid

The resonance Raman spectra

butyryl-CoA

the complex of reduced

and the

of

labeled spectrum

the SCAD 13

with of

purple

complex

C are shown in Fig.

the

MCAD

purple

The band was assigned to the

reduced flavin (6).

have been oxidized to crotonyl-CoA. 10 frequency with 1-,

2-,

with 1 - ^ C labeling,

or 3~

with

moiety

of

the substrate must

The 1577-cm - ^ band shifted

C labeling.

The

labeling of

C(10a)=C(4a)-C(4)=0

As the enzyme was already reduced,

5.

complex

The 1615-cm - ^ band was not affected upon ^ C

(6).

with

to

a

low

As the isotopic shift is large

the band is mainly associated with

a

C(1)=0

double

bond stretching mode of crotonyl-CoA. The band is at too low a frequency to be

a C=0 double bond stretching mode.

pure keto-form,

but

should

polarized form at the C(1)=0.

contain

Thus, an

the crotonyl-CoA is not in a

appreciable

contribution

hancements of both bands of reduced flavin and crotonyl-CoA gest

that

the

purple

the

a

strongly

sug-

complex is the charge-transfer complex between the

reduced enzyme and crotonyl-CoA. in

of

Whichever the case may be, the resonance en-

To compare the structure of

purple complex and free in solution,

crotonyl-CoA in buffer solution (Fig.

6).

crotonyl-CoA

we observed Raman spectra of

The 1584 cm"* band was also ob-

served in the spectrum of CoASH. Thus the band is probably derived from the adenine moiety of CoA, band varied in strength,

whose band appears around 1580 cm~*. but was scarcely affected in the

The 1655-cm - ^ band

frequency

307 with

these

isotopic

frequency with 1-,

substitutions.

2-,

or 3 -

13

The

1 6 2 7 - c a ~ * band s h i f t e d t o a low

C l a b e l i n g of crotony1-CoA.

Figure 7

• a r i z e s the i s o t o p i c f r e q u e n c y s h i f t s f o r o b s e r v e d Raaan bands o f CoA

in

the p u r p l e c o n p l e x and f r e e in s o l u t i o n .

s i a i l a r to each other, purple

conplex

sua-

crotonyl-

Both i s o t o p e e f f e c t s

r e c o n f i r m i n g t h a t the s u b s t r a t e butyryl-CoA in

has b e c o a e c r o t o n y l - C o A .

t o 1577 ca~* in t h e p u r p l e c o a p l e x of SCAD.

in

solution

This large frequency s h i f t

i n t e r a c t i o n between the C(1)=0 and t h e enzyae,

a d i r e c t p i e c e of e v i d e n c e f o r an a p p r e c i a b l e c o n t r i b u t i o n of f o r i of C(1)=0 a o i e t y

in the e n z y a e - b o u n d enoyl-CoA.

O

632.8 nm

the

The f r e q u e n c y of t h e C(1)=0 c a r -

bonyl a o i e t y of c r o t o n y l - C o A s h i f t e d f r o a 1627 c n - ^ when f r e e

dicates a substantial

are

Kia e t al_.

a

in-

and i s

polarized reported

A 88.0 nm

non-

1577 1615 A

Iqkgieru

1525

non-labeled

—

15 51

, 1519 A A

JUA

[1- 13 C]

/^V

1800

1700 1600 RAMAN SHIFT (cm" 1 )

1500

VVVV

1571

15 26

1616 A

.

AJ ^Av ^

[2- 1 3 Cl

,569 A

161A

\ 1525

Jvv 1577

16 16

,

1700

\

i 1600

V/NA'

1523

' 1500

[3-13a

[Aj3C]

RAMAN SHIFT (cm )

Fig. 5 ( l e f t ) : Resonance Raaan s p e c t r a e x c i t e d a t 6 3 2 . 8 na of t h e p u r p l e c o n p l e x of SCAD f o r a e d upon t h e a d d i t i o n of n o n - l a b e l e d or i s o t o p i c a l l y l a b e l e d b u t y r y l CoA. Fig. 6 ( u p p e r ) : Raaan s p e c t r a e x c i t e d a t 4 8 8 . 0 na of nonl a b e l e d and i s o t o p i c a l l y l a b e l e d crotonyl-CoA.

308

Free, Complex non-labeled

I.

)

1 -13C

Fig. 7: Summary of t h e i s o t o p i c frequency s h i f t s f o r observed Raman bands of crotonyl-CoA in the SCAD p u r p l e complex and f r e e in s o l u t i o n .

[

2-13C 3-13C 4-13C

that

, 1700

_j 80 o

5

a> N |

60 40

k_

o z

20

4

6

8

10 12 14 16 Substrate chain length

18

20

Figure 2. Chain length dependance of the activity of MCADH, MLCADH, LCADH (all human), and rat LCADH. The activity of rat LCADH was measured with the DCPIP [8], all others with the ferricenium assay [7], For specific activities see Table 1. The K m values for the C12 and C14 substrates are = 4|j.M and = 5 |J,M, respectively, and thus essentially the same as for MCADH (=4(iM for C12); values for LCADH are not yet available. Isovaleryl-CoA is not a substrate for MLCADH. Table 1. Comparison of chain length dependance of activities of "human MLCADH", human MCADH, human LCADH expressed in E.coli, and rat liver LCADH. Acyl-CoA Substrate

MLCADH Humana) (min-1)

MCADH Humana) (min-l)

LCADH Human3) (min-l)

C4-CoA C6-C0A C8-C0A ClO-CoA C12-CoA C14-CoA CI6-C0A CI8-C0A C20-CoA

0 3 6 12 203 158 34 3 3

500 851 1101 665 613 259 116 0 0

0 43 56 119 148 93 69 47 13

a) Ferricenium assay in 0.1M phosphate buffer pH 7.6 [7] b) Data from [8], PMS/DCPIP assay.

LCADH Rat b ) (U/mg) 0 0 0.7 1.5 1.8 2.1 2.0 1.6 0.9

330 Human MCADH and the corresponding enzymes from pig or beef have a characteristic pH dependence of their activity, which reflects pK a values between 7 and 8.5 depending on the substrate [1], The pH dependence of the activity of MLCADH determined using dodecanoylCoA shows a pK a = 8, which is clearly in the same range as that found with MCADH (pK ~ 8.2, [1]). This indicates that the factors at the enzyme active site, which influence basicity of functional groups, have not been altered significantly by the double mutation.

Conclusions The preliminary results presented in this report clearly demonstrate that the hypothesis according to which in LCADH and IVDH Glu261 and Glu254 respectively have the same catalytic role as Glu376 in MCADH is tenable. The similar velocity of turnover of MLCADH as compared with LCADH indicate that the orientation of Glu255 with respect to the substrate and the flavin cofactor must be very similar to that assumed to be present at the active center of LCADH [4], The finding of K m values comparable to those of the other MCADH indicates that substrate binding also must be very similar. The most interesting, and perhaps unexpected, finding is the chain length dependence of the activity, which is surprisingly narrow, but which is clearly closer to that of LCADH than that of human MCADH (Figure 2). Therefore we conclude that the position of the proton abstracting base affects the interaction of acyl-CoA dehydrogenases with substrates i.e., the chain length specificity. Crystallographic studies currently carried out in collaboration with Dr. J.J. Kim might help clarify this point.

Acknowledgements: This work was supported by grant Gh 2/4-7 from the Deutsche Forschungsgemeinschaft. References 1. 2. 3. 4. 5. 6. 7. 8.

Ghisla, S., S. Engst, P. Vock, V. Kieweg, P. Bross, A. Nandy, I. Rasched, A.W. Strauss. 1994. This volume Powell, P.J. and C. Thorpe. 1988. Biochemistry 27, 8022-8028 Bross, P., S. Engst, A.W. Strauss, D.P. Kelly, I. Rasched, S. Ghisla. 1990. J Biol Chem 265,7116-7119 Kim, J.-J.P. and J. Wu. 1988. Proc. Natl. Acad. Sci. USA 85, 6677-6681 Nandy, A., P. Bross, I. Rasched and S. Ghisla. In preparation Thorpe, C., R.G. Matthews and C.H. Williams. 1979. Biochemistry 18, 331-337. Lehman, T.C., D.E. Hale, A. Bhala and C. Thorpe. 1990. Anal. Biochem. 186, 280-284 Ikeda, Y., K. Okamura-Ikeda and K. Tanaka. 1985. J.Biol.Chem. 260, 1311-1325

Glutaryl-CoA dehydrogenase from anaerobic, benzoate degrading Pseudomonas sp.: An FAD-dependent glutaconyl-CoA decarboxylase

Ulrich Härtel and Wolfgang Buckel Laboratorium für Mikrobiologie, Fachbereich Biologie, Philipps-Universität, D-35032 Marburg, Germany

Introduction It is well established that many bacteria are able to degrade aromatic compounds in the presence of molecular oxygen. This reactive molecule not only serves as terminal electron acceptor but also is required for the cleavage of the aromatic 6jt-system. In recent years it has been shown, however, that several bacteria are capable of degrading aromatic compounds in the complete absence of oxygen provided that suitable electron acceptors such as nitrate or sulfate are present. The anaerobic pathways are different from those found in aerobic organisms. A central intermediate of the anaerobic aromatic metabolism is benzoyl-CoA which can be formed from such diverse compounds as toluene, phenol, 4-hydroxybenzoate, 2-aminobenzoate, phenylacetate or benzoate (1). The 671-system of benzoyl-CoA then is cleaved by reduction rather than by oxgenation to cyclohex-l,5-diene-l-carboxyl-CoA. Hydration of both double bonds followed by oxidation and hydrolytic ring cleavage yields 3hydroxypimelyl-CoA which probably undergoes P-oxidation to glutaryl-CoA (2). This paper describes the purification and properties of glutaryl-CoA dehydrogenase from Pseudomonas sp. grown anaerobically on benzoate and nitrate. The enzyme also catalyses the decarboxylation of glutaconyl-CoA to crotonyl-CoA at a somewhat higher rate. Although the latter activity does not involve a redox reaction, it is strictly dependent on oxidized FAD present in the enzyme (3).

Results Glutaryl-CoA dehydrogenase catalysing the oxidative decarboxylation of glutaryl-CoA to crotonyl-CoA was induced in all organisms when grown anaerobically on aromatic compounds. Furthermore, glutaconyl-CoA decarboxylase activity was likewise increased in

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

332 COO"

o—d

COSCoA

COSCoA

COSCoA

H2O ,

2H

3 steps Fig. 1. Anaerobic degradation of benzoate COSCoA "OOC'^^^COSCoA Glutaryl-CoA

(i-oxidation

_

ooc

KJ

OH

FAD FADH 2 " O O C ^ ^ ^ C OS C o A Glutaconyl-CoA

H+

C02 H3Cv

FAD

-4, the enzyme was slowly reduced to a colorless protein and readily reoxidized on aeration. These spectral characteristics were very similar to those of N.winogradskyi enzyme(2). The prosthetic group of R.palustris enzyme was determined to be FAD with a thin layer chromatography of the extract of the enzyme treated by 10% trichloroacetic acid. As shown in Fig.2„ N.winogradskyi enzyme is composed of only one kind of subunit with the molecular weight of 36000, while the native enzyme exists as dimer of identical 36kDa polypeptides (2). On the other hand, R.palustris enzyme showed two migrating bands with the molecular weight of 38000 and 76000, respectively, on SDSpolyacrylamide gel electrophoresis.

This suggests that R.palustris enzyme is composed

of two kinds of subunits. However the antibody against N.winogradskyi enzyme crossreacted with both subunits(data not shown), indicating that the large subunit may be a dimeric form of the small subunit.

457

Fig. 1 Absorption spectra of R.palustris NADF"" reductase

Comparison

of enzymatic

properties

M

(I)

fet

. ",.

(2)

M

¡Mm

Fig. 2 SDS-PAGE of the purified enzyme. Lane M;Marker proteins; Lane IJf.winogradskyi enzyme; lane 2 Jt.palustris enzyme. of R.palustris

NADP+ reductase

with

N.winogradskyi NADP+ reductase. The R.palustris flavoenzyme oxidized NADPH with various electron acceptors such as DCIP and Menadione. Kinetic parameters for NAD(P)H-DCIP reductase activity were summalized in Table 1.

It is of interest that the enzymes reduce NADP f with

benzylviologen radical: benzylviologen photoreduced in the presence of proflavin and EDTA was reoxidized by the enzyme in the presence of NADP f under anaerobic conditions.

Although Fd is considered to be the physiological electron donor for

R.palustris NADFrt"reductase(3), the soluble non-haem iron sulfur proteins as Fd have not been purified from N.winogradskyi.

It should be determined by further studies in future.

Discussion In the present study, we have purified Fd-NADP* reductase(FNR)-like flavoenzyme to an electrophoretically homogenous state from R.palustris and shown that the enzymatic properties and molecular features are very similar to those of NADP + reductase purified from the chemoautotrophic bacterium N.winogradskyi (2). Recently, Seewaldt et al.

458 Table 1. Comparison of enzymatic properties of R.palustris N.winogradskyi NADP + reductase. N. winogradskyi enzyme

NADP + reductase and

R.palustris enzyme

NADPH-DCIP reductase activity

Km* Vmax**

1.89 7.25

1.22 13.5

NADH-DCIP reductase activity

Km* Vmax**

9.7 0.77

23 0.052

NADPH-BV reductase activity

Vmax**

1.38

1.07

+

+

BVH-NADP+ reductase activity *Km(fiM) **Vmax(sec"1) have reported that N.winogradskyi

is included in the alpha group among the purple

bacteria and is phylogenetically close to R.palustris as judged from the 16S ribosomal RNA{4).

Further, Tanaka et al. have reported that the primary sequence of

N.winogradskyi

cytochrome c is homologous to that of R.viridis cytochrome C2(5). In

this study, we have found that FNR-like flavoenzyme exists in R.palustris

and

N.winogradskyi

and further, the R.palustris enzyme cross-reacts with the antibody to

N.winogradskyi

enzyme. These results suggest that the chemoautotrophic bacterium,

N.winogradskyi

may be evolved from a photosynthetic bacterium which is a common

ancestor of all bacteria in alpha group. As far as is known, the chemoautotrophic bacteria such as Nitrosomonas europaea and Thiobacillus novellus are included in the beta group and use the Calvin cycle for C02 fixation. enzyme and the N.winogradskyi

With the purifications of the R.palustris

enzyme and the development of bacterial FNR-like

flavoenzyme-specific antibodies, studies on the universal distributions of FNRs among chemoautotrophic bacteria and photosynthetic bacteria are now possible. Reference 1.Klerk.H.DE., R.G.Bartsch, and M.D.Kamen. 1965. Biochim.Biophys.Acta 97:275 2.Kurokawa,T., Y.Fukumori and T.Yamanaka. 1987. Arch.Microbiol. 148:95-99 3.Yamanaka,T. and M.D.Kamen. 1967, 131:317 4.Seewaldt E„ K.Schleifer, E.Bock and E.Stackebrandt. 1982. Acta Microbiol. 131:287 5.Tanaka,Y., Y.Fukumori and T.Yamanaka. 1982. Biochim.Biophys.Acta 707:14

NMR Investigation of NADPH-Adrenoferredoxin Reductase with NADP+ and Adrenoferredoxin

Shigetoshi Miura and Yoshiyuki Ichikawa Department of Biochemistry, Kagawa Medical School, Miki-cho, Kita-gun, Kagawa 761-07, Japan

Introduction NADPH-adrenoferredoxin oxidoreductase (adrenodoxin reductase) is an FAD-containing flavoprotein with the relative molecular mass of 54000, which locates on the inner membrane of adrenal mitochondria. It functions in the mitochondrial electron transport system supporting the cytochrome P-450 dependent steroidogenic hydroxylation reactions. Adrenodoxin reductase receives two reducing equivalents from NADPH at once, and then delivers one reducing equivalent to adrenoferredoxin (adrenodoxin), an iron-sulfur protein containing a 2Fe-2S cluster. The present study was conducted to elucidate the interaction of adrenodoxin reductase with adrenodoxin and NADP+ in dynamic and equilibrium states. Use of proton NMR spectroscopy for the small molecules interacting with a macromolecule provided some insight into their static and dynamic binding modes (1, 2, 3). We measured proton NMR spectra for adrenodoxin and NADP + in the presence of adrenodoxin reductase.

Results and Discussion 'H NMR spectra of the reductase during the titration of the reductase with NADP + indicates no signals arising from NADP + appeared in the spectra until more than the stoichiometric amount of NADP + was added, indicating that there is no free NADP+ under the condition with a substoichiometric amount of NADP + . This suggests that the dissociation constant (Kd) of NADP + is much lower than the concentration of the reductase employed in this NMR study. During the titration of the reductase with more than the stoichiometric amount of NADP1", the line width of the resonances due to NADP + are getting sharper as the concentration of NADP + increases. Under the slow-exchange condition, the line width depending on the mole ratio of NADP + to the reductase could be due to the life time broadening. As the Ki of NADP1" is much lower than the concentration employed in the present experimental conditions, the binding site for NADP +

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

460

A8

N2

i °

0.9

0

02

OA

0.6

03

1.0

"

cc

§ 0.9

0

02

DURATION TIME(Sec)

OA

0.6

oa

1.0

DURATION TIME(Sec)

Figure 1. Time dependent transferred NOEs are plotted as a function of irradiation time. Irradiation frequency was placed at A2'H (A), and at NI 'H (B). The sample contained 0.3 mM adrenodoxin reductase and 3 mM NADP + in 50 mM phosphate buffer with 0.2 M KC1 at pH* 7.6, 28° C. is fully occupied with NADP+. Thus, the life time of free NADP + is dominated by the off rate constant

(koff)

of NADP + from the reductase. The estimated off rates of NADP + from the

reductase are calculated from the line width of N2H proton for nicotinamide moiety and A8H proton for adenine moiety. As expected from the above equation,

koff

remains constant over

wide range of the mole ratio examined. The resonances from both adenosine and nicotine moieties of NADP+ indicate a similar off rate of about 15-20 s_1. Time dependent transferred NOE of NADP + in the presence of adrenodoxin reductase were measured by one dimensional NOE measurement (Figure 1). Only between the resonances showing the cross-peaks in NOESY spectrum, time dependent negative NOEs are observed (data not shown). Time dependent profiles for development of NOE exhibit a long lag phase of as long as 80 ms. Usually, a long lag phase was observed in such a system with a slow crossrelaxation rate. This was thought as an indication of the spin diffusion, which is no longer directly connected with the distance between the specific pair of protons (4). However, the observed negative NOEs are highly specific for the pairs of resonances that form cross-peaks in the NOESY spectrum. Recent simulation demonstrated the exchange lag phase for the system with an intermediate exchange rate comparable with the cross-relaxation rate (3).

This

+

implication is consistent with the off rate of NADP from the reductase. It is possible to estimate qualitatively the conformation of the bound NADP + from the NOESY spectrum. Within the adenine-ribose moiety of bound NADP + , the intense NOESY cross-peak between A2'H/A8H protons and the weaker signal between A1'H/A8H protons suggest the adenineribose glycosidic torsional angle to be an anti conformation. Likewise, the NOESY cross-peaks between N1'H/N2H and N2'H/N6H protons for the nicotinamide-ribose moiety confine the

461

Figure 2. NMR spectra of adrenodoxin reductase for the aromatic region during the titration with NADPH. The samples contained 0.25 mM of the reductase without (a), and with 1.1 eq (b), 2.2 eq (c), 12 eq (d) of NADPH in 50 mM phosphate buffer at p H * 7.6, 2 8 ° C. ( • ) indicates resonances due to NADPH, and those marked with (o) are from NADP+. nicotinamide-ribose glycosidic torsional angle to an anti conformation. 'H NMR spectra for the reductase during the anaerobic titration with NADPH are shown in Figure 2. No NMR signal due to free NADP(H) was observed until more than stoichiometric amount of NADPH was added. 'H NMR signals due to NADP+ showed up in the spectra after addition of more than stoichiometric amount of NADPH. During further addition of excess NADPH, resonances due to free NADP+ appeared in the spectrum before those of NADPH emerged. This indicates that reduced adrenodoxin reductase has higher affinity for NADPH than that for NADP+. Nonselective Ti values for the C2H protons of His-10 and His-62 residues in adrenodoxin, which were assigned previously (5), were measured in the system containing adrenodoxin reductase and adrenodoxin. The complex formation with the reductase affects the relaxation time of these resonances. Chemical modification of adrenodoxin with diethyl pyrocarbonate indicated that the side residues of his-10 and his-62 were not directly involved in the site for interaction with redox partners (6,7). Thus, the effects on the relaxation time could be due to the change in the rotational correlation time upon complex formation. In the presence of the reductase at the concentration of one sixth of adrenodoxin, Ti values of histidine residues in

462 adrenodoxin become shorter as the concentration of NaCl increases. Assuming the slowexchange condition at low ionic strength, the upper limit of the off rate is estimated to be less than 4 s 4 (Figure 3). The estimated off rate constant is comparable with the kcat in cytochrome c reductase activity at low ionic strength.

ko« = 15-20 s

1

His-10

His-62

Figure 3. Schematic representation of the ternary complex among adrenodoxin reductase, NADP+ and adrenodoxin at low ionic strength. Adrenodoxin reductase possesses distinct binding sites for both NADP + and adrenodoxin. koff of adrenodoxin is estimated to be less than 4 s_1. koff of NADP+ is determined to be about 15~20 s 1 . The isoalloxazine ring of FAD is located at the site close to both NADP + and adrenodoxin. Acknowledgments This work was supported by grant-in-aid for scientific research on priority areas (No. 04225224 and 05209222) from the Ministry of Education, Science and Culture of Japan.

References 1, Ferrin, L. J., and Mildvan, A. S. (1985) Biochemistry 24, 6904-6913. 2, Banerjee, A., Levy, H. R„ Levy, G. C., LiMuti, C„ Goldstein, B. M„ and Bell, J. E. (1987) Biochemistry 26, 8443-8450. 3, Kouda, D., Kawai, G., Yokoyama, S., Kawakami, M., Mizushima, S., and Miyazawa, T. (1987) Biochemistry 26, 6531-6538. 4, Clore, G. M„ and Gronenborn, A. M. (1982) J. Magn. Reson. 48, 402-417. 5, Miura, S., and Ichikawa, Y. (1991a) Eur. J. Biochem. 197, 747-757. 6, Miura, S„ and Ichikawa, Y. (1991b) J. Biol. Chem. 266, 6252-6257. 7, Miura, S„ Tomita, S., and Ichikawa, Y. (1991) J. Biol. Chem. 266, 19212-19216.

One electron reduction of adrenodoxin reductase as studied by pulse radiolysis

M. Miki, K. Kobayashi The Institute of Scientific and Industrial Research, Osaka University, Mihigaoka 8-1, Ibaraki, Osaka 567, Japan S. Miura, Y. Ichikawa The Department of Biochemistry, Kagawa Medical School, Miki-cho, Kita-gun, Kagawa 761-07, Japan

Introduction NADPH-adrenodoxin oxidoreductase [adrenodoxin reductase, AdR] is an FAD-containing flavoprotein which is an essential component of the electron transport system for the cytochrome P-450-dependent steroid hydroxylation. AdR receives two electrons from NADPH and then delivers one electron to adrenodoxin (Ad), an iron-sulfur protein containing a 2Fe-2S cluster. The molecular mechanism of electron transfer from NADPH to cytochrome P-450 in the adrenal hydroxylation system via AdR has been studied. For the electron transport to cytochrome P-450, shuttling of Ad between AdR and cytochrome P-450 (1) or ternary complex formation among them was proposed (2). In this process, however, the formation of the semiquinone state of the enzyme has not been clearly identified. Some of the advantages of the pulse radiolysis technique for determining the spectral and kinetic behavior of one-electron reduction products of flavoproteins have been demonstrated (3). The present paper describes the reduction of the flavin of AdR by hydrated electron (e^ - ) in the presence and absence of Ad.

Results The reduction of flavin in AdR by e aq " was investigated by pulse radiolysis. The decay of eaq" accompanied an absorption decrease at 460 nm and an absorption increase at 600 nm. The kinetic difference spectrum at 20 jus obtained after the pulse shows that e aq " reduces

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

464 directly the flavin of AdR to form the blue semiquinone of the enzyme, as shown in Reaction (1). H+ eaq-

+

E-FAD

E-FADH*

(1)

Subsequently the semiquinone form was found to decay in the time range of seconds. From the difference spectrum, it is concluded that the semiquinone reacts by dismutation to form the oxidized and the fully reduced forms of the enzyme, as shown in Reaction (2). 2 E-FADH'

•

FAD

+

FADH 2

(2)

The reduction of the enzyme with eaq~ was performed in the presence of equilimolar NADP+. Under the condition, nearly all of NADP4" is bound to the enzyme. The flavin of the NADF^-AdR complex was reduced effectively by eaq". In contrast to NADP4" free enzyme, the rate of the formation of the semiquinone is independent on the concentration of the enzyme (6.1 x 10 4 s 1 at pH 7.5). In this process, the formation of NADP' bound to the enzyme with an absorption maximum at 400 nm was observed transiently. Subsequently, the decay of N A D P accompanied the absorption decrease at 460 nm. From these results, it is concluded that eaq" reacts with bound NADP+ to the enzyme to form NADP" and that an electron flows from NADP" to the flavin of the enzyme by an intramolecular electron migration, as shown in Scheme I. The intramolecular electron transfer in step (i) is rate-determing step in Scheme I.

e "

+

E-FAD-NADP + O

H+ >~ E-FAD-NADP' I ^ J 00

E-FADH-NADP +

(I)

A similar result was obtained in the reaction of CO2" with the NADP^-AdR complex. The reaction of the enzyme with e^" was performed in the presence of 1 mM NADP+. Since [ N A D P f ] / [ A d R - N A D P + ] is sufficiently high under the condition, eaq~ reacts preferentially with free NADP1", not bound to the enzyme. Then N A D P thus formed reacts with the NADP1"-enzyme complex to form the semiquinone of the enzyme. The rate of the formation of the semiquinone is similar to that obtained in the presence of equilimolar NADP1" and is independent on the concentration of the enzyme. In contrast, in the presence of either 1 mM N A D + or 1-methylnicotinamide (NMA), the formation of the semiquinone obeys second order kinetics, indicating that NAD" or NMA radical directly reacts with the flavin of the enzyme. The difference between NADP' and NAD' cannot be

465 explained by the redox potential of the reductants. Therefore, it is concluded that N A D P transfers an electron to NADP4" bound to the enzyme initially, and then electron flows to the flavin by an intramolecular migration. Similar experiments were performed in the presence of Ad. The complex of Ad-AdR was little reduced by e aq ", although the redox sites of both proteins can be reduced by eaq". This indicates that two proteins are bound tightly each other and mask their reduction sites completely. In contrast, in the reaction of eaq~ with the ternary complex of Ad-AdRNADP4", eaq" reacts with NADP4" bound to the enzyme to form NADP'. Subsequently the NADP' reduces both flavin of AdR and iron-sulfur cluster of Ad by intramolecular electron transfer.

Discussion According to pulse radiolysis experiments, flavoproteins can be classified as either e aq " reducible or -unreducible. The former class contains NADH-cytochrome bs reductase (3), adrenodoxin reductase, and flavodoxin (4). The latter class contains glucose oxidase (5), D-amino acid oxidase (5), lipoamide dehydrogenase (6), and xanthine oxidase (7). The flavins of this class cannot be reduced by e^", and e aq " reacts with amino acid residues in these proteins. The former class is functionally similar to serve as an electron transport between flavoprotein and corresponding oxidoreductase. This difference can be explained by the assumption that the flavin moiety of the former class is accessible to solvent. The reaction with eaq" is assigned mainly to a direct reaction proceeding via the exposed flavin. This is supported by the evidence that the complexes of Ad-AdR, putidaredoxinputidaredoxin reductase, and ferredoxin-ferredoxin NADP4" reductase were little reduced, although the redox sites of the flavin and corresponding oxidoreductase could be reduced by e aq ". In fact, the edge of the dimethylbenzyl ring in both flavodoxin (8) and ferredoxin NADP 4 " reductase (9) is well exposed to the solvent, as has been suggested by X-ray analysis. An important finding in the present work is that eaq~ or CO2" reacts NADP4" bound to the enzyme directly, not the flavin. This suggests that the nicotinamide moiety of NADP4" bound to the enzyme is accessible to the solvent and masks the FAD completely. On the other hand, in the ternary complex of Ad-AdR-NADP4", the NADP bound to the enzyme transfers electron both flavin of AdR and the iron-sulfur cluster of Ad. This raises the possibility that the flavin, the nicotinamide of NADP4", and the iron-sulfur cluster in the

466 ternary complex are located closely each other, as shown in Fig. 1. Similar structure has1 been proposed by X-ray analysis of ferredoxin NADP+ reductase (9).

Fig. 1. Schematic presentation of reaction of eaq~ with Ad-AdR-NADP+ complex

References 1. Lambeth, J. D„ Seybert, D. W„ and Kamin, H. 1979. J. Biol Chem. 254, 7255-7264. 2. Kido, T. and Kimura, T. 1979. J. Biol. Chem.

254,11806-11815.

3. Kobayashi, K., Iyanagi, T., Ohara, H., and Hayashi, K. 1988. J. Biol. Chem. Fa, 7493.7499. 4. Faraggi, M. and Klapper, M. H. 1979. J. Biol. Chem. 254, 8139-8142. 5. Kobayashi, K., Hirota, K., Ohara, H., Hayashi, K., Miura, R., and Yamano, T. 1983. Biochemistry. 22, 2239-2243. 6. Elliot, A. J., Münk, P. L., Stevenson, K. J., and Armstrong, D. A. 1980. Biochemistry. 22. 4945-4950. 7. Anderson, R. F., Hille, R„ and Massey, V. 1986. J. Biol. Chem. 261, 15870-15876. 8. Burnett, R. M., Darling, G. D., Kendall, D. S., LeQuesne, M. E , Mayhew, S. G., Smith, W. W„ and Ludwig, M. L. 1974. J. Biol. Chem. 249, 4383-4392. 9. Karplus, P. A., Deniels, M. J., Herriott, J. R. 1991. Science. 251, 60-66.

STRUCTURE-FUNCTION STUDIES ON NADPH-CYTOCHROME P450 REDUCTASE USING UREA-PERTURBATION AND

19

F NMR SPECTROSCOPY

R. Narayanasami, P. M. Horowitz, and B. S. S. Masters Department of Biochemistry, University of Texas Health Science Center at San Antonio, San Antonio, TX 78284-7760, U.S.A. J. D. Otvos Department of Biochemistry, North Carolina State University, Raleigh, NC 27695, U.S.A.

Introduction

NADPH-cytochrome P450 reductase is an ubiquitous protein belonging to a rare group of enzymes containing both FMN and FAD (1,2). It is a participant in the microsomal electron transport system, wherein reducing equivalents from NADPH are utilized in the metabolism of a variety of endogenous and exogenous substrates (3). The direction of electron flow is known to be: NADPH -» FAD -» FMN -» cytochrome P450 (4). Although NADPH-cytochrome P450 reductase has been under investigation for over three decades, our understanding of this enzyme at the structural level is very limited (5). Neither the exact mechanism nor the amino acid residues involved in the electron transport in this system is clearly understood. Our laboratories have been interested in exploring these problems by the application of techniques such as fluorescence, NMR, and urea-perturbation. Recently, we have utilized 31p NMR spectroscopy to further our understanding of the structural interactions in reductase (6,7). In the present studies, urea perturbation of reductase has been utilized to study the relationship between the loss of flavins and protein structural changes induced by urea by following the recovery of catalytic activity. Our preliminary results on the application of

NMR spectroscopy to reductase, in which all nine

tryptophan residues have been replaced with 5-fluorotryptophan, are also presented. This provides us with a powerful tool to explore the structure of reductase (8), with the trp residues acting as sensitive probes at or near crucial binding regions for flavins, NADPH, and various substrates.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

468 Results and Discussion

When 0 . 2 | i M reductase was incubated with various concentrations of urea for 4 hr, loss of cytochrome c reductase activity directly related to the concentration of urea was observed (Figure 1). When the same phenomenon was followed as a function of time, 95% of the original activity was lost within 2 0 min of incubation, even in 2 M urea (data not shown). When incubation mixtures were dialyzed in the presence of excess flavins, partial recovery of activity was obtained, suggesting a role for flavin loss in the urea-induced inactivation of reductase. Preliminary experiments revealed that the quenching of flavin fluorescence by bound reductase was abolished in the presence of urea in a time-dependent fashion and that the rate of fluorescence increase was a direct function of urea concentration.

FUrlatddad: to IscBUdoi / to the u**f iMa. Tim* " 72 hn.

1

2 [Urea] (M)

Fig. 1. Residual cytochrome c reductase activity of reductase samples containing urea.

0.00

0.S0

1.00 1.S0 [Urea] (M)

2.00

Fig. 2. Influence of added FMN on the activity of reductase samples containing urea.

Thus, the reconstitutability of enzyme samples containing urea was examined by addition of flavins to the samples, either during urea incubation or during the cytochrome c reductase assay. Figure 2 suggests that, in general, the largest proportion of original activity was attained when F M N was added both during urea incubation and to the assay mixture. In comparison, experiments in which the urea incubation samples were supplemented with F M N only during the assay exhibited less residual activity. Samples which were not supplemented with F M N displayed the smallest percentage of original activity. The flavin fluorescence enhancement, mentioned earlier, argues in favor of release of flavins in the presence of urea. Free flavins were isolated by ultrafiltration on a Centricon filtration

469 apparatus (Amicon, Inc.), and their concentrations were estimated by the Faeder-Siegel procedure (8). It can be seen in Figure 3 that concentration of free FMN increases with increasing concentrations of urea and reaches a plateau at 1 M urea, suggesting complete FMN removal. Concentration of free FAD is negligible at urea concentrations of 0-2 M and increases to a constant value at 4 M urea. This result is in agreement with the fact that FAD is much more tighdy bound than FMN and with the prior observation that addition of FAD to the urea-treated mixtures containing FMN does not increase the activity any further.

Fig. 3. Correlation of flavin release to changes Fig. 4. 1 9 F NMR spectrum of 5-fluorotryptophanin protein structure in the presence of urea labeled reductase (0.25 mM) at 470 MHz. It would be desirable to correlate all of these observations, pertaining to flavin binding and catalytic activity, to more direct structural changes. One approach is the determination of tryptophan/tyrosine fluorescence of reductase at different concentrations of urea to measure structural changes brought about by urea (Figure 3). Between 0-1 M urea there is a negligible change in the percentage of unfolded protein, the amount of which increases steeply in the higher urea concentration range (1-3 M), reaching a steady value at 4 M urea. All of these results are consistent with a model in which FMN is reversibly released from holo-reductase, a process accelerated by urea. The FMN-free enzyme subsequently loses FAD in a reversible manner. Once both flavins are lost, aggregation of the protein occurs, presumably due to exposed hydrophobic surfaces, by an irreversible process. The longer the duration of incubation with urea, the larger is the proportion of reductase lost to aggregation and misfolding and, thus, the less recoverable activity. This is consistent with the observation that the longer the time of incubation with a given concentration of urea, the smaller was the recoverable activity (data not shown). We are currently in the process of

470 studying the intermediates involved in this model by other biophysical techniques such as sedimentation velocity, light-scattering, and interaction with hydrophobic probes as monitored by fluorescence. Figure 4 is a

NMR spectrum acquired at 470 MHz of

reductase (0.25 mM) labeled with 5-fluorotryptophan, demonstrating the resolution of 8 of the 9 fluorotryptophan signals. Preliminary estimates of the spin-lattice relaxation time, T j , for all of the fluorine signals, are of the order of 600-800 msec. These values are higher than those anticipated for a molecule the size of reductase (79 kD) as result of aggregation (7). We are currently in the process of trying to obtain sufficient quantities of two of the tryptophan mutants of reductase to facilitate signal assignment in the ^ F NMR spectrum of fluorotryptophan-labeled reductase.

Acknowledgements This research was supported by NIH Grant No. HL30050 and Robert A. Welch Foundation Grant No. AQ-1192.

References 1. Iyanagi, T. and Mason, H. S. 1973. Biochemistry. 12, 2297-2308 2. Masters, B. S. S., Prough, R. A., and Kamin, H. 1975. Biochemistry. 14, 607-613 3. Masters, B. S. S. and Okita, R. T. 1980. Pharmacol. Ther. 9, 227-244 4. Vermilion, J.L., Ballou, D.P., Massey, V., and Coon, M.J. 1981. J. Biol. Chem. 256, 266-277 5. Porter, T. D. 1990. Trends. Biochem. Sci. 16, 154-158 6. Otvos, J. D„ Krum, D. P., and Masters, B. S. S. 1986. Biochemistry. 25, 7220-7228 7. Narayanasami, R., Otvos, J.D., Kasper, C.B., Shen, A., Rajagopalan, J., McCabe, T.J., Okita, J.R., Hanahan, D.J., and Masters, B.S.S. 1992. Biochemistry. 31, 4210-4218 8. Rule, G.S., Pratt, E.A., Simplaceanu, V., and Ho, C. 1987. Biochemistry. 26, 549-556 9. Faeder, E. J. and Siegel, L. M. 1973. Anal.. Biochem. 53, 332-336

Cerebellar Nitric Oxide Synthase Behaves as a Bi-Domain Structure

Bettie Sue Siler Masters, Essam Sheta, and Kirk McMillan Department of Biochemistry, The University of Texas Health Science Center, San Antonio, Texas 78284-7760 U.S.A.

Introduction

Cerebellar nitric oxide synthase (NOS) catalyzes the formation of NO and citrulline from L-arginine (1) and molecular oxygen (2). Two monooxygenation steps are required for product formation and N°-hydroxy-L-arginine (NOHArg) has been identified as an oxygenated intermediate in the reaction (3). The NOS isoform from rat brain is a 160 kDa enzyme containing heme (4-6), FAD and FMN (7), and tetrahydrobiopterin (8) prosthetic groups. The C-terminal 641 amino acid sequence of cerebellar NOS displays consensus regions for flavin and nucleotide binding and 58% sequence similarity to rat NADPHcytochrome P450 reductase (9).

This domain presumably serves as the oxidoreductase

domain. Carbon monoxide has also been reported to inhibit NO formation by -80% and to produce a reduced-CO difference spectrum with an absorbance maximum at about 445 nm (4). White and Marietta (10) also reported CO inhibition of both macrophage and cerebellar NOS activities. The apparent analogies between NOS and microsomal NADPHcytochrome P450 reductase- and cytochrome P450-mediated systems have led to the search for additional structural and mechanistic similarities. Cerebellar NOS has been purified from stably transfected human kidney 293 cells (1), generously provided by Dr. Solomon H. Snyder, using a modification of growth conditions and purification procedure (4). In the following studies, the effects of calmodulin addition on enzymatic activity and subsequent proteolytic cleavage by trypsin of NOS have led to the conclusion that the flavoprotein and heme domains interact in the production of NO and citrulline, dependent upon the binding of calmodulin (and its interaction with Ca+2) to a specific region between these domains.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

472 Results and Discussion

The effects of Ca +2 /calmodulin (Ca +2 /CaM) on the electron transport properties of cerebellar nitric oxide synthase, cytochrome c, 2,6-dichlorophenolindophenol, and ferricyanide reduction were measured in the presence and absence of Ca +2 /CaM. Only in the reduction of cytochrome c by NOS was any effect noted and, in this case, a >14-fold increase was obtained in the presence of Ca +2 /CaM (Figure 1A). 1 1 1 1 1 120 E

>3.

-

/

90 60

/

-

30 0

'

-CaM/

O

+CaM 0

* 1 I I " 20 40 60 80 100 Cyt c OiM) Figure 1A

Figure IB

Interestingly, the increased cytochrome c reduction was inhibited by increasing amounts of superoxide dismutase (SOD). The presence of either arginine or carbon monoxide inhibited by 38% or 20%, respectively, superoxide anion-mediated cytochrome c reduction (Figure IB), suggesting that NOS-bound heme participates in the formation of superoxide anion. In order to determine the effect of CaM on tryptophan fluorescence of NOS, incremental additions of CaM were made to a solution containing 500 nM NOS (Figure 2A). The spectra were obtained by excitation at 295 nm and emission at 340 nm and represented only the tryptophan residues of NOS, since CaM lacks tryptophan.

350 Wavelength

Figure 2A

400 (nm)

[Calmodulin] (nM)

Figure 2B

473 The Hill plot derived from the fluorescence data (Figure 2B) gave an n value of ~1 and a Kd of ~1 nM, with concentrations of CaM between 0.2-4 nM and 10 pM CaCl 2 .

Data

indicate that NOS is undergoing a conformational change with CaM addition to its oxidized form in the absence of any other added cofactors. The Kd for CaM determined in these experiments is in agreement with that of Bredt and Snyder (1) who reported an EC 50 of - 1 0 nM for activation of NOS by Ca +2 /CaM. The consensus binding sequence for calmodulin, located approximately in the middle of the NOS sequence, suggested that CaM could modulate the interaction of the flavoprotein domain (641-residue C-terminus) with the putative heme-binding domain (788-residue Nterminus, including the CaM-binding sequence). Therefore, limited proteolysis experiments were performed using immobilized trypsin at 4° C in the absence of Ca +2 /CaM. Incubation with trypsin produced an increase in cytochrome c reduction concomitant with a decrease in N O formation, measured by NO-hemoglobin capture (Figure 3A).

The increased

NADPH-cytochrome c reductase activity was not inhibitable by superoxide dismutase (data not shown), indicating that the flavoprotein domain, when separated from the N-terminal heme-binding domain, can directly reduce cytochrome c, as with the microsomal NADPHcytochrome P450 reductase.

S e p a r a t i o n of the N- a n d C - t e r m i n a l d o m a i n s of N O S y n t h a s e

Figure 3A

Figure 3B

474 Experiments in which NOS was digested with trypsin in the absence of CaM (Figure 3B) produced two major proteolytic fragments.

The production of these fragments was

prevented by the addition of CaM (data not shown) due to its binding to the region of NOS subject to cleavage by trypsin. Tryptic cleavage yields a - 8 9 kDa fragment, identified by sequence analysis as the N-terminal domain of NOS, and a ~79 kDa fragment. Neither fragment binds to a calmodulin affinity column due to tryptic digestion of the CaM binding site. Recent spectral studies (data not shown) indicate that the N-terminal fragment contains heme.

The C-terminal 641 residues constitute the flavin binding domain of NOS by

homology with other flavoproteins (1) and direct determination of FAD and FMN content (4, 11). Purification of these fragments to homogeneity will permit the determination of interactions between the domains in reconstituted systems. Acknowledgements Supported by NIH Grant No. HL 30050 awarded by the National Heart, Lung, and Blood Institute and by Grant No. AQ1192 from The Robert A. Welch Foundation to BSSM. References 1.

Bredt, D.S. and Snyder, S.H. 1990. Proc. Natl. Acad. Sci. USA. 87, 682-685.

2.

Kwon, N.S., Nathan, C.F., Gilker, C., Griffith, O.W., Mathews, D.E., and Stuehr, D.J. 1990. J. Biol. Chem. 265, 13442-13445.

3.

Stuehr, D.J., Kwon, N.S., Nathan, C.F., Griffith, O.W., Feldman, P.L., and Wiseman, J. 1991. J. Biol. Chem. 266, 6259-6263.

4.

McMillan, K„ Bredt, D.S., Hirsch, D.J., Snyder, S.H., Clark, J.E., Masters, B.S.S. 1992. Proc. Natl. Acad. Sci. USA. 89, 11141-11145.

5.

Stuehr, D.J. and Ikeda-Saito, M. 1992. J. Biol. Chem. 267, 20547-20550.

6.

Klatt, P., Schmidt, K„ and Mayer, B. 1992. Biochem. J. 288, 15-17.

7.

Stuehr, D.J., Cho, H.J., Kwon, N.S., Weise, M.F., and Nathan, C.F. 1991. Natl. Acad. Sci. USA. 88, 7773-7777.

8.

Mayer, B„ Mathias, J., Heinzel, B„ Werner, E.R., Wachter, H., Schultz, G„ and Bohme, E. 1991. FEBS Lett. 288, 187-191.

9.

Bredt, D.S., Hwang, P.M., Glatt, C.E., Lowenstein, C„ Reed, R.R., and Snyder, S.H. 1991. Nature. 351, 714-718.

10.

White, K.A. and Marietta, M.A. 1992. Biochemistry. 31, 6627-6631.

11.

Bredt, D.S., Ferris, C.D., and Snyder, S.H. 1992. J. Biol. Chem. 267, 10976-10981.

Proc.

STUDIES ON NAD(P)H-QUINONE OXIDOREDUCTASE Gabriella Tedeschi*. Shiuan ChenA and Vincent Massey* *From the Department of Biological Chemistry, The University of Michigan Medical School, Ann Arbor, Michigan 48109-0606 A Division of Immunology, Beckman Research Institute of the City of Hope, Duarte, California 91010

Introduction NAD(P)H-quinone oxidoreductase (EC 1.6.99.2) (DT-diaphorase) is an FAD-containing reductase that catalyzes the two-electron reduction of quinones to hydroquinones using either NADH or NADPH as the electron donor. The rat liver enzyme expressed in E. coli is a dimeric protein containing one non-covalently bound FAD per protein monomer of 30 kDa (1). In this study the flavin cofactor was removed and synthetic analogs of FAD were substituted in the flavin binding site as structural probes.

Results and discussion DT-diaphorase apoprotein was obtained by dialyzing the holoenzyme for 3 days at 4 °C against 200 mM potassium phosphate pH 6.0 containing 0.3 mM EDTA, 20 % glycerol, 2 M KBr and activated charcoal in the dialyzing fluid to adsorb FAD as soon as it passed through the dialysis membrane. The yield of apoenzyme in terms of protein recovery was about 60 - 70 % and the enzyme preparation regained 65 - 80 % of its specific activity with respect to the native holoenzyme after incubation in ice for 30 minutes with excess of FAD. The spectral properties of the FAD-reconstituted enzyme were virtually identical with those of the native protein. 8-Chloro-, 8-mercapto-, 6-mercapto-, 6-thiocyanato-, 6-azidoand 6-amino-flavins also bind to the apoenzyme with dissociation constants < 10'^M. However replacement of the flavin N(5) by carbon (5-deaza-FAD) results in a very large decrease in binding affinity (Kd 1.4 x 10"^ M at 25 °C). All these analogues are enzymatically active and can be reduced by NAD(P)H under anaerobic Conditions.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

476 Accessibility of the 8 position to solvent Reconstitution of DT-diaphorase with 8-chloro-FAD yields a non resolved spectrum closely similar to that of the native enzyme without formation of any intermediates. The result rules out the possibility of a covalent bound between the protein and the flavin, by displacement of the 8-Cl-group as found in lipoyl dehydrogenase (2). Accessibility of the flavin 8 position was explored by reacting the flavin with sulphur nucleophiles. As shown in Table 1 it is clear that the 8-C1 group of 8-Cl-FAD-DT-diaphorase reacts faster than does the 8-C1 group of the free flavin. This result suggests that the 8 position is accessible to solvent and that the protein can stabilize the tetrahedral intermediate of the reaction (3). In particular sodium sulphide at pH 5 and 8 also effects the displacement of chloride giving a spectrum typical of the paraquinoid form of 8-mercapto-FAD (4) (Fig.l) with an extinction coefficient of 30200 M~1 cm"' at 590 nm. The result is consistent with the fact that the native flavoprotein forms the anion radical upon photoreduction and suggests the presence of a positive charge near the N(1)C(2)=0 region of the flavin. As expected the decreased nucleophilicity of this form results in much lower reactivity of the enzyme-bound-8mercapto-FAD toward iodoacetamide and MMTS. Fig.l. Titration of 8-mercapto-FAD with apo-DT-diaphorase. 8-mercapto-FAD 3.4 (J.M (1) was titrated with apo-DT-diaphorase: 0.77 (J.M (2), 1.73 (J.M (3) and 3.4 |i.M (4) in 0.2 M KPi pH 8.0 containing 20% glycerol and 0.3 mM EDTA at 25 °C.. Fig.2. Spectral properties of 6-mercapto-FAD-DT-diaphorase in the absence (1) and in the presence (2) of 46 |iM dicumarol at pH 7.0. 0.12 1 i i i | i i i i | i i i i | i i i i | i i i i

o 12 0.1 0.08 « 0.06 I

0.04

0.02

0 388

488

588

688

wavelength (nm)

Fig.l

788

888

0 308

408

508

668

wavelength (nm)

Fig.2

788

888

477 Table 1. Second Order Rate Constants for the Reaction of 8-Cl-FAD-DT-diaphorase with Thiol Reagents.

8-C1-FAD

8-Cl-FAD-DT-diaph. 8-Cl-FAD-DT-diaph. + dicumarol 40 |iM (M" 1 min" 1 ) (M" 1 min"!) (M"l min"*) B-mercaptoethanol Thiophenol Thiophenol DTT

pH 8.0 pH6.0 pH7.0 pH 8.0

1.9 1.2 x l O 3 1.8 x 10 3 5.8

4.3 5.4 x l O 4 1.1 x 10 5 114.0

Second order rate constants of 1.07 M~1 min~l and 342

n.d. 1.08 x 10 3 n d 1.71

min~ 1 were calculated for the

reaction of 8-mercapto-FAD enzyme with iodoacetamide and MMTS respectively. These rates were not affected by the presence of dicumarol. However when the enzyme reconstituted with 8-C1-FAD was saturated with this competitive inhibitor the rates were dramatically decreased (Table

1). The oxidation of 8-mercapto-FAD by m-

chloroperbenzoic-acid or H2O2 occurs in two distinct steps with rate constants of 37.5 M"1 min"l and 3.5 M~1 min~l for the formation of S-oxide and sulfinate/sulfonate derivatives respectively. The reactivity of native enzyme and 8-mercapto-FAD with sulfite was also examined. No spectral perturbations were observed by reacting normal FAD-DTdiaphorase with 10 mM Na2SC>3 indicating no adduct formation between sulfite and the coenzyme. As expected the reaction with 8-mercapto-FAD enzyme yields 8-sulfonyl-FADenzyme and no further reaction occurs at N(5) position (5). Accessibility of the 6 position to solvent 6-Azido-, 6-amino-, 6-mercapto- and 6-thiocyanato-flavins were used as structural probes for the 6 position. Light irradiation of 6-azido-FAD enzyme yields the 6-amino derivative and it does not result in any covalent fixation of the flavin to the protein either in the presence or in the absence of dicumarol or NADP + . The reaction of 6-thiocyanato-FAD with dithiothreitol is fast (k=1750 M"1 min'l, the corresponding free flavin rate is 1000 M~1 min"l) suggesting that the 6 position is open to solvent. Also in this case the anionic form of 6-mercapto-flavin was highly favoured as shown in Fig. 2 by the presence of long wavelength absorbance.. Dicumarol binding has little effect on the rates of the reactions but

478 increases the pKa of the enzyme-bound flavin from below pH 5.0 to greater than pH 9.0 (Fig.2). 6-Mercapto-flavin reacts with iodoacetamide with a second order rate constant of 85 M"1 min'l (k=150 M~1 min'l for the free flavin). The reaction with MMTS is very fast

and the 6-mercapto-FAD spectrum can be regenerated

by the addition of

dithiothreitol. The 6-mercapto-FAD-enzyme reacts with mCPBA or H2O2 to produce the 6-S-oxide. As observed for Old Yellow Enzyme, this species is surprisingly resistant to further oxidation and extended incubation with H2O2 or excess peracid is required for sulfinate/sulfonate formation. Use of 5-deaza-FAD as mechanism probe Semiquinone formation with 5-deazaflavin is energetically highly unfavourable, thus precluding any role of 5-deaza-flavin radicals in one electron catalysis (6). It is thus a good probe for the differentiation between 1- and 2- electron processes. DT-diaphorase reconstituted with this FAD analogue is enzymatically active using 2-hydroxy-l,4naphthoquinone as electron acceptor in the standard assay. The specific activity

was

about 10 % of that of the native enzyme. In contrast no activity was observed with ferricyanide as one electron acceptor. In accordance with these data no radical intermediates were detected in rapid reaction studies using AZQ (diazaquinone) as electron acceptor. The results confirm that DT-diaphorase may be an obligatory two-electron transfer enzyme and may play a role in the detoxification of quinones and quinoid compounds by reducing them to the relatively stable hydroquinones. Acknowledgements This research was supported by a grant from the U.S. Public Health Service GM-11106. References 1. Chen, H„ Ma J.X., Forrest, G.L., Deng, P. S. K. Martino, P. A., Lee, T.D. and Chen, S., (1992), Biochem. J. 284, 855-860 2. Moore, E.G., Cardemil, E. and Massey, V., (1978), J. Biol. Chem. 253, 6413-6422 3. Schopfer, L.M., Massey, V. and Claiborne, A., (1981), J. Biol. Chem. 256, 7329- 7337 4. Massey, V., Ghisla, S. and Moore, E.G., (1979), J. Biol. Chem. 254, 9640-9650. 5. Fitzpatrick, P.F. and Massey, V., (1983), J. Biol. Chem. 258, 9700-9705. 6. Blankenhorn, G., (1976), Eur. J. Biochem. 67, 67-80.

EFFECT OF RIBOFLAVIN DEFICIENCY ON DT-DIAPHORASE

K. Yagi, S. Komura, M. Nakashima, N. Ishida, N. Ohishi Institute of Applied Biochemistry, Yagi Memorial Park, Mitake, Gifu 505-01, Japan L. Ernster Department of Biochemistry, Arrhenius Laboratories for Natural Sciences, Stockholm University, Stockholm, Sweden

Introduction DT-Diaphorase (EC 1.6.99.2) catalyzes the two-electron reduction of quinones to hydroquinones, thereby preventing O2" formation through redox cycling of semiquinones via one-electron quinone reductase (1). Therefore, this flavoprotein is generally regarded as an antioxidant enzyme. DT-Diaphorase is known to be induced by various compounds such as 3-methylcholanthrene (2), azo dyes, and r-butylhydroxyanisole (3). The administration of 3-methylcholanthrene to rats resulted in an increase in the mRNA level of this enzyme (4). These facts suggest an important physiological role of this enzyme in protecting the living body from toxic effects of xenobiotics. We previously reported that riboflavin deficiency brought about an increase in the lipid peroxide level in the body and considered that such increase is partly due to a decrease in the activity of another antioxidant enzyme, glutathione reductase (5). However, the possible change in DT-diaphorase activity and its implication in lipid peroxidation in riboflavin deficiency still remained to be clarified. The present study was undertaken to examine this problem.

Results and Discussion Effect of Riboflavin Deficiency on the Activity ofCytosolic and Mitochondrial

DT-Diaphorase

in Rat Liver Male weanling rats of the Wistar strain were divided into two groups and fed either a

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

480 Table 1. Effect of riboflavin deficiency on the activity of DT-diaphorase Group

Mitochondrial DT-diaphorase (mU/mg protein)

Cytosolic DT-diaphorase (mU/mg protein) +FAD

-FAD Riboflavin-deficient

166 ±

96*

Control

6 1 6 ± 198

-FAD

+FAD

197 ± 115*

70.8 ± 32.6

76.4 ± 31.8

555 ± 224

73.4 ± 34.1

88.4 ± 39.5

Mean ± SD is given, n = 9. Significantly different from the control: *pE.GSSG —»E.NADPH.GSSG pathway, a greater proportion of the reaction flux will proceed via the E —»E.NADPH —»E.NADPH.GSSG route and, as a result, the observed initial velocity will pass through a maximum and then decrease to a limiting plateau dictated by the rate constants of the kinetically less favourable route. The results of such an experiment are shown in Fig. 2.

504 The juxtaposing of like charges in a tightly packed area of the dimer interface of E.coli glutathione reductase has resulted in the acquistion of co-operative behaviour with respect to glutathione. A detailed steady-state kinetic characterisation suggests that this may come about as a result of the enzyme mechanism changing to become one in which a ternary complex is formed via two pathways. In order to investigate this possibility further, rapid reaction kinetic analysis of the enzymes is in progress. The structural basis of these observations is, as yet, undetermined and awaits crystallographic analysis to show how the introduced residues can be accommodated in such a densely packed region of the protein. Acknowledgments This work was supported by the Science and Engineering Research Council. A. Bashir was supported by a Science and Engineering Research Council Studentship and a bursary from Wolfson College, Cambridge. A. Berry and N. S. Scrutton are Royal Society 1983 University Research Fellows. References 1.

Williams, C.H. Jr., 1992. In: Chemistry and Biochemistry of Flavoproteins (F. Muller, ed.). CRC Press, Boca Raton, pp. 121-211

2.

Karplus, P.A. and G. E. Schulz. 1987. J. Mol. Biol. 195 , 701-729.

3.

Ermler, U. and G. E. Schulz. 1991. Proteins: Structure, Function, Genetics. 9 , 174-179.

4.

Greer, S. and R. N. Perham. 1986. Biochemistry. 25 , 2736-2742.

5.

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

6.

Scrutton, N.S., M. P. Deonarain., A. Berry and R. N. Perham. 1992. Science. 258, 1140-1143.

7.

Scrutton, N.S., M. P. Deonarain., A. Berry and R. N. Perham 1993. This Volume.

8.

Segel, I. H., 1975. Enzyme Kinetics. Wiley Interscience, New York. pp. 60-61.

ROLE OF CONSERVED GLYCINE RESIDUES IN THE NADPH BINDING MOTIF OF GLUTATHIONE REDUCTASE.

Maria Rescigno & Richard N. Perham Cambridge Centre for Molecular Recognition, Department of Biochemistry, University of Cambridge, Tennis Court Road, Cambridge, UK., CB2 1QW.

Introduction

In most nicotinamide

dinucleotide-binding

proteins, a common pa(J-fold has been identified in the domain primarily responsible for binding the coenzyme (1, 2; Fig.l). The positive dipole charge of the N-terminus of the a-helix can interact favourably with the negatively charged pyrophosphate moieties of the dinucleotides. This interaction is facilitated by the presence of a glycine residue at the beginning of the a-helix, the second conserved residue of the "fingerprint" motif, GXGXXG(A), (where X= any amino acid, (1, 3)), that has been identified in the (ia[3-fold. Inspection of the three-dimensional structure

Fig.l Schematic drawing of the ¡lap fold of E.coli glutathione reductase with bound NADPH (4, 9)

of Escherichia coli glutathione reductase (4) suggests that this residue (Gly-176) is so close to the pyrophosphate bridge of the bound coenzyme that the introduction of a larger side chain at this position would sterically hinder the interaction of nucleotide with protein. In this paper we study the effect of systematically changing the size and charge of the side chain at this site by generating eight point mutations at the codon for Gly-176 in the gene (gor) encoding E.coli glutathione reductase.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

506 Results and discussion

Site directed mutagenesis was carried out as described previously (5). The mutant genes were expressed and the proteins purified from an E.coli strain carrying a chromosomal deletion of the gor gene (5). Measurement of apparent kinetic parameters. The first mutation carried out (G176A) replaced glycine with alanine. Although this is considered a conservative mutation, the G176A mutant showed a very low catalytic efficiency and was essentially inactive (Table I). Its apparent Km for NADPH was 15 fold higher than that of the wild-type enzyme. Thus, a minimal change, the introduction of a methyl group, is sufficient to abolish activity presumably by disrupting the electrostatic interaction between the a-helix dipole and the pyrophosphate bridge of NADPH that helps correct orientation of the bound coenzyme.

Table I. Apparent Kinetic Parameters for Wild-Type and Mutant Proteins. ENZYME Wild-Type G176A G176S G176 V G176L G1761 G176E G176H G176R

tfmfor NADPH GxM) 19±2.7 334+51 38±6 >2500« 627 ±140 31+2.8 192±27 921 ±158 >2500°

(^m mut/ ^mWT)

¿cat (min -1 )

(% to WT)

kaitjKm (min-1, /¿M)

(1) (17.5) (2) (>130) (33) (1.63) (10) (48.5) (>130)

32000 18 4986 N.D.fr 603 49 74 229

(100) (0.06) (15.6) N.D.& (1.9) (0.15) (0.23) (0-71) N.D.&

684 0.05 131 N.D.fc 0.96 0.63 0.39 0.25 N.D. 6

Km for GSSG (pM) 97±9.1 485) (220) (485) (32.3) (5.9) (198) (485) (8.8)

a The apparent value of Km for NADPH could not be measured since at concentrations as high as 600 mM the enzyme remained unsaturated, b These kinetic parameters could not be determined because saturating levels of NADPH could not be reached. cThe apparent value of Km for GSSG could not be measured since discrimination in rate could not be achieved even at concentrations as low as 0.02 fjM. Glutathione reductase activity was measured at 30°C in 100 mM potassium phosphate buffer, pH 7.5. N.D., not determined. The serine mutant (G176S) is the most efficient enzyme, retaining 20% of the wild-type catalytic efficiency with an apparent K m for NADPH (38/iM) only slightly higher than that of the wild-type (19/iM). This is an interesting result since serine replaces the second glycine in some mercuric reductases. Although the crystal structure of these proteins is not known, it is

507 conceivable that a hydrogen bond is formed between the serine and the pyrophosphate, stabilizing the binding. Further increases in the size of the side chain at position 176 produced a marked decline in the catalytic efficiency of the mutant enzymes (Table I). Surprisingly, the apparent K m for NADPH of the isoleucine mutant, G176I, is very similar to that exhibited by G176S, whereas the ¿cat of the former is 1% of that of G176S. This suggests that even if the enzyme structure can rearrange to accommodate a larger side-chain, the nicotinamide moiety is incorrectly positioned for the electron transfer to the enzyme-bound flavin. In spite of the inherent electrostatic repulsion between the pyrophosphate and a negatively charged side chain of a residue at position 176, the G176E mutant exhibited a Km for NADPH only 10-fold higher than that of the wild-type enzyme. These results indicate a marked preference for a glycine residue at the N-terminal end of the a-helix. The absence of a side chain at this position is required to allow a close interaction between the pyrophosphate moiety and the a-helix dipole, enabling the correct orientation of the pyridine nucleotide in the binding pocket. All the above mutants showed lowered apparent Km values for GSSG. Site-directed mutagenesis studies carried out previously on a residue (Tyr-177) in the NADPH binding site of E.coli glutathione reductase (6) also revealed a lower Km value for GSSG. This result was unexpected since NADPH and GSSG bind in separate sites, and there was no evidence of major conformational changes. An explanation for this finding lies in the fact that in a kinetic mechanism such as that displayed by E.coli glutathione reductase, a mutation generated at one of the two separate sites which affects the ratio of the catalytic activities of the individual half-reactions, may cause a decrease in the Km value for the substrate of the half reaction that becomes less rate-limiting (7). Primary deuterium isotope effects. In order to determine whether the mutations had changed the rate-determining step in the overall reaction, primary deuterium kinetic isotope effects were measured for each enzyme (Table II). At saturating levels of GSSG, only a small D V value was observed when NADPH was used as the variable substrate for the wild-type enzyme. This is in accord with the value previously obtained for the same enzyme (8). For most of the mutants, the D V values were much higher (to a maximum of 8.5 in G176H). This suggests that reduction of the enzyme by NADPH has become rate-limiting in the overall reaction. Assuming that the mutations did not change the intrinsic isotope effect of E.coli glutathione reductase, then its value can be estimated to be >8.5. The DV/Xr values for most of the mutants and for wild-type enzyme were not significantly greater than 1.0.

508 Table II. Primary Deuterium Isotope Effects for Wild-Type and Mutant Proteins. ENZYME

D

V/K

Dy

1.15 1.45 1.035 2.31 1.657 1.947 N.D.a N.D.a 2.012 4.765 0.82 1.08 0.98 1.3 1.51 8.5 N.D.a N.D.a These values could not be determined (see Table I).

Wild-Type G176A G176S G176V G176L Gl 761 G176E G176H G176R a

Replacement of Gly-176 in E.coli glutathione reductase resulted in enzymes with very low catalytic efficiencies, indicating a great preference for a glycine residue at this site for the correct orientation of the coenzyme in the binding pocket. The increase of the isotope effects on V max shows that in the mutant proteins the reduction of the enzyme by NADPH has become a rate-limiting step of the overall reaction.

Acknowledgments MR was supported by the Ministero dell'Università', Università' di Milano, Italy.

References 1. 2. 3. 4. 5. 6. 7. 8. 9.

Wierenga, R.K., De Maeyer, M.C.H., & Hoi, W.G.I. 1985. Biochemistry. 24, 1346. Rossmann, M.G., Moras, D„ & Olsen, K.W. 1974. Nature. 25Q, 194. Scrutton, N.S., Berry, A., & Perham, R.N. 1990. Nature. 242, 38. Mittl, P.R.E., Berry, A., Scrutton, N.S., Perham, R.N. & Schulz, G.E.1993J. Mol.Biol. 221, 1921. Deonarain, M.P., Berry, A., Scrutton, N.S., & Perham. R.N. 1989. Biochemistry 2L9602 Berry, A., Scrutton, N.S., & Perham, R.N. 1989. Biochemistry. 2&, 1264. Matthews, R.G., 1990, Flavins and Flavoproteins (B. Curti, S. Ronchi & G. Zanetti, eds), Walter de Gruyter, Berlin-New York, 593 Vanoni, M.A., Wong, K.K., Ballou, D.P., & Blanchard, J.S. 1990. Biochemistry. 22, 5790 Kraulis, P.J., 19911. Appl. Crystallogr. 24, 946

Eukaryotic Lipoamide Dehydrogenase: Molecular Genetic and Structural Aspects

Kichiko Koike and Masahiko Koike Department of Pathological Biochemistry, Atomic Disease Institute, Nagasaki University School of Medicine, Sakamoto-1-chome, Nagasaki 852, Japan Akio Takenaka Department of Life Science, Faculty of Bioscience and Biotechnology, Tokyo Institute of Technology, Midori-ku, Yokohama 227, Japan

Introduction Lipoamide dehydrogenase (LD) (EC 1.8.1.4.) is the common flavoprotein component of three 2-oxo acid dehydrogenase multienzyme complexes, the pyruvate dehydrogenase complex (PDC), the 2-oxoglutarate dehydrogenase complex (OGDC) and the branched-chain 2-oxo acid dehydrogenase complex (BCODC)( 1,2). LD is also found in the glycine cleavage

O

, CoA-SH

R -C-S-Llp-SH ] I

(E1-p,o,b) Y ( E 2 - p , o , b ) [ U p ^ r ' S H [ FAD]

-s-s-

L i p ( E 3 )

í'-M J

R : CH 3 - (p), HOOCHßH ¿C - (o), CH 3CH - (b)

ch 3

NAD-* V ETC

Net Reaction :

O R -CCOOH + CoA-SH + NAD+ —

Yp-nadh+H*

O R

-C-S-CoA + COjJ + NADH + H+

Figure 1. Multistep reaction sequence in an oxidative decarboxylation of 2-oxo acids (p, Pyruvate; o, 2-oxoglutarate; b, branched-chain-2-oxo acid) by 2-oxo acid dehydrogenase complexes. E l , 2 - o x o acid d e h y d r o g e n a s e ; E2, a c y l t r a n s f e r a s e ; E3, lipoamide dehydrogenase (LD); TPP, thiamin pyrophosphate; Lip-S2 and Lip(SH)2, lipoic acid and its reduced form.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

510 system (3). LD catalyzes the transfer of two electrons to an electron transfer chain (ETC) via NAD+ from dihydrolipoic acid that is covalently bound to lysine residues of the central core, t h e c o m p l e x - s p e c i f i c d i h y d r o l i p o a m i d e a c y l t r a n s f e r a s e (E2), as shown in Figure 1. Eukaryotic LD cDNA clones have been isolated and characterized (4-6), however its gene has not yet been isolated. Human LD gene has been mapped to c h r o m o s o m e 7 (7) and the sequence of the 5' flanking region has recently been characterized (8). The three-dimensional structure of dimeric flavoprotein, human erythrocyte glutathione reductase similar to LD has been elucidated by an X-ray diffraction analysis (9). Yeast LD has been crystallized and structurally elucidated by X-ray method (10). Having amino acid sequence and three-dimensional structures would allow comparison with other flavoproteins to help determine the role of LDs in medicine. As a part of structurefunction studies of the 2-oxo acid dehydrogenase complexes, we separated LD component from the PDC and OGDC by gel-permeation high-performance liquid chromatography (HPLC) in a single operation. The amino acid sequence of the porcine LD was deduced from the nucleotide sequences of PCR products amplified from porcine heart cDNA. The crystallization and X-ray analysis of porcine LD have been attempted. The detailed structure of yeast LD are also discussed.

Results and Discussion

Simple-Rapid Isolation Procedure of Porcine Heart LD

The PDC and OGDC were purified from the extract of Keiline-Hartree particle preparation of porcine heart muscle, and LD, namely the common component of these complexes was isolated by gel-permeation HPLC in the presence of the protein dénaturants, in a single operation. Three component enzymes of the PDC were first separated by a gel-permeation HPLC on a TSK-Gel G4000SWG column containing 4M urea at 7°C within 1 h, as shown in Figure 2, followed by the ion-exchange chromatography (11). Porcine O G D C was also separated into three component enzymes by the similar procedure in the presence of 0.7 M guanidine hydrochloride, 0.05% Triton X-100 and 2 mM dithiothreitol at 10°C (12). The TSK-Gel column permitted very rapid simultaneous dissociation and fractionation of three component enzymes accompanied by good preservation of their activities and high overall yield of each component.

511

Retention

Time(min)

Figure 2. Elution profile of three component enzymes of porcine PDC by gel-permeation H P L C . P D H , Pyruvate dehydrogenase ( E l - p ) ; LAT, dihydrolipoamide acetyltransferase (E2-p);Fp,LD(E3). 63

168

1590

2323 b

(A) 0 ATG

Kb ,N

(B)

-29 40 iNh

560 H i e

521b 2N

(C)

1066

566b

H

2C

974

3N H 1N 2N

1027b 5(j1

1683

710b

•H

3C

1066 2c 1863 1183b

• H 3C

Figure 3. Amplification of the full length porcine LD cDNA by P C R . (A), scale (kilobase pair, kb). (B),structure of LD c D N A ; A T G , ATG codon; N and C, amino- and carboxyltermini. (C), location of forward ( 1 N ~ 3 N ) and reverse ( l C ~ 3 C ) p r i m e r s , nucleotide number and sizes (base pair, b) of amplified specific segments on cDNA.

512 PCR Amplification of Porcine cDNA Porcine mRNA was isolated by guanidine isothiocyanate preparation method followed by oligo(dT)-cellulose chromatography (13) from heart tissues obtained from newborn pigs. The forward and reverse oligonucleotide primers essential for amplification of full length of LD cDNA (beginning at ATG codon and ending at stop codon) were synthesized. The forward primer (29-mer) contained an additional 12 nucleotides coding for an artificial £coRI site. The reverse primer (29-mer) contained an additional 12 nucleotides coding for an artificial BamHl site. The location of primers for amplification and size of specific segment of LD cDNA are given in Figure 3. PCR products were subcloned into pUC19 vector and sequenced by the dideoxynucleotide chain-termination method with T7 DNA polymerase (13). As shown in Figure 4, the amino acid sequence deduced from the nucleotide sequence of PCR products was identical with that from the cloned cDNA for porcine LD (4) except two amino acid replacements.

These two porcine LDs exhibit a strikingly high sequence identity of 96%

with human LD (5). 10 20 30 40 50 60 70 80 90 P I :MQSWSRVYCTLAXRGHFNRIAHGLQGVSAVPLRTYADQPIDADVTVIGSGPGGYVAAIKAAQLGFKTVCIEKNETLGGTCLNVGCIPSKA P2 : L H : S S L 100 110 120 130 140 150 160 170 180 PI :LLNNSHYYHMAHGKDFASRGIEMSEVRLNLEKMMEQKSNAVKALTGGIAHLFKQNXVVRVNGYGKITGKNQVTATKADGSTEVINTKNIL P2 : H : D T H R G-Q—D 190 200 210 220 230 240 250 260 270 PI :IATGSEVTPFPGITIDEDTWSSTGALSLKKVPEKMWIGAGVIGVELGSVWQRLGADVTAVELLGHVGGIGIDMEVSKNFQRILQKQGF P2 : F H : 1 F V 1 280 290 300 310 320 330 340 350 360 PI :KFKLNTKVIGATKKSDGNIDVSIEAASGGKEVITCDAVLLVCIGRRPFTQNLGLEELGIELDPRGRIPVNTRFQTKIPNIYAIGDWAGP P2 : H : T K K 370 380 390 400 410 420 430 440 450 P I :MLAHKAEDEGIICVEGMAGGAVHIDYNCVPSVIYTHPEVAWVCTSEEgLKEEGIEYKVGKFPFAAHSRAKTNADTDGMVKILGQKSTDRV P2 : I : 460 470 480 490 500 510 PI:LGAHIIGPGAGEMINEAALAXjEYGASCEDIARVCHAHPTLSEAFREANLAASFGFAINF P2: H : L V S

Figure 4. Comparison of amino acid sequences of porcine (P) and human (H) LD deduced from the nucleotide sequences of their cDNAs. P I , sequence reported (4); P2, sequence deduced from PCR products; H, sequence reported (5).

Crystallographic Studies of Lipoamide Dehydrogenases The 2-oxo acid dehydrogenase complexes are the highly organized multienzyme systems, consisting of multiple copies of three component enzymes, 2-oxo acid dehydrogenase (El-

513 p,o,b), dihydrolipoamide acyltransferase (E2-p,o,b), and LD (E3), which catalyses the irreversible serial reactions specifically and efficiently, as shown in Figure 1 (1,14). The complexes are classified into two types of the central core structures composed of E2s with different symmetries depending on organisms. One has the 432 symmetry (15) in Gram negative bacteria and the other the 532 symmetry (16) in Gram positive bacteria and in eukaryotes. The crystal structure of the isolated component of LD from yeast was determined for the latter type (10). As suggested from sequence homology (17), the tertiary structures of LDs has been confirmed to be similar to that of glutathione reductase, forming a family of flavoproteins. As for the former type, the X-ray structures of LDs from Azotobactor vinelandii (18) and Pseudomonas fluorescens (19) have been reported, and the central core structure composed of E2s is discussed by Hoi and his collaborators (20). Thus it is revealed that the tertiary structures of LDs are essentially the same even in the different architecture between the two types. It is noticed that such a strong conservation of the structure may be the result of the functional restraint imposed by the enzyme. In the complexes, however, the LDs are assembled into the different core structures. The structural differences between the two types and their effects on the reaction are important knowledge for understanding the organization of this multienzyme system, and for modifying it or for designing new systems. For comparison of the details, it is necessary to know the higher resolution structure of LD from yeast. Recently it was found that the crystals of yeast LD newly obtained by a desalting method have the same space group as, but are slightly different in the cell parameters reported previously (10). They gave X-ray diffraction patterns more than 3.0 A resolution. The crystal structure, solved by the molecular replacement method shows the differrent molecular packing. Refinements of the atomic parameters are in progress . The details of the structure will be discussed in elsewhere. Preliminary data for yeast LD model construction with computer graphics by fitting into electron density maps are shown in Figure 5, A (helical part) and B (p strand). The flavin, adenine, ribose and diphosphate moieties are easily located on the omit map of FAD (Figure 5, C). On the other hand, the architecture of the OGDC has much in common with the PDC. The LD isolated from porcine heart OGDC (12) was crystallized by the hanging drop vapor diffusion method with ammonium sulfate as precipitant. The crystals are hexagonal plate in shape, as shown in Figure 6. They diffract more than 4.5 A resolution. The unit cell has a size of a=b=380 A, c=139 A and y =120° with hexagonal space group, P6 n or P6 n 22 (n=0~5). As the asymmetric unit contains several molecules, another crystalline form suitable for X-ray analysis is being surveyed.

514

515

Figure 5. Electron density maps of helical part (A) and several p strand ( B ) of yeast LD. On the omit map of F A D (C), the flavin, adenine, ribose and diphosphate moieties are easily located.

Figure 6. Crystal of porcine LD, grown using the sitting-drop vapor diffusion method.

516 Acknowledgments

We are grateful to Dr. Howard J. Jacob for helpful comments.This research was supported by grants from the Ministry of Education , Science, and Culture of Japan and the Vitamin B Research Committee.

References 1. Koike, M., Koike, K. 1976. Adv. Biophys. 9: 182-227 2.

Yeaman, S.J. 1989. Biochem. J. 257: 625-632

3. Kikuchi, G., Hiraga, K., Yoshida, T. Biochem. Soc. Trans. 8: 504-506 4.

Otulakowski, G. Robinson, B.H. 1987. J. Biol. Chem.

232:17313-17318

5.

Pons, G., Raefsky-Eatrin, C., Carothers, D.J., Pepin, R.A., Javed, A.A., Jasse, B. W., Ganapathi, M.K., Samols, D., Patel, M.S. 1988. Proc. Natl. Acad. Sci. USA 85: 1422-1426

6.

Browning, K.S., Uhlinger, D.J., Reed, L.J. 1988. Proc. Natl. Acad. Sci. USA 85: 1831-1834

7.

Otulakowski, G., Robinson, B.H., Willard, H.F. 1988.Somat. Cell Mol. Genet. 14: 411-414

8. Johanning, G.L., Morris, J.I., Madhusudhan, K.T.,Samols, D., Patel, M.S. 1992. Proc. Natl. Acad. Sci. USA 89: 10964-10968 9. Thieme, R„ Pai, E.F., Schirmer, R.H., Schulz, G.E.1981. J. Mol. Biol. 152: 763-782 10. Takenaka, A., Kizawa, K., Hata, T., Sato, S., Misaka, E.-T., Tamura, C., Sasada, Y. 1988. J. Biochem. 103: 463-469 11. Urata, Y., Koike, K„ Goto, S., Koike, M. 1991. J. Nutr.Sci.Vitaminol. 37: 257-267 12. Moriyasu, M., Koike, K„ Urata, Y., Tsuji, A., Koike, M. 1986. J. Nutr. Sci.Vitaminol. 32: 33-44 13. Sambrock, J., Fritsch, E.F., Maniatis, T. 1989. Molecular Cloning : A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York 14. Reed, L.J. 1974. Acc. Chem. Res. 7: 40-46 15. Oliver, R.M., Reed, L.J. 1982. In : Microscopy of Proteins, Vol. 2 (Harris, J.R., ed.). Academic Press, London, pp. 1-47

517 16. Stoops, J.K., Baker, T.S., Schroeter, J.P., Kolodziej,S.J., Niu, X.-D., Reed, L.J. 1992, J. Biol. Chem. 267: 24769-24775 17. Rice, D.W., Schulz, G.E., Guest, J.R. 1984. J. Mol.Biol. 174: 483-496 18. Mattevi, A., Obmolova, Kalk, K.H., Westphal, A.H., Kok, A., Hol, W,G.J. 1993. J. Mol. Biol. 230: G„ 1183-1199 19. Mattevi, A., Obmolova, G., Kalk, K.H., Berkel, W.J.H., Hol, W.G.J. 1993. J. Mol. Biol. 230: 1200-1215 20. Mattevi, A., Obmolova, G., Kalk, K.H., Teplyakov, A., Hol, W.G.J. 1993. Biochemistry 32: 3887-3901

Disruption of the Gene

Coding

for

Dihydrolipoamide

Dehydrogenase

in

Haloferax volcanii by Homologous Recombination.

N.N. Vettakkorumakankav and KJ. Stevenson Department of Biological Sciences, University of Calgary, Calgary T2N 1N4, Alberta, Canada L. C. Schalkwyk and W. F. Doolittle Department of Biochemistry, Dalhousie University, Halifax B3H 4H7, Nova Scotia, Canada

Introduction Dihydrolipoamide dehydrogenase (DHLipDH, EC 1.8.1.4) is an essential component of the 2oxoacid dehydrogenase (1) and glycine cleavage multienzyme complexes (2) of many eubacteria and eukaiyotes where it catalyzes the reaction: Dihydrolipoamide + NAD* = Lipoamide + NADH + H+

(Ref.3)

In Archaea, however, 2-oxoacids are metabolised by oxidoreductase systems (4) thus the discovery of DHLipDH activity in halobacteria was unexpected (5) and led to the purification and characterization of DHLipDH from Halobacterium halobium (6) and Haloferax volcanii (7,8). A physical map, transformation procedures and shuttle vectors are available (9-12) for analysis of the gene structure and function in Hf. volcanii. Here we report the localizing of the DHLipDH on the Hf. volcanii gene and the disruption of that gene allowing to us to determine whether DHLipDH activity is essential for growth under standard laboratory conditions.

Results The ipd gene (DHLipDH activity) was localized by dot hybridization to cosmid 126, near the gene for threonine deaminase, of the minimal cosmid set used in constructing a physical map of the genome of Hf. volcanii (13). Digestion of cosmid 126 DNA with Hind HI and Ssp I restriction endonucleases followed by hybridization with the lgd probe led to the identification of a 7 kb fragment. A sub-fragment containing the DHLipDH open reading frame was cloned

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

520 to create plasmid pT7-03. The mutant gene coding for 3-hydroxyl-3-methylglutaryl-coenzyme A reductase (hmg gene) of Hf. volcanii (which confers mevinolin resistance and has been used to construct a shuttle vector (11)) was employed to disrupt the Ipd gene in plasmid pNAT 01. Endonuclease Xho I cleavage sites within the lpd gene of plasmid pT7-03 and Xho I cleavage sites in the hmg gene in mutant Hf. volcanii (pBE HI) led to the incorporation of a 2 kb fragment imparting mevinolin resistance within the lgd gene thereby disrupting the production of DHLipDH. (cf\ Figure). pT7-03

pBE III Xho I

Xho I

\ 2.0 Kb , KBÜBIiMW

Sma I

r

—

Xho I

I0.6K I Kb I

mevinolin resistance

1.0 Kb

1

DIILipDH

Digest with Xho I to linearize the plasmid DNA.

Digest with Xho I and gel purify the 2.0 Kh fragment.

Xho I

Xbal

Xho I

T7-03 linearized by Xho I

Ligate Mev r fragment to linearized T7-03 to obtain DIILipDII-mev r insertion mutant. Sma I Xho I

t

t

sssssssss 0.6 Kb

Xho I

Xbal

t

-

SSSSSSSSï

2.0 Kb

1.0 Kb

pNATOl

Figure. Construction of the plasmid pNAT 01 carrying the disrupted lpd gene for dihydrolipoamide dehydrogenase through insertion of the hmg gene from Haloferax volcanii.

521 Hf. volcanii strain WFD11 (lacking plasmid pHV2 (11)) was transformed with plasmid pNAT 01 linearized with endonuclease Sma I. Transformants were selected on plates with rich or minimal media containing mevinolin. Genomic DNA from mevinolin-resistant colonies was isolated by lysis with SDS, phenol extraction and ethanol precipitation and subsequently digested with endonucleases Hind III and Ssp I.

The fragments were separated by

electrophoresis through 0.7% agarose gels, Southern transferred to Hybond N membrane and hybridized separately to lgd and hmg genes labelled with [alpha 32-P] dCTP. The ¡pd hybridization pattern expected for a endonuclease Hind III/Ssp I double digest of genomic DNA from wildtype WFD11 is a single 7 kb band. For strains with an lgd gene disrupted by a double recombination event the expected pattern was two bands with a summed molecular size of 9 kb since the 2 kb hmg fragment insert carries a Hind HI cleavage site. Transformants identified as VNN 21 and VNN 28 show this double recombination pattern yielding Ipd hybridization signals at 6.5 kb and 2.5 kb. Hybridization with an hmg gene probe produced a signal with one of these fragments for each of the transformants. Growths of Hf. volcanii strains VNN 21 and VNN 28 on rich and minimal media containing mevinolin were compared with the growth of wildtype WFD11 and a mevinolin-resistant transformant (VNN 23) produced by recombination at the resident hmg locus and thus having an intact ¡pd gene. The strains carrying the disrupted ]pd gene did not show any differences from the controls in growth rate in rich and minimal (glycerol) media. Assays for DHLipDH activity in controls in crude extracts yielded values of 0.006 units/mg for WFD111 and 0.005 for VNN 23. DHLipDH activity in strains VNN 21 and VNN 28 carrying the disrupted lpd gene was zero.

Discussion The lpd gene coding for DHLipDH activity in the extreme halophilic Archaea Haloferax volcanii has been disrupted. The loss of this dithio reductase activity has not affected the growth of the organism on either rich or minimal (glycerol as sole carbon source) media. Thus, dihydrolipoamide dehydrogenase activity does not appear to be essential for survival and optimal growth for Hf. volcanii. Archaebacteria remains unknown.

The role of this enzyme in the extreme halophilic

522 Acknowledgements Financial support from the Natural Sciences and Engineering Research Council of Canada (KJS) and the Medical Research Council of Canada (WFD) is gratefully acknowledged. The assistance of Dr. Steven W. Cline with transformations of Haloferax volcanii is gratefully acknowledged.

References 1.

Yeaman, S.J. 1989. Biochem. J. 257:625.

2.

Kikuchi, G., K. Hiraga. 1982. Mol. Cell. Biochem. 45:137.

3.

Williams, C.H. Jr. 1992. In: Chemistry and Biochemistry of Flavoenzymes Vol. i n (F. Muller ed.) CRC Press, Boca Raton, Ann Arbor, London, p. 121.

4.

Kercher, L., D. Oesterhelt. 1982. Trends in Biochem. Sci. 7:371.

5.

Danson, M.J. 1988. Adv. Microbiol. Physiol. 29:165.

6.

Danson, M.J., A. McQuattie, KJ. Stevenson. 1986. Biochemistry 25: 3880.

7.

Vettakkorumakankav, N., M.J. Danson, D.W. Hough, KJ. Stevenson, M. Davison, J. Young. 1992. Biochem. Cell Biol. 70: 70.

8.

Vettakkorumakankav, N., KJ. Stevenson. 1992. Biochem. Cell Biol. 70:656.

9.

Cline, S.W., W.F. Doolittle. 1989. Can. J. Microbiol. 35:148.

10.

Cohen, A., W.L. Lam, R.L. Charlebois, W.F. Doolittle, L.C. Schalkwyk. 1992. Proc. Natl. Acad. Sci. USA 89:1602.

11.

Lam, W.L., W.F. Doolittle. 1989. Proc. Natl. Acad. Sci. USA 86:5478.

12.

Niewlandt, D.T., C. J. Daniels. 1990. J. Bacteriol. 172:7104.

13.

Charlebois, R.L., L.C. Schalkwyk, J.D. Hofman, W.F. Doolittle. 1991. Biol. 222:509.

J. Mol.

Redox Properties of the FAD in the Active Site Mutants C44S and C49S of Escherichia coli Lipoamide Dehydrogenase

Nancy Hopkins and Charles H. Williams Jr. Department of Biological Chemistry, University of Michigan, Ann Arbor, MI 48109 Dept. of Veterans Affairs Medical Center, 2215 Fuller Rd. Ann Arbor, MI 48105

Introduction E. coli lipoamide dehydrogenase is a dimeric flavoprotein and a member of the pyridine-nucleotide disulfide oxidoreductase family of enzymes. As the terminal enzyme in the pyruvate dehydrogenase complex, it catalyzes the reduction of N A D + by dihydrolipoamide (1). Previous studies have shown that the enzyme is capable of accepting four electrons but, catalysis proceeds only through the two electron reduced state (1). The enzyme has two redox centers, the FAD and the disulfide. The cyteines which comprise the disulfide have distinct functions; Cys-44, participates in interchange with dihydrolipamide and Cys-49, particpates in a charge transfer interaction with the flavin (1). Both of these residues have been mutated to serines (C44S and C49S) (2) making it possible to study the redox behavior of the FAD in the absence of the disulfide. Previous studies of the wild type enzyme have used indirect methods to determine the redox potential of the FAD (3, and calculated from data in ref. 4). Two-electron reduced lipoamide dehydrogenase (EH2) is an equilibrium mixture of several species. In the pig heart enzyme, the charge transfer complex predominates so that the redox pair is effectively E o x and the charge transfer species. In lipoamide dehydrogenase from E. coli on the other hand, more than one EH2 species is present in a redox equilibrium including the charge transfer complex and the disulfide/FADH2 species. In wild type enzyme therefore, it is important not to associate the E/EH 2 potential with the disulfide/dithiol potential or the EH2/EH4 potential with the FAD/FADH2 potential except as an approximation. In C44S and C49S, the disulfide is absent and the redox potential of the FAD can be measured. These potentials can be compared to the EH2/EH4 potentials of wild type enzyme keeping the caveat in mind.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

524 Results Reduction of the enzymes at pH 7.6 using NADH and NADPH indicated that C44S was more difficult to reduce than C49S since a larger excess of reductant was required to approach full reduction. This indicated that the redox potential for C44S was lower than that of C49S. An excess of NADH and NADPH was needed to sufficiently reduce both enzymes which also showed that both enzymes had redox potentials lower than both reduced pyridine nucleotides. Using methyl viologen, a low redox potential dye (Em = -440 mV), with the xanthine/xanthine oxidase (X/XO) system as electron donor (5), simultaneous reduction of the dye and the enzymes confirmed that the redox potential of the enzymes was quite low. In order for the X/XO system to work, the dye couple and the enzyme couple must remain in equilibrium. Nernst plots were constructed (Figure 1) using a theoretical slope of 29.5 mV to fit the data. Redox potentials were calculated for both enzymes from the NADH, NADPH and X/XO data (Table 1). The plot relating the redox potential of C49S to pH showed a break in slope at pH 7.6 (data not shown). This indicated pK a is presumably associated with the ionization of the FADH2 in the reduced form of the enzyme.

Discussion The higher redox potential determined using NADH indicated that this physiological substrate, bound more tightly to the reduced forms of the enzymes than the oxidized forms. The redox potentials determined using the X/XO system were lower than those determined using the reduced pyridine nucleotides which also indicated binding of NADPH as well as NADH to the enzymes. This behavior was not unexpected since NADPH is an analog of NADH. It can also be concluded from these data that NADH binds more tightly to the enzymes than does NADPH and that the binding is tighter for C44S than for C49S because of the larger differences in redox potentials as seen in Table 1. At pH 7.6, both enzymes have a lower redox potential than the wild type enzyme (EH2/EH4, Em = -314 mV) (calculated from data in ref. 4). The low redox potential for C44S (-417 mV) can be explained by the thiolate which causes the enviroment around the flavin to be more negative, making it more difficult to reduce. The low redox potential for C49S (-386 mV) in comparison to wild type enzyme (EH2/EH4) is difficult to explain without further

525

J3 W

W

-440

-440

-1.0

0.0

1.0

-1.0

Log (Eox/Ered)

0.0

1.0

Log (Eox/Ered)

Figure 1. Nernst Plots of Redox Titrations of (A) C44S and (B) C49S at pH 7.6. The system potentials, Eh, of the three titration methods (NADH, NADPH and X/XO) are plotted as a function of the log (Eox/Ered) at 25°C. The solid lines represent the data fitted to a theoretical slope of 29.5 mV. Squares, NADH data; triangles, NADPH data and circles, X/XO data. TABLE 1. Comparison of Redox Potentials for E. C. LipDH C44S and C49S Difference (mV)

Enzyme

Method

Em (mV)

C44S

X/XO

-417

NADPH

-375

42

NADH

-353

64

X/XO

-386

NADPH

-363

23

NADH

-335

51

C49S

526 experiments but, it is comparable to wild type enzyme from pig heart which has a redox potential (EH2/EH4) of -382 mV (3). In conclusion, these data show that the enviroment surrounding the flavin modulates the redox potential of the flavin. Thus, the thiolate present in C44S which causes the enviroment around the flavin to be more negatively charged, lowers the redox potential of the flavin, making its redox potential lower than that of C49S and wild type enzyme (EH2/EH4). It is possible that having the hydroxyl group of the serine close to the flavin as seen in C49S increases the electronegativity of the enviroment surrounding the flavin and this would explain why C49S has a lower redox potential than wild type enzyme (EH2/EH4). These Cys to Ser mutants of lipoamide dehydrogenase have similar relative redox potentials to those observed in analogous mercuric reductase active site mutants (C135S, Em = -428 mV; C140S, Em = -375 mV, ref. 6) and this is also an indication that these types of mutations cause the redox potential of the flavin to be lowered.

Acknowledgements Support of the Department of Veterans Affairs (CW), NIGMS grant GM21444 (CW), and Rackham School of Graduate Studies, Univ. Michigan (NH) is gratefully acknowledged. References 1.

Williams, C. H. Jr. 1992. In: Chemistry and Biochemistry of Flavoenzymes, Vol.III (Muller, F., ed.). CRC Press, Inc., Boca Raton, pp. 121-211.

2.

Hopkins, N., Russell, G. C., Guest, J. R. and Williams C. H., Jr. 1991. In: Flavins and Flavoproteins (Curti, B., Zanetti, G. and Ronchi, S., eds.). Walter de Gruyter and Co., Berlin, 581-584.

3.

Matthews, R. G. and Williams, C. H. Jr. 1976. J. Biol. Chem. 251, 3956-3964.

4.

Wilkinson, K. D. and Williams, C. H. Jr. 1979. /. Biol Chem. 254, 852-862.

5.

Massey, V. 1991. In: Flavins and Flavoproteins (Curti, B., Zanetti, G. and Ronchi, S., eds.). Walter de Gruyter and Co., Berlin, 581-584.

6.

Distefano, M. D., Au, K. G. and Walsh, C. T. 1989. Biochemistry. 28, 11681183.

R- AND S-DIHYDROLIPOIC ACID DERIVATIVES AS SUBSTRATES OF LIPOAMIDE DEHYDROGENASE

L. David Arscott and Charles H. Williams, Jr. Department of Veterans Affairs Medical Center and Department of Biological Chemistry, University of Michigan, Ann Arbor, MI 48105 USA

The R-lipoic acid isomer was found to be the natural substrate isolated from the a ketoglutarate multienzyme complex where it was covalently attached in amide linkage with the e-amino group of a specific lysine residue in dihydrolipoyl transacetylase, the core component of the complex (1). Previous studies indicated that both isomers react with the complex although at disparate rates (2). Dihydrolipoic acid is a very poor substrate of lipoamide dehydrogenase above pH 6.3 (2); thus, dihydrolipoamide is the preferred substrate. Reduction of lipoamide dehydrogenase by R,S-dihydolipoamide displays biphasic kinetics. It was hypothesized that R - dihydrolipoamide reacted rapidly and that S - dihydrolipoamide accounted for the slow rate either by reacting with the enzyme directly or by nonenzymatically reducing the R-lipoamide already formed (3,4,5). Recent work, measuring the rate of the interchange reaction of S-lipoic acid with R,S-dihydrolipoic acid, indicated that the rate of the nonenzymatic reaction was very slow (6). The apparent lack of significant biological activity of the S-dihydrolipoamide and the lack of the diastereomers has meant that most reported data involving the biological or catalytic properties of this substrate have used the R,S-racemic mixture. In the course of a study of a lipoamide dehydrogenase altered in the active site base catalyst, H444Q, data was obtained suggesting that both R- and S-dihydrolipoamide could react directly with the enzyme (7). Thus, this study to characterize the specificity of the lipolyl binding site. Results and Discussion The kinetics of the reaction of R,S-dihydrolipoamide with excess 3-acetylpyridine adenine dinucleotide catalyzed by pig heart lipoamide dehydrogenase are distinctly biphasic as shown in Figure 1 with half the reaction taking place in each phase. APyAD + , a more readily oxidized NAD+ analog, allows the reaction to go virtually to completion.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

528 The insert in Figure 1 plots both the fast and slow rate versus the enzyme concentration. A constant ratio of 10 was found for the fast to slow rate. This would be expected only if both isomers were direct substrates. If the slow rate was a response of the nonenzymatic interchange of S-dihydrolipoamide with R-lipoamide, the slow rate should not increase as a function of increased enzyme concentration since there is no increase in the initial concentration of R,S-dihydrolipoamide. It should be noted that these kinetics are not at saturating conditions, however steadystate kinetics with the S-isomer a possible alternate substrate should be competitive with the R-isomer(8). 1.0

0.0 0.4 0.8 Enzyme ((iM) 100

200

300 400 Time (s)

500

Figure 1. Full reaction turn-over kinetics following the reduction of APyAD+ by R,S-dihydiolipoamide with pig heart lipoamide dehydrogenase at 25 °C in 0.1 M Na-K PO4, 0.30 mM EDTA, pH 7.6. The solution of 247 )iM APyAD + plus enzyme was initiated by adding R,Sdihydrolipoamide to a concentration of 60 fiM. Curves are for increasing lipoamide dehydrogenase concentration of: 1, 0.097

/iM; 2, 0.29 /iM; 3, 0.57 /xM ; 4, 0.77 ftM. Curves were fitted

60 to two exponentials and the respective rate constants plotted in the insert.

The methyl esters of R- and S-dihydrolipoate have been synthesized from R- and Slipoic acids (reduction preceded esterification). Steady-state studies at pH 8.9 with the methyl ester of R,S-dihydrolipoate gave an extrapolated V , ^ of 9060 min-1 which is the same as had been determined for R,S-dihydrolipoamide (5). The double reciprocal plots in Figure 2A graph the steady-state kinetics using the R-isomer alone. Both substrates are varied maintaining their concentrations at a constant ratio. A Ping-Pong Bi Bi mechanism under these coonditions will give converging straight lines with increasing slopes when plotted in double reciprocal form (8). Secondary plots gave Km values of 263 /¿M and 580 /xM for the methyl ester of Rdihydrolipoate and APyAD+, respectively and a V^^ of 7755 (± 1070) min-1. The S-isomer also gave converging lines in reciprocal format with a V , ^ of 1274 (±

529 250) miir 1 (Figure 2B). Secondary plots gave KJJ, values of 753 /xM and 418 /xM, for the methyl ester of S-dihydrolipoate and APyAD+, respectively. Also, the sum of the extrapolated V , ^ values for the R- and S- isomer of 9030 mirr 1 is the same as the value with the racemic R,S-dihydrolipoic methyl ester of 9060 min -1 .

Figure 2. Double reciprocal plots of the kinetics of reaction of R-dihydrolipoic methyl ester (DHLME, the methyl ester of R- or S-dihydrolipoic acid) or S-DHLME and A P y A D + catalyzed by pig heart lipoamide dehydrogenase. Assays were done in 70 mM PPj, 0.3inM EDTA, pH 8.9, 10% ethanol (v/v), 23 °C. A: ca. 3 nM enzyme started the reaction. Curves representing concentrations at constant ratios are: 1, 0.6; 2, 1.3; 3, 2.5; 4, 4.2; 5, 6.3. B: ca. 30 nM enzyme started the reaction. Curves representing constant ratios are: 1, 0.3; 2, 0.9; 3, 1.9; 4, 3.7.

Figure 3. Inhibition» of the reaction of R - dihydrolipoic methyl ester with APyAD + by S-dihydrolipoic methyl ester. Assays at 25 °C, contained 70 mM Tris, pH 8.9, 2 0 % e t h a n o l , 8 6 2 /iM APyAD + and ca 6 nM enzyme. The concentration of S-DHLME is: 1, 0; 2, 760 /jM; 3, 1520 /iM; 4, 1900 /¿M.

500 1000 R-DHLME (jiM)

1500

530 Inhibition kinetics varying the R-isomer in the presence of fixed levels of S-isomer as shown in Figure 3, also indicate that it is an alternate substrate (8). In this reaction the turn-over decreases as the S-isomer is increased and the apparent K m decreases (the APyAD+ was at non-saturating conditions). Solubility of S-DHLME limited the level of the inhibitor. In summary these experiments indicate that the S-isomer can react directly with lipoamide dehydrogenase, since the turnover rate using the S-isomer in the enzymatic reaction is much faster than the interchange reaction of S-lipoic acid with R,S-dihydrolipoic acid (6). Acknowledgements Samples of R- and S-lipoic acids were kindly supplied by Drs. J. R. Guest and V. Massey. Support of the Department of Veterans Affairs and grant # GM21444 from the National Institute of General Medical Sciences is gratefully acknowledged. References 1.

Reed, L.J., Gunsalus, I.C. et al., (1953) J. Am. Chem. Soc. 75,1267-1270.

2.

Sanadi, D.R., Langely, M. and Searls, R.L. (1959) J. Biol. Chem. 234, 178-182.

3.

Massey, V., Gibson, Q.H. and Veeger, C. (1960) Biochem. J. 77, 341-351.

4.

Wilkinson, K.D. and Williams, C.H.,Jr. (1979) J. Biol. Chem. 254, 852-862.

5.

Matthews, R.G., Ballou, D.P. and Williams, C.H.,Jr. (1979) J. Biol. Chem. 254, 4974-4981.

6.

Yang, Y-S. and Frey, P.A. (1989) Arch. Biochem. Biophys. 268, 465^t74.

7.

Williams, C.H. Jr., Arscott, L.D., Gamm, D. Hopkins, N., Allison, N. and Guest, J.R. (1990) In: Flavins & Flavoproteins (Curti, B., Ronchi, S. and Zanetti, G. Eds.) Walter de Gniyter, Berlin-New York pp. 577-580.

8.

Segel, I.H. (1975) In: Enzyme Kinetics (Segel, I.H., Ed.) p. 793, John Wiley and Sons, New York.

Comparative Study on Flavin Radical Formation of Llipoamide Dehydrogenase and Glutathione Reductase by Photoreduction

Kazuko Maeda-Yorita

and

Kenji Aki

Division of Enzyme Regulation, Institute for Enzyme Research, The University of Tokushima, Tokushima 770, Japan

Introduction Among the members of the pyridine nucleotide - disulfide oxidoreductase family, lipoamide dehydrogenase (LD) and glutathione reductase (GR) are closely related in their structure and reaction mechanism (1). One of the differences in these two enzymes is the stability of the charge transfer (CT) complex from an active site thiolate anion to the oxidized form of FAD in the two-electron-reduced state (EH2).

In order to get information on redox states of the flavin, we

photoreduced each enzyme to its one-electron-reduced species in the presence of EDTA and 3,10-dimethyl-5-deazaisoalloxazine (2), and analyzed by ESR and absorption spectra.

Results and Discussion The first observed spectral species after photoreduction of pig heart LD in the absence of /3-NAD+ has an absorbance peak at 455 nm like that of the oxidized form and has increased absorption up to 680 nm, indicating an equilibrium between oxidized, reduced, and neutral radical species (Fig 1A, curve 2).

This

spectrum is labile, especially at acidic pH, and changes to the second spectral species, the CT complex characteristic of the catalytically important EH2 form, where the absorption peak is blue-shifted and the longer wavelength band is centered at 530 nm (Fig.lA, curve 3). (Fig.lA inset)

Disappearence of a weak ESR signal

corresponds to the above absorbance spectral

change.

Photoreduction of pig heart LD in the presence of /3-NAD+ results in a stable

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

532

Figure 1. Photoreduction of pig heart LD and yeast GR. A; 18 nM LD (curve 1) was reduced photochemically in 0.1 M potassium phosphate buffer, containing 0.3 mM EDTA, at pH 6.3. In the absence of /J-NAD+, the first transient species (curve 2) changed to the subsequent stable species (curve 3). In the presence of five equivalents of /J-NAD+, the single stable species(ctv/ve 4) was obtained. inset, ESR signal development of 180 nM LD on irradiation. ESR signal intensity was normalized by the enzyme FAD concentration for comparison with that of GR. B; 17 |iM GR (curve 1) was photoreduced in the absence (curve 2) or presence (curve 3) of five equivalents of /J-NADP+ in 0.1 M potassium phosphate buffer, containing 0.3 mM EDTA, at pH 7.6. inset, ESR signal development of 79 ^M GR on irradiation.

533 spectrum (Fig.lA, curve 4).

A broad long wavelength absorption band of

unknown origin beyond 800 nm is also present. 600 nm is increased at acidic pH.

The absorption intensity around

The observed ESR signal is intense, with a g-

value of 2.005 and a peak to peak signal width of 1.5 mT, with a shoulder and remarkable power saturation.

These spectral properties indicate an equilibrium

between neutral and anionic flavin radicals and CT interactions between some redox states of /3-NAD+ and FAD. spectral properties. photoreduction.

Other NAD(P)+ analogs do not cause these

E. coli LD shows

similar

absorption

spectra

on

Relatively intense absorbance around 600 nm of E. coli LD is an

indication of the stability of the neutral flavin radical. Photoreduction of yeast GR at pH 7.6 in the absence of 0-NADP+, on the other hand, results in the stable absorption spectrum of the anionic flavin radical (Fig. 1B, curve

2).

This species is ESR positive, and has identical ESR

parameters to those found in photoreduction of LD in the presence of /3-NAD+. The presence of /J-NADP+ perturbs the absorption spectrum of the anionic flavin radical, and produces an additional longer wavelength band beyond 800 nm (Fig.lB, curve 3).

No significant pH dependence on the absorption intensity

beyond 500 nm was observed. The order of the stability of the neutral flavin radical of these enzymes in the presence of /3-NAD(P)+ is inversely parallel to that of the stability of the CT complex from thiolate anion to FAD.

Alkylation of the redox active dithiol by

iodoacetamide produces different spectral species, LD(EHR) and GR(EHR), respectively (3,4).

Although photoreduction of LD(EHR) shows complete

bleaching at 455 nm as in free flavin, JGR(EHR) resists against bleaching at 462 nm except for that due to release of FAD from the enzyme.

Photoreduction of

LD(EHR) or GR(EHR) in the presence of /J-NAD(P)+ is, however, similar to that of the intact enzymes.

This indicates that the direct interaction of bound /3-NAD(P)+

with FAD at the active site is responsible for the stability of the flavin radical in both the enzymes, as in scheme 1. Therefore the observed phenomena could be due to a correlation between interaction of the redox active disulfide with FAD at the C4a position and that of pyridine nucleotide with FAD at the N5 position, indicating subtle orientational differences of the bound pyridine nucleotide and the redox active disulfide-FAD pair at the active sites of the two enzymes.

534

Scheme

1

References

1.

Williams, C.H., Jr. (1976) in The Enzymes, 3rd Ed.Vol.13. (P. D. Boyer, ed.) pp89-173

2.

Massey, V., and Hemmerich, P. (1978) Biochemistry, 17, 9-17

3.

Thorpe, C „ and Williams, C.H., Jr. (1976) J. Biol. Chem., 251, 7726-7728

4.

Arscott, L.D., Thorpe, C., and Williams, C.H., Jr. (1981) Biochemistry, 20, 1513-1520

The Pyruvate Dehydrogenase Complex from Azotobacter vinelandii

A. de Kok, A. Berg, W. van Berkel, A. Fabisz-Kijowska and A. Westphal Department of Biochemistry, Agricultural University, Wageningen, NL F. van den Akker, A. Mattevi1 and W.G.J. Hoi Department of Structural Biology, University of Washington, Seattle, USA

Introduction Pyruvate dehydrogenase multienzyme complexes from prokaryotes consist of three components, pyruvate dehydrogenase or Elp, acetyltransferase or E2p and lipoamide dehydrogenase or E3. The E2p component from Azotobacter vinelandii is a highly segmental structure and consists of three lipoyl domains, an Elp/E3 binding domain and a core-forming catalytic domain. The domains are separated by flexible linkers of 20 to 40 amino acids. The cofactor lipoamide, covalently linked via an amide bond to a lysine of each lipoyl domain, acts as a "swinging" arm and shuttles between the active sites of the three components. Progress in structural insight in pyruvate dehydrogenase complexes is described in a recent review paper (1). The crystal structures of A. vinelandii (2) and Pseudomonas fluorescens (3) E3 are closely related. The rms difference for all 932 Ca atoms is 0.55A with 84% amino acid identity. Small structural variations will be discussed here that must provide the basis for the differences in thermostability and redox properties. The intact E2p component could not be crystallized, probably due to the disorder created by the flexible linker regions. The N-terminal lipoyl domain was solved by NMR (to be published), while the catalytic domain was solved

by X-ray crystallography (4).

So far structural information on the integration of the individual components into a functional complex is lacking. The availability of a structure of the binding domain in complex with E3 would be a first approach to gain insight in the E2p/E3 interaction. Some properties of the E3-binding domain interaction and efforts to purify this complex for crystallization purposes are described here. ') Present address: Dept of Genetics and Microbiology, University of Pavia, Italy

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

536 Results and discussion The lipoamide dehydrogenase component The three dimensional structures of the lipoamide dehydrogenases from A. vinelandii, P. putida and P. fluorescens

(2,3,5) revealed a dimer of approximately 100

kDa with subunits consisting of four domains. The two active sites reside at the interface of the subunits with the redox active disulfide close to the C4a position of the flavin ring. This ring separates the nicotineamide binding pocket from the lipoamide binding region of the active site (fig 1). The latter is at the si-side of the flavin. In the P. fluorescens and P. putida structures, the C-terminal residues form a tail which folds back into the active site approaching the si-side of the flavin. The tail significantly contributes to the subunit-subunit interaction and provides, besides hydrophobic contacts, two H-bonds with residues (Tyrl6 and Gln24, P. fluorescens numbering) of the opposite chain (fig 1). The C-terminus of the A. vinelandii structure is much less well defined. All biochemical data indicate however that in solution the C-terminus in A. vinelandii E3 adopts the same conformation as in the P. fluorescens structure (6).

Fig 1. A schematic diagram of the catalytic center of P. fluorescens lipoamide dehydrogenase. The drawing outlines the two channels separated by the flavin ring and forming the substrate binding sites. The side chains of Tyrl6 and His470 are H-bonded with each other and more than 6 A away from the reactive disulfide bridge. Nevertheless they exercise a key influence on the redox properties of the enzyme.

537 The properties of several active site mutants of the A. vinelandii enzyme have been described (6-8). Tyrl6 and the C-terminal tail are quite far removed from the redox active disulfide, but mutants caused rapid overreduction to the four-electron reduced state (EH4). In Y16F it is obviously not possible to form the hydrogen bond with His470 in the other subunit as is observed in wild type (3). In a C-terminal mutant lacking the last nine amino acid residues (A9) an additional H-bond and many hydrophobic contacts with the other subunit are missing. In Y16F the melting temperature (t,,,) is 8° lower than in wild type, while in A9 it is 15° lower. A relation between oveireduction and dimer (thermo) stability therefore has been suggested (6). As it is well known that lipoamide dehydrogenases from several sources differ in their stability against overreduction, the question arises whether this relation is a general feature. A comparison was therefore made between the structures from A. which can be overreduced easily, and P. fluorescens,

vinelandii,

which is quite resistant to

overreduction (3). P. fluorescens E3 has a ^ which is 6° higher than the enzyme from A. vinelandii (6). The flavin and disulfide bridge environment is identical and no side chain mutations occur within 6.7 Á from the flavin. All amino acid charges within 9 Á distance from the isoalloxazine are conserved. The most significant change is a replacement of Ser449 and Thr452 in P. fluorescens

E3 by two alanines in A. vinelandii E3. Both

residues are not only part of the upper interface region, but also part of the active site, because they flank the His450-Pro451 sequence (fig 1). The replacement does not affect the protein conformation. In A. vinelandii E3, the buried carbonyl oxygen of residue 449 is an unsatisfied H-bond acceptor. This burial is entirely caused by interactions with the other subunit. Therefore the T449A replacement could affect the dimer stability. The reverse effect is also observed: overreduction leads to dissociation (9). Limited proteolysis provides a sensitive tool for probing dissociation. The oxidized wild type enzymes are stable against a variety of proteases, but the overreduced enzymes are rapidly attacked. With trypsin the first cleavage is at Arg391 (9). This residue is buried in the subunit interface of the oxidized enzyme and forms a salt-bridge with Asp61 in the other subunit (3). These and other experiments indicate that reduction of lipoamide dehydrogenase to the EH4 state is accompanied by conformational changes promoting subunit dissociation. A tighter subunit association could decrease the redox potential of the EH2/EH4 couple, thus preventing overreduction. The structural basis for a diminished subunit interaction in the EH4 state is unknown. Binding of FAD to the monomeric apoenzyme yields initially a monomeric holoenzyme (10). The FAD is weakly bound and its spectroscopic properties (weak fluorescence, blue shift of visible absorption) indicate

538 that it is solvent accessible. In a slow reaction with a high activation energy the holo dimeric enzyme is formed. It is speculated that binding of FAD causes strain in the molecule. Part of the energy increase could be compensated by

subunit-subunit

interactions. Removal of FAD or reduction of the flavin (leading to a nonplanar conformation) may lead to a relaxed state with less favorable subunit-subunit interactions, especially because the N3 of the isoalloxazine itself is part of the interface. So far it has not been possible to obtain structures with bound cofactors or of reduced enzymes. The exception is a structure of the P. putida enzyme in complex with NAD + (5). In this structure the nicotineamide ring is in the "out" conformation, comparable to the conformation of NADP* in glutathione reductase (11). To obtain more insight into the reaction mechanism the P. fluorescens

structure was used to dock the

substrate R-dihydrolipoamide into the active site by a molecular modelling experiment. Electron and proton transfer of dihydrolipoamide to NAD+ is interfaced by the catalytic center of E3: His450, the Cys48-Cys53 disulfide and FAD, all located deep in the active site. In the course of catalysis one of the sulfurs of the substrate donates a proton to the active site base His450 followed by the formation of a mixed-disulfide bond intermediate with Cys48 (12). Therefore an important criterium for a correct binding mode is the ability of one of the sulfurs of dihydrolipoamide to approach closely these two residues. It is not known at present which of the two sulfurs, S6 or S8, of the substrate forms the mixed-disulfide bond with Cys48. The starting conformation of dihydrolipoamide was obtained by energy minimizations using the DREIDING II forcefield implemented in the BIOGRAF software package (13). Two starting models were obtained, one for each sulfur positioned near Cys48 and no initial bad contacts were obtained. In both orientations this was followed by energy minimizations and molecular dynamics simulation, resulting in two possible binding modes of the substrate. The ability of the particular sulfur of the substrate to approach the Cys48, as required for catalysis, was tested to distinquish between the two possibilities. This was achieved by decreasing that distance by applying, in small repeating steps, a decreasing harmonic distance restraint which is each time followed by energy minimizations molecular dynamics simulation at 300K to allow the substrate to relax. When the S6-atom is positioned near Cys48, the initial S(lip(SH)2)-S(Cys48) distance is 5.6 Á. Upon decreasing this distance to 3.5 Á, a 50 kcal/mol increase in internal energy plus van der Waals energy is calculated. When the S8 atom was positioned near Cys48 with an initial distance of 4.1 Á, only 4 kcal/mol increase was obtained. More extensive models in which also surrounding side chains of E3 were

539

Ser389'

Fig 2. Schematic drawing of lipoamide bound to P. fluorescens docking. Broken lines indicate distances in A .

E3 as revealed by

allowed to move showed a similar trend. The S(lip(SH)2)-Ne2(His450) distance, important for proton abstraction, was also monitored upon approaching Cys48. This distance is 3.5 A for the S8-position and 4.1 A for the S6-position at the beginning of the approaching Based upon these results it is postulated that the S8-atom of dihydrolipoamide forms the mixed-disulfide intermediate and the binding mode obtained by docking is shown schematically in fig 2. The residues postulated in being involved in substrate recognition are conserved in other species (14) strengthening the postulated binding mode. Comparing the binding pocket in E3 with that of dihydrolipoamide transacetylase with bound dihydrolipoamide

(15), showed no conserved finger-print for dihydro-

lipoamide binding. The conformations of dihydrolipoamide in both proteins are also different. However, the solvent accessible surface and hydrophobicity

of both the

substrate and the protein buried upon binding shows a remarkable similarity. Very interesting is the presence of a long second channel perpendicular to the substrate binding channel also ending at the active center (fig 3). Because of its length of more than 20 A to reach the disulfide this channel cannot be involved in dihydrolipoamide binding. It is possible that this channel might be an outlet for water molecules pushed away by the substrate entering via the other channel. Of interest is that three a -

540

Fig 3. Dihydrolipoamide bound to P. fluorescens E3 with a solvent accessible surface of the interior of this protein reveals a second channel to the active center. Also shown are the His450-Glu455 diad, the disulfide and the flavin ring.

helices located along the second channel (involving residues 12-25, 325-342 and 452-466, P. fluorescens

numbering), together with a fourth helix (involving residues 48-65)

point with the positive end of their dipole towards His450. This could compensate the rise in pKa expected from the interaction with Glu455. In this manner the pKa of the histidine can be suitably balanced for both proton abstraction and donation during catalysis. The acetyl transferase component The structure of the N-terminal lipoyl domain was recently solved by homo- and heteronuclear NMR techniques (to be published). It consists of two antiparallel (J-sheets, each consisting of four strands. The lipoyllysine group is exposed to the solvent and located in a single type-I turn at the comer of one of the sheets. Its overall structure is very similar to that of Bacillus

stearothermophilus

(16), despite its low amino acid

identity of 31%. The C-terminal catalytic domain was solved by X-ray crystallography (4). It consists of eight trimers in the corners of a hollow, truncated cube. The active sites consist of 30A long channels running across the entire dimer interface within each trimer. The entrance of the channel at the outer face of the cube forms the lipoamide binding site, while (Acetyl)CoA enters the channel from the hollow center of the cube. Soaking experiments with substrates and crystal structures of active site mutants have yielded a

541 wealth of information on binary complexes with E2p and the mechanism of the transacetylase reaction (15). The S8 thiol of dihydrolipoamide is the reactive SH group, involved in the initial proton

abstraction

by

a histidine,

His610

(15). In

binding

of

CoA,

but

not

dihydrolipoamide, Asn614 moves from a position in which it forms a hydrogen bridge with Asp508 to a position in which it is hydrogen-bonded to N8l of His610. In this manner the protonated histidine is stabilized. Surprisingly, S0 2 0H", generated by hydrolysis of dithionite used to prevent oxidation, is bound in the center of the active site. The binding mode shows that the active site can accomodate a negative charge, stabilized by the positively charged His610 and by the formation of a H-bond with the side chain of Ser558. This complex may therefore serve as a model for the stabilization of the putative negatively charged tetrahedral intermediate. Interaction between acetyltransferase and lipoamide dehydrogenase Protein engineering experiments with E2p indicate that E3 interacts only with the binding domain, while Elp interacts with both the binding domain and the catalytic domain (1,17). At present the residues of E3 that are involved in the interaction with the E2p component are unknown. The E3-monomer, both the apo and holoenzyme, does not interact with E2p (18). Therefore it seems likely that structural elements from both subunits near the dimer interface of E3 are involved in the E2p-E3 interaction. The interaction has several effects on catalysis by E3. Bound in the complex, E3 is less sensitive to inhibition by NADH. The initial rate in the lip(SH)2 -> NAD+ reaction increases, especially at low ionic strength. The rapid modification of Cys48 by maleimides in NADH reduced unbound E3 is fully prevented in the E2pE3 subcomplex. Quite remarkably, binding of the Tyrl6 and A9 mutants to E2p restore the physiological activity to values between 55 and 100% (Table I). This activation was also observed with the purified binding domain preparation and was used to distinguish between bound and unbound E3 during purification (see below). Turnover studies by stopped flow indicated that E3(Y16F) was as active as wildtype during the initial phases of the reaction. The addition of E2p had no effect on this initial phase. The activating effect of E2p was completely due to the protection against dead-end inhibition by NADH. When E2p was added to E3(Y16F) within seconds after the initial phase no activation was observed, demonstrating that irreversible structural changes take place upon reduction, preventing the interaction with E2p. Activation provides a sensitive assay for binding and also allows conclusions to be drawn on the integrity of the binding domain obtained by engineered E2p proteins. A set

542 Table I. Effect of E2p on Activity of E3 Wild-type and Mutants. Enzyme

Lip(SH)2 -> NAD*

Lip(SH)2 -> AcPyNAD +

activity (%)

activity (%)

-E2p

+E2p

-E2p

+E2p

100

138

100

119

A5

85

138

86

90

A9

ROH + H2O + NAD(P)+; Ref. 1], This activity has been shown to serve a protective function directed against oxygen-linked DNA damage. Characteristics of AhpF and AhpC proteins are summarized in Table 1 below. The flavin-containing AhpF component was shown by sequence comparisons to be related to thioredoxin reductase (TR), a well-characterized flavin- and redox-disulfide-dependent oxidoreductase (2). Although AhpC is not similarly homologous with the small redoxdisulfide-containing protein substrate of TR, thioredoxin, the direct interaction of the two

Table 1. Characteristics of the Protein Components of S. typhimurium AhpR. AhpF

AhpC

—57 kDa/subunit, dimeric — 1 FAD/subunit —6 half-cystine residues/subunit, in 3 pairs -homology with thioredoxin reductase (TR) (36% sequence identity over amino acids 209-515 of AhpF, redox-active halfcystine residues of TR fully conserved) - A h p C reductase, DTNB reductase, diaphorase, and oxidase activities

- 2 1 kDa/subunit —no chromophoric cofactor - 2 half-cystine residues/subunit, in intrasubunit disulfide bond —homology with proteins from a wide range of organisms, first and possibly second half-cystine residue(s) of AhpC conserved -peroxide reductase activity in the presence of AhpF and reduced pvridine nucleotides

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

584 proteins and subsequent redox activity of the small protein indicate similarities between the two systems. Mechanistically, AhpC as a heme- and metal-independent peroxide reductase can be related to two other relatively well characterized enzymes, the streptococcal NADH peroxidase (3) and the eukaryotic glutathione peroxidase (4). In each of these cases, single cysteine or selenocysteine residues cycle catalytically through unusual reversibly oxidized derivatives, cysteine-sulfenic acid (Cys-SOH) and selenenic acid (Cys-SeOH), respectively. Based on comparisons with the TR-thioredoxin system, the two heme-independent peroxidatic mechanisms described above, and the half-cystine contents of the AhpR proteins, we have proposed the following catalytic mechanism, including redox-active protein disulfides on each component protein. /S(H+)

R 0 0 H

We report here data confirming the ability of AhpF to catalyze electron transfer between NAD(P)H and the cystine disulfide of AhpC; the resulting dithiol of reduced AhpC is readily reoxidized on addition of ethyl hydroperoxide.

Results and Discussion In order to establish the stoichiometry of the reduction of AhpC by pyridine nucleotides (with AhpF mediating the electron transfer) and to determine whether or not the single protein disulfide of AhpC is the acceptor of those electrons, anaerobic NADH titrations of AhpC in the presence of a catalytic amount of AhpF were carried out, followed by DTNB assays to detect protein thiols. Initial experiments indicated that the stoichiometry of NADH-linked reduction could be readily determined by spectral analysis focusing on the absorbance change at 340 nm during the titration; addition of the first eq of NADH results in very little increase in

585 absorbance at 340 nm, while subsequent additions result in the increase in absorbance expected for the known amount of NADH added (Fig. 1). The overall stoichiometry for oxidation of added NADH in the presence of AhpC was 1.13 ± 0.03 (N=5) mol of NADH per mol AhpC. DTNB assays for free thiols performed following the anaerobic addition of one or more eq of NADH to AhpC indicate the generation of 2.00 ± 0 . 1 3 (N=6) reactive thiols per AhpC. These DTNB titrations require prior removal of interfering excess NADH by exposure to air to take advantage of the NADH oxidase activity of AhpF. Where ethyl hydroperoxide is added prior to DTNB assay, the thiol titer obtained is proportionally lower. As shown in Figure 2, 0.97 eq of ethyl hydroperoxide is required to fully oxidize the two thiols of reduced AhpC and regenerate the cystine disulfide. Our results strengthen the proposal that AhpF acts on AhpC in a manner analogous with the TR/thioredoxin system, reducing the redox-active disulfide of AhpC to the dithiol. These

0.12

0.08

0.04

0

300

350

400

450

500

Wavelength, nm Figure 1. NADH titration of AhpC. AhpC was titrated anaerobically with NADH in the presence of a catalytic concentration (200-fold less) of AhpF. Spectra shown, in order of increasing A340, were obtained following the addition of 0, 0.38, 0.76, 1.14, 1.23, 1.33, 1.42, 1.52 and 1.61 eq NADH/subunit AhpC, respectively.

586

Ethyl hydroperoxide, eq Figure 2. Oxidation of the dithiol of reduced AhpC by ethyl hydroperoxide. Reduced AhpC generated by the anaerobic addition of NADH was reacted with varying amounts of ethyl hydroperoxide prior to spectral thiol assays using DTNB. Depicted here is the resulting thiol titer expressed as a function of ethyl hydroperoxide added. results also strongly support the direct involvement of the nascent dithiol of AhpC in the reduction of organic hydroperoxides. As shown in the proposed catalytic mechanism, oxidation of one of the two cysteine thiols of reduced AhpC followed by condensation between the oxidized cysteine derivative and the second cysteine thiol is chemically feasible, given the known hydrogen peroxide-mediated oxidation of a cysteine thiol to generate the cysteine-sulfenic acid in NADH peroxidase (3). The redox-active cystine of AhpC, if of catalytic importance, is unusual in that the separation between the two half-cystine residues (CyS46 and Cysigs) in the primary sequence is substantial.

References

1.

Jacobson, F. S„ R. W. Morgan, M. F. Christman and B. N. Ames. 1989. J. Biol. Chem. 264, 1488-1496

2.

Tartaglia, L. A., G. Storz, M. H. Brodsky, A. Lai and B. N. Ames. 1990. J. Biol. Chem. 265, 10535-10540

3.

Poole, L. B. and A. Claiborne. 1989. J. Biol. Chem. 264, 12330-12338

4.

Epp, O., R. Ladenstein and A. Wendel. 1983. Eur. J. Biochem. 133, 51-69

FLAVOPROTEIN PEROXIDE AND DISULFIDE REDUCTASES AND THEIR ROLES IN STREPTOCOCCAL OXIDATIVE METABOLISM

A1 Claiborne, R. Paul Ross, Don Ward, Derek Parsonage, and Edward J. Crane, III Department of Biochemistry, Hake Forest University Medical Center, Winston-Salem, North Carolina 27157

Introduction The enteric facultative anaerobe Streptococcus (now Enterococcus) faecalis is easily distinguished from other Gram-positive bacteria such as Bacillus and Staphylococcus in its catalase-negative phenotype, which is a consequence of its biochemical deficit in heme biosynthesis (1). S. faecalis also lacks respiratory cytochromes, and its strictly fermentative metabolism compares more favorably with that of Gram-positive obligate anaerobes such as Clostridium.

Still, S. faecalis enjoys a

robust aerobic lifestyle; in fact, early characterization of its potent pyruvate oxidase activity led to the discovery of lipoic acid (2). Studies by Gunsalus and coworkers (3) and by Dolin (4), and by Hoskins et al. (5) showed that streptococcal extracts were able to use molecular oxygen in the reoxidation of NADH, as generated in part by the S. faecalis pyruvate dehydrogenase system and its associated lipoamide dehydrogenase. An unusual FAD-dependent NADH oxidase catalyzes the tetravalent reduction of O2 -» 2H2O, and the FAD-containing NADH peroxidase catalyzes the reduction of H2O2 -» 2 ^ 0 .

While the S. faecalis pyruvate dehydrogenase,

NADH oxidase, and NADH peroxidase were the subjects of numerous reports in the 1950's and 1960's, considerable progress in this laboratory since 1986 has led to a much clearer understanding of their similarities and distinctions relative to the flavoprotein disulfide reductase family. Streptococcal NADH Peroxidase Since the 1990 Flavin Symposium the structural relationship of this unique flavoprotein peroxidase to other members of the disulfide reductase family has been elucidated through the 2.16 A x-ray structure for the enzyme from S. faecalis 10C1 (6).

In general, the NADH peroxidase structure reveals

considerable similarity to those now determined for glutathione reductase (GR), lipoamide dehydrogenase, and other disulfide reductases of the GR class, with respect to chain fold, domain organization, and FAD location.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

588 The high-resolution peroxidase structure places the active-site cysteine (Cys42) in a position superimposable with that of the charge-transfer Cys63 of human GR. In spite of this degree of detailed structural similarity, Cys42 of the peroxidase is stabilized as an unusual redoxactive cysteine-sulfenic acid (Cys-SOH) which cycles catalytically between the sulfenic acid and thiol forms (7), thus representing the first example of such a two-electron redox cycle in an enzyme mechanism. The presence of this unique Cys-SOH redox center and the efficient catalytic reduction of H2O2 provide the major basis for the distinction between the peroxidase and the GR claBS of disulfide reductases (Fig. 1). Another major structural distinction, in addition to the absence of an active-site disulfide in the peroxidase, is the location of the active-site base, HislO, which is derived from the FAD-binding domain that also contains Cys42. In GR the active-site His467' is derived from the C-terminal Interface domain of the complementary subunit. HislO appears to play both a catalytic role in H2O2 reduction and a structural role in stabilizing the peroxidase Cys42-SOH redox center.

Figure 1. Left. active-site structure of lipoamide dehydrogenase from Azotobacter vinelandii. focusing on the redox-active disulfide. Right. active-site structure of NADH peroxidase; CS042 represents the non-native Cys42-sulfonic acid. Recently we reported (8) the extensive characterization of the recombinant NADH peroxidase as purified from Escherichia coli, in addition to determining the steady-state kinetic mechanism and demonstrating that the protein is well-suited for one-dimensional NMR work with labeled amino acids. Site-directed mutagenesis experiments have now shown that replacement of Cys42 with Ser or Ala eliminates the non-flavin redox center and peroxidatic activity, generating a simple flavoprotein which is stoichiometrically reduced with one equivalent of NADH (Fig. 2). Although

589 substitution of HislO with Gin or Ala does not alter the behavior of the peroxidase in NADH titrations, the oxidized form of the HislOGln mutant does exhibit a distinctive long-wavelength spectral feature which indicates that HislO in wild-type enzyme has a strong influence on the electronic interaction between Cys42-SOH and FAD. In addition, the HislOGln mutant is very sensitive to peroxide inactivation (irreversible oxidation of Cys42-SOH), further supporting the role of HislO in stabilizing the sulfenic acid.

Figure 2. Anaerobic NADH titrations of wild-type recombinant NADH peroxidase (left) and of the active-site Cys42Ser mutant (right). In the GR-like disulfide reductases, the two cysteines of the disulfide are separated by four residues, and the charge-transfer cysteine of GR aligns with Cys42 of the peroxidase (9). A major question given the functional distinctions between GR and NADH peroxidase has to do with the selection and stabilization of the Cys42-SOH redox center. We have explored the role played by localized secondary structural elements in this selection by individually replacing residues Ile37-Ser41 with Cys and analyzing the resultant mutants for active-site disulfides involving Cys42 and the new Cys. Two mutant peroxidases, Phe39Cys and Leu40Cys, do form disulfides, but the disulfide form of Phe39Cys does not bind FAD due to the concomitant perturbation of active-site structural elements. The Leu40Cys disulfide form (Cys40:Cys42), however, is quite stable and has been subjected to a variety of kinetic, spectroscopic, and redox analyses. The protein has no peroxidase activity, and NADH titrations lead directly to the FADHj enzyme form (Fig. 3). Intramolecular electron transfer from FADH2 to the active-site disulfide, however, does not occur. We have modeled the Cys40:Cys42 disulfide form of this mutant, based on the 2.16 A structure of the wild-type peroxidase, and subjected the model to energy minimization using the Kollman force field. Formation of the disulfide effectively moves the Cys42-Sy to a new position 5.5 A from the flavinC(4a)-position (Fig. 3). In addition, the redox potential of this

590 disulfide appears to be at least 90 mV lower than that of the FAD, based on dithionite titrations in the presence of a redox indicator dye plus methyl viologen. This combination of unfavorable factors explains the altered redox behavior of the Leu40Cys mutant and provides further support for the conclusion that the redox-active disulfides of GR and other disulfide reductases exist in Bpecial conformations enhanced by activesite secondary structural elements (10). Taken together, our analyses indicate that the exclusion of vicinal cysteine thiols and an active-site secondary structure that will not support introduction of a redox-active disulfide represent two of the major factors contributing to the selection and stabilization of the Cys42-SOH redox center in the streptococcal NADH peroxidase (11).

WAVELENGTH, nm

Figure 3. Left. NADH titration of NADH peroxidase Leu40Cys mutant. Right. active-site superimposition of GR (human erythrocytes) and the NADH peroxidase Leu40Cys disulfide model. Peroxidase residues 40-42 are displayed; the Cys40:Cys42 disulfide is shown as a dashed line. Streptococcal NADH Oxidase:

A Second Flavoprotein Peroxide Reductase

In 1989 (12) we presented evidence for a second non-flavin redox center in the streptococcal NADH oxidase as well; we further demonstrated that one cysteinyl peptide sequence from the oxidase was closely related to the active-site sequence for the peroxidase. Other spectroscopic and redox properties of the oxidase, combined with this active-site sequence homology, supported the conclusion that this flavoprotein also contained a Cys-SOH redox center. In 1992 (13) we reported the cloning, sequence, and overexpression of the NADH oxidase gene from S. faecalis 10C1; the oxidase sequence is 44% identical to that of the NADH peroxidase. Overall, the homology between peroxidase and oxidase extends to all four structural domains, and 11 of the 15 residues within 5 & of the FAD isoalloxazine in

591 the peroxidase are conserved in the oxidase sequence.

Among the most

highly-conserved segments is the cysteinyl peptide containing Cys42, which appears in the identical position as Cys42-SOH in the peroxidase sequence. The active-site base HislO is conserved in an identical position in the NADH oxidase sequence as well.

These and other considerations support the

conclusion that streptococcal NADH peroxidase and NADH oxidase constitute a distinct claBS of flavoprotein peroxide reductases (14), easily contrasted with disulfide reductases such as GR and thioredoxin reductase. The results of an active-site structural comparison of streptococcal NADH peroxidase and NADH oxidase, based on the chemical, spectroscopic, and redox properties of the two proteins reconstituted with artificial flavins, fully supports this classification (15).

One of the principal

differences between the two native proteins is the modulation of the respective FAD redox potentials, with that of the two-electron reduced oxidase being considerably higher than that in the EH2 form of the peroxidase.

This leads to the generation of a catalytically-competent

NADH oxidase EH4 species, which carries out the four-electron reduction of 02

2H 2 0 (16).

This oxygen reactivity has been explored (17) through the

use of artificial flavins such as 1-deaza-, 2-thio, and 4-thio-FAD, which proved to be valuable mechanistic probes with p-hydroxybenzoate hydroxylase.

In general, these results suggest that the detailed

mechanism of oxygen activation employed by NADH oxidase differs from that of the aromatic hydroxylases.

By modeling the C(4a)-peroxyflavin analog

of Schreuder et al. (18) within the active site of the homologous NADH peroxidase, we were also able to demonstrate that the opposite enantiomer would be required for such a hypothetical intermediate in NADH oxidase oxygen reactivity, as compared with both p-hydroxybenzoate hydroxylase and glycolate oxidase.

Evidence for Multiple Lipoamide Dehydrogenases in Streptococcus faecalis In addition to lacking hemeproteins, the streptococci are generally thought to lack most enzymes of the citric acid cycle; the predominant aerobic fate of acetyl-CoA is conversion to acetate, and this process generates one ATP per mole of acetate (19).

A phosphotransacetylase

converts acetyl-CoA to acetyl phosphate, using inorganic phosphate, and ATP is produced through the action of acetate kinase.

Direct evidence for

a lipoic acid-dependent pyruvate dehydrogenase system in S. faecalis 10C1 was first presented by Dolin and Gunsalus (20), who also demonstrated activity with a-ketobutyrate.

More recently, the S. faecalis pyruvate

dehydrogenase has been shown (21) to exhibit unexpectedly high activity in vivo when grown anaerobically on pyruvate, and Snoep et al. (22) purified both the active complex and the associated E3 (lipoamide

592 dehydrogenase) component from S. faecalis NCTC 775 in 1992. Based on an examination of both the NADH inhibition of thiB lipoamide dehydrogenase and its redox behavior in the presence of excess NADH, Snoep et al. (22) concluded that the enhanced in vivo pyruvate dehydrogenase activity in anaerobic S. faecalis was partly due to the relative insensitivity of the E3 component to over-reductxon by NADH and concomxtant xnhxbxtxon. Allen and Perham (23) reported the cloning and sequence of a DNA fragment from S. faecalis (ATCC 29212) which encoded the entire dihydrolipoyl transacetylase (E2) gene and partial upstream and downstream open reading frames corresponding to the pyruvate dehydrogenase Elf) and E3 components, respectively. The S. faecalis lipoamide dehydrogenase IpdhD) gene was thus shown to be located within a gene cluster, as has been found for the pyruvate dehydrogenase components in E. coli (24), Bacillus stearothermophilus (25), and Staphylococcus aureus (26). The S. faecaliB 10C1 lpd Gene Based on multiple sequence alignments of several lipoamide dehydrogenases from bacterial sources, consensus segments corresponding to the redoxactive disulfides and active-site histidines were selected for the design of oligonucleotide primers. A 1.1-kb product was consistently obtained in low yields by PCR amplification, using these primers, from S. faecalis 10C1 DNA. Using this fragment as a probe, two distinct clones containing inserts of 2 kb and 4.2 kb were isolated from a Xgtll genomic library. Restriction mapping and sequence analysis localized the S. faecalis lpd gene to a Pstl-EcoRI fragment at the 3' end of the 4.2-kb pLD03 clone; this 1.7-kb segment contained 300 bp of upstream nucleotide sequence and all but seven of the 3' codons of the lpd open reading frame. On the other hand, the complementary pLD09 clone (2 kb) contained the full-length lpd gene on an overlapping Pstl-BamHI fragment. The entire lpd sequence was completed on both strands by combining complementary data from these two clones. The lpd gene (1.4 kb) encodes a polypeptide of 469 amino acid residues; the protein has been overexpressed in E. coli with the T7 RNA polymerase system and is active in the NADH:1ipoamide assay at pH 8.0. The apparent molecular weight of 51,000 from SDS/PAGE agrees well with the calculated value, based on the deduced sequence. FASTA alignments reveal 33-38% identity with the lipoamide dehydrogenases from E. coli (24), Azotobacter vinelandii (27), S. aureus (26), and B. stearothermophilus (25). Comparisons with the NADH oxidase (13) and NADH peroxidase (9) from S. faecalis 10C1 yield identities of 24-25% as expected. Based on the 2.2 A structure of the A. vinelandii lipoamide dehydrogenase (10), there are no indications of functionally-significant substitutions in the S. faecalis Lpd. One major difference in the streptococcal sequence,

593 however, involves the basic dipeptide His44-Lys45 that replaces the very strongly conserved Asn50-Val51 sequence (28) found between the two cysteines of the redox-active disulfide (Fig. 4). The functional and/or structural consequences of this change are not known.

Npx S. Nox 3. PdhD S. Lpd S. PdhD B. PdhD B. PdhD S. Lpd A. Lpd P. LpdV P. LpdG P. Lpd3 P. Lpd E.

faecalis faecalis faecall faecalis stearot subtlll aureus vlneland flouresc putida putida putida coll

FAD-BINDING •REDOX-ACTiyE,CYStEiNE( MKV IVLGSSHGGY-EAVEELLNLHPDAEIQWYEKGDFISFLS--MGMQLYLEGKV MKV VWGCTHAGT - SAVKSILANHPEAEVTVY ERNDNIS FLS - -f(£GI ALYVGGW MWGDFAIELDTWIGAGPGGYVAAIRAAEMGQKVAIIER EYIGGVCLNVGCIPSKA MAE QTDLLILGGGTGGYVAAIRAAQKGLNVTIVEKYK LGGTCLHKGCIPTKA MWGDFAI ETETLWGAGPGGYVAAIRAAQLGQKVTIVEK-G NLGG\fPIiNVGC: PSKA MWGDFPIETDTLVIGAGPGGYVAAIRAAQLGQKVTWEK-A TLGG^CIitiVGCI PSKA MWGDFPIETDTIVIGAGPGGYVAAIRAAQLGQKVTIVEK-G NLGGVCLNVGCI PSKA MS-QKFDV IVIGAGPGGYVAAIKSAQLGLKTALIEKYKGKEGKTALGGTCLNVGCIPSKA MS-QKFDV WIGAGPGGWAAIRAAQLGLKTACIEKYIGKEGKVALGGTCLNVGOIPSKA M- - -QQTIQTTLLIIGGGPGGYVAAIRAGQLGIPTVLVEG QALGGTCLNIGqi PSKA MT-QKFDV WIGAGPGGYVAAIKAAQLGLKTACIEKYTDAEGKLALGGTCLNVGCI PSKA M--KSYDV VIIGGGPGGYNAAIRAGQLGLTVACVE GRSTLGG®it®i32MPSKA ST EIKTQVWLGAGPAGYSAAFRCADLGLETVIVERYN TLGGVCLNVGGIPSKA

Figure 4. Multiple alignment of FAD-binding domain sequences for bacterial lipoamide dehydrogenases, NADH peroxidase, and NADH oxidase. Comparison of the full-length lpd sequence with the 57-residue N-terminal sequence reported for the S. faecalis (ATCC 29212) pdhD gene product (23) reveals 57% identity, clearly distinguishing the two streptococcal lipoamide dehydrogenases and supporting the suggestion that these enteric streptococci may have at least two lpd and/or pdhD genes. Further evidence for this heterogeneity comes from genomic Southerns and from sequences immediately upstream of the S. faecalis 10C1 lpd gene, which demonstrate that lpd is not linked to any known a-ketoacid dehydrogenase E2 gene. We have now demonstrated (Fig. 5) that the S. faecalis lpd gene is, however, linked with two open reading frames encoding gene products homologous to the phosphotransbutyrylase (29) and butyrate kinase (30) of Clostridium acetobutvlicum (GenBank accession number L04468). Indeed, the initiation codon for the streptococcal lpd gene is separated from the butyrate kinase (buk) termination codon by only five nucleotides, and a probable ribosome binding site seven base pairs upstream from the ATG start codon overlaps with the 3'-terminus of the buk coding region. This aspect of lpd gene organization closely resembles that for the pdhD genes from the Gram-positive S. faecalis (ATCC 29212; Ref. 23), S. aureus (26), and B. stearothermophilus (25); in all four cases there is very tight linkage between the lipoamide dehydrogenase gene and the corresponding upstream gene (pdhc or buk). The S. faecalis lpd gene differs from other lpd and pdhD genes, however, in that no potential rho-independent transcription terminator is evident downstream of the 3'-terminus; in fact

594 a fourth putative ORF initiation codon comes 17 nucleotides after the lpd stop codon. HlndlU

Htn98% D 2 0

using a 600

MHz spectrometer. Flavocytochrome b2 concentrations were 0.1 mM tetramer (i.e. 0.4 mM subunit) and cytochrome c aliquots were titrated into the flavocytochrome b2 sample (both proteins were kept oxidised throughout). The linewidths due to cytochrome c haemmethyl resonances at 34.0 and 32.5 ppm were monitored throughout the titration. If significant binding of cytochrome c to flavocytochrome b2 occurs then the linewidths should broaden significantly whereas if there were no binding at all then the linewidths would be identical to free cytochrome c.

A plot of linewidth at 34.0 ppm against

l/[cytochrome c] for both wild-type and HA3-enzyme is shown in Figure 5.

614 O

200-1

140-

N

X

2

i

80-

c

o 20

0

2 5 1 / [Cytochrome c j

5 0 -1

mM

Figure 5. Plot of the linewidth at 34.0 ppm against 1/[cytochrome c] for the wild-type and HA3 flavocytochromes b2 The degree of binding is indicated by the gradient of the lines; the steeper the gradient the stronger the binding. Thus from Figure 5 it is clear that the wild-type enzyme binds cytochrome c far more strongly than the HA3-mutant enzyme. Thus, at least qualitatively, we can confirm from the NMR studies that the HA3-mutation does disrupt cytochrome c binding to flavocytochrome b2. From our work on the HA3-mutant enzyme we conclude that: the interdomain hinge has little importance in the lactate dehydrogenase function of the enzyme; the hinge plays a central role in mediating interdomain electron transfer from FMN to ¿>2-haem; the hinge is also important for binding and electron transfer to cytochrome c.

Acknowledgements

This work was supported by the Science and Engineering Research Council through research studentships awarded to PW, SD and RES and research grants to GAR and SKC. We are grateful to John Parkinson for help with the nmr experiments and to Florence

615 Lederer, Scott Mathews, Mariella Tegoni and Christian Cambillau for fruitful discussions.

References 1.

Chapman, S.K. and G.A. Reid. 1993. Chem. in Brit. 29:202-204.

2.

Daum, G., P.C. Bohni and G. Schatz. 1982. J. Biol. Chem. 257:13028-13033.

3.

Appleby, C.A. and R.K. Morton. 1954. Nature 173:149-152.

4.

Xia, Z.-X. and F.S. Mathews. 1990. J. Mol. Biol. 272:837-863.

5.

Jacq, C. and F. Lederer. 1974. Eur. J. Biochem. 47:311-320.

6.

Guiard, B. 1985. EMBO. 1. 4:3265-3272.

7.

Black, M.T., S.A. White, G.A. Reid and S.K. Chapman. 1989. Biochem. J. 258:255-259 .

8.

Pallister, R.L., G.A. Reid, C.E. Brunt, C.S. Miles and S.K. Chapman. 1990. In Flavins and Flavoproteins (B. Curti, S. Ronchi, G. Zanetti eds.), Walter de Gruyter, Berlin, pp. 787-790.

9.

Brunt C.E., M.C. Cox, A.G.P. Thurgood, G.R. Moore, G.A. Reid and S.K. Chapman. 1992. Biochem. J. 285:87-90.

10. Dubois, J., S.K. Chapman, F.S. Mathews, G.A. Reid and F. Lederer. 1990. Biochemistry 29:6393-6400. 11. Miles, C.S., N. Rouvifcre-Fourmy, F. Lederer, F.S. Mathews, G.A. Reid, M.T. Black and S.K. Chapman. 1992. Biochem.J. 285:187-192. 12. White, S.A., M.T. Black, G.A. Reid and S.K. Chapman, 1989. Biochem. J. 263:849-853. 13. White, P., F.D.C. Manson, C.E. Brunt, S.K. Chapman and G.A. Reid. 1993. Biochem. J. 297:89-94. 14. Miles, C.S., F.D.C. Manson, G.A. Reid and S.K. Chapman. 1993, Biochim. Biophys. Acta, (in press). 15. Ogura, Y. and T. Nakamura. 1966. J. Biochem. 60:77-86. 16. Capeillfcre-Blandin, C., M. Iwatsubo, G. Testylier and F. Labeyrie. 1980. In: Flavins and Flavoproteins (K. Yagi and T. Yamamoto, eds.) Japan Scientific Societies Press, Tokyo, pp. 617-630.

The L-Mandelate Dehydrogenase from Rhodotorula gratninis is a Flavocytochrome b2.

Stephen K. Chapman, Otto Smekal Department of Chemistry, University of Edinburgh, West Mains Road, Edinburgh EH9 3JJ, Scotland, UK. Graeme A. Reid Institute of Cell and Molecular Biology, University of Edinburgh, Edinburgh EH9 3JR, Scotland UK

Introduction The L-(+)-mandelate dehydrogenase (L-MDH) from the yeast Rhodotorula graminis is a flavocytochrome which catalyses the oxidation of L-mandelate to phenylglyoxylate (1,2). It has recently been shown that L-MDH shows many similarities to the L-lactate dehydrogenase (L-LDH), or flavocytochrome b2, from the yeasts Saccharomyces cerevisiae and Hansenula anomala. Like L-LDH, the L-MDH from R. graminis is a homotetramer and each subunit contains one FMN and one haem group in a polypeptide of around 59 kDa. The N-terminal sequence exhibits 50% identity with S. cerevisiae L-LDH over 32 residues (2). The apparent similarity between L-MDH and L-LDH raises many interesting questions relating to substrate specificity, particularly since the two enzymes are mutually exclusive towards their primary substrates. Thus the substrate for L-MDH, L-mandelate, is a potent inhibitor of L-LDH whereas L-lactate strongly inhibits L-MDH activity. To understand the molecular basis for this difference in specificity we have investigated the catalytic properties of L-MDH.

Results and Discussion It has been shown previously that proton abstraction at C-2 of lactate is the major ratedetermining step in the reaction catalysed by L-LDH from S. cerevisiae (3-5). Using D,L-

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

618 [2- 2 H]lactate and with ferricyanide as electron acceptor a kinetic isotope effect, KIE, of 5 was measured (3). In contrast, it has been reported that there is no observable 2 H-KIE for the oxidation of mandelate by L-MDH from R. graminis (1). This has been confirmed by a recent study in which the oxidation of D,L-[2- 2 H]mandelate by L-MDH gave rise to a KIE value of 1.1 ± 0.1 which was invariant over the temperature range 5-35°C (6). Thus we can completely rule out the possibility that abstraction of the C-2 hydrogen of mandelate makes a significant contribution to the rate-determining step of the reaction. This conclusion raised questions about the nature of the highest energy transition state in the reaction catalysed by L-MDH. To investigate this we have carried out a detailed kinetic analysis of the oxidation of eight different monosubstituted D,L-mandelates catalysed by L-MDH. These were: 4-chloro-; 4bromo-; 4-fluoro-; 4-methyl-; 3-methoxy-; 4-methoxy-; 3-hydroxy-; and 4-hydroxy-D,Lmandelates. Values of K m and kCit (at 25°C in Tris/HCl buffer, pH 7.5, I = 0.10 M and with ferricyanide, 1 mM, as electron acceptor) were determined for D,L-mandelate and all eight analogues and revealed that D,L-4-hydroxymandelate was the best substrate with a ifccat value of 146 s 1 and a Km value of 0.08 mM. These values compare to 94 s 1 and 0.36 mM found with D,L-mandelate itself. Activation parameters, A H i and AS+ (determined over the range 5-37°C) for all the D,L-mandelate analogues were compensatory, resulting in similar values for the free energy of activation, AG±, of around 60 kJmoH at 298.15 K. If a given reaction series follows a common mechanism then one would expect a quantitative relationship between AH+ and AS± - the so called isokinetic relationship or compensation law (7). Thus the near constancy of A G i values for the oxidation of all the substituted mandelates by L-MDH is strongly indicative that the same mechanism applies in all cases. To provide further mechanistic information we analysed the oxidation of the various D,Lmandelate analogues using a Hammett type approach. The Hammett treatment aims to correlate the effect of meta- and para- ring substituents on reaction rate. Attempts to correlate &cat values with the classical Hammett relation, equation 1, gave reasonable results. log(* cat ) R = log(* cat ) H + op

[1]

Where (&cat)R = kcat for substituted D,L-mandelate R; (&cat)H = kCit for D,L-mandelate itself; a = the sum of the total electronic effects of the ring substituent; and p = the

619 reaction constant (reactions with a positive p are helped by electron withdrawing groups and vice versa). Fits to equation 1 gave a relatively low value of p of around 0.5 indicating that the ring substituents have only a small demand on the electron density at the C-2 carbon in the transition state. This would imply a fairly electron rich C-2 carbon in the transition state. To gain insight into the separate contributions of the inductive and resonance effects of the ring substituents we analysed the data from all the substituted D,L-mandelates in terms of Taft's dual substituent parameters equation, equation 2 (8). l ° g ( * c a t ) R = !°g(*cat)H + P l a I +

Where

X o

R+]

t2l

= the value for the inductive effect and 95% homogeneous. MDH-GOX was purified from the soluble fraction of E. coli JM105. Following ammonium sulfate fractionation, the soluble chimeric protein was purified by chromatographies on DEAE-Sephacel and Phenyl Superose. FMN was included in all buffers to prevent the loss of cofactor. By SDS-PAGE, the purified MDH-GOX is > 98% homogeneous.

624 Kinetic properties of wild type MDH and MDH-GOX Both wild type MDH and MDH-GOX are specific for the S-enantiomer of mandelate and also catalyze the oxidation of S-phenyllactate. MDH-GOX does not produce any H2O2. The kinetic parameters of wild type MDH and MDH-GOX are summarized in Table II. Table II. Kinetic parameters for purified preparations of MDH and MDH-GOX. wild type MDH S-[^H]-mandelate 2

S-[ H]-mandelate R,S-[^H]-phenyllactate 2

R,S-[ H]-phenyllactate

kcat(sec~*)

Km (mM)

220 ± 4

0.18 + 0.02

MDH-GOX kcat (sec~^)

Km(mM)

1.6 + 0.01

0.18 ± 0 . 0 1

103 ± 1 . 5

0.18 ±0.01

0.35 ± 0 . 0 1

0.28 ± 0.02

0.33 ± 0.01

3.2 ±0.23

0.82 ± 0 . 0 1 '

1.02 ± 0 . 0 6

0.07 ± 0.01

5.8 ± 1 . 8

0.15 ± 0 . 0 1

0.87 ± 0.08

Small a-hydroxyacids, including glycolate, lactate, and a-hydroxybutyrate are neither substrates nor inhibitors for both enzymes. R-Mandelate, phenylacetate, and phenyl-1,2ethanediol are weak inhibitors of both enzymes. Preliminary stopped flow experiments indicate that the lower activity of MDH-GOX is the result of a slower rate of reduction of FMN by bound substrate.

References 1.

Tsou, A. Y„ S. C. Ransom, J. A. Gerlt, D. D. Buechter, P. C. Babbitt, G. L. Kenyon. 1990. Biochemistry 29: 9856.

2.

Diep Le, K. H„ F. Lederer. 1991. J. Biol. Chem. 266: 20877.

3.

Lindqvist, Y. 1989. J. Mol. Biol. 209: 151.

4.

Xia, Z„ F. S. Mathews. 1990. J. Mol. Biol. 212: 837.

5.

Lindqvist, Y„ C.-I. Branden, F. S. Mathews, F. Lederer. 1991. J. Biol. Chem. 266: 3198.

6.

Volokita, M., C. R. Somerville. 1987. J. Biol. Chem. 262: 15825.

7.

Hoey, M. E„ N. Allison, A. J. Scott, A. J., C. A. Fewson. 1987. Biochem. J. 248: 871.

The

D282N

and

Y254L

active-site

mutants

of

expressed in E. coli as a mixture of holoenzyme and

flavocytochrome flavin-free

b2 are

protein

Muriel Gondry*, Myriam Schaffner*, Forbes D.C. M a n s o i A Stephen K. Chapman*, Graeme A. & Florence Lederer* *CNRS URA 1461, Hôpital Necker, 75743 Paris Cedex 15, France Departments of ''Chemistry and ^Microbiology, University of Edinburgh, Edinburgh EH9 3JG, UK

Introduction

Flavocytochrome

¿2 from Saccharomyces

cerevisiae

(L(+) lactate cytochrome c

oxidoreductase) is a tetramer of identical subunits with M r 57 000. Each subunit consists of two domains, one containing FMN, the other a protoheme IX group. The enzyme catalyses the oxidation of L-lactate to pyruvate : L-lactate reduces FMN, reducing equivalents then pass sequentially to the ¿>5-type cytochrome which is the electron donor to cytochrome c. The three-dimensional structure of the protein has been solved to 2.4 A (1). Studies in solution have led to propose a chemical mechanism with formation of a carbanion intermediate as the initial step and to assign specific roles to the active-site side chains (2). The proposed mechanism has been further refined by the study of site-directed mutants. In wild-type enzyme, the catalytic base H373 is predicted to hydrogen bond to D282. After a proton abstraction by H373, the incipient imidazolium ion would be stabilized by an electrostatic interaction with the carboxylate of D282. The mutant protein D282N is being studied in order to define the importance of these interactions. The mutant protein Y254L is also being characterized. The previously published study of the Y254F mutant (3) showed that the hydrogen bond between the phenolic -OH and the substrate stabilized the transition state. Before carrying out a kinetic characterization for the D282N and Y254L proteins, we had to solve a new problem, namely that flavin insertion is affected during mutant biosynthesis in E. coli.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

626 Results and Discussion Flavin Insertion is Affected during Biosynthesis in E. coli. The purification of the D282N and Y254L mutants cannot be carried out as for that of the wild-type: their chromatographic behaviour is particular i.e. they are strongly bound to the hydroxyapatite gel and elution is very difficult. The problem is solved by applying a steep linear phosphate gradient instead of the usual ammonium sulfate gradient (as an example: 5.57 (imol of D282N mutant are loaded onto a column (2.5 x 20 cm) and eluted with a gradient from 350 ml 0.1 M Na+/K+ phosphate buffer pH7 to 350 ml 0.7 M Na+/K+ phosphate buffer pH7). If one pools all the heme-containing fractions, flavin assays by fluorescence under denaturing conditions indicate a flavin-to-heme ratio of less than unity. Indeed, the elution profiles from the hydroxyapatite column for the two mutant proteins show a shift beween the heme elution and the specific activity profiles which is not observed with the wild-type enzyme. This shift results from a flavin deficit compared to heme in the early part of the peak. Already before chromatography, the D282N and Y254L mutants have a flavin-to-heme ratio of about 0.6. There is neither a significant decrease in the total activity nor an important drop in the flavin-to-heme ratio between any two successive steps of purification. Therefore, the partial lack of flavin appears to already exist in the cell. Wild-type flavocytochrome ¿2 aid the previously studied mutants (Y143F, Y254F) produced by the same E. coli MM294 strain present a 1/1 FMN-to-heme stoichiometry. Moreover, since the mutants D282N and Y254L are not expressed at a higher level than the wild-type, it appears that the FMN synthesizing capacity of the cell should be sufficient. We propose that these last two mutations alter the pathway or the rate of intracellular folding in such a way that flavin insertion is affected during biosynthesis. A blue ultrogel A4R column can be used to separate the weakly adsorbed D282N holoprotein from the flavin-free enzyme eluted at high ionic strength (Fig. 1). The same separation is obtained under the same conditions for the Y254L mutant. The flavin-free fractions cannot be displaced specifically by about 10 |iM FMN, contrary to the deflavoenzyme prepared from wild-type holoenzyme by rapid Sephadex filtration at acid pH (4). Flavin addition to the supernatant immediately after cell disruption does not increase the specific activity. These results show that once folded the mutant flavin-free enzymes present a local structural perturbation which prevents flavin binding. But selective proteolysis experiments as well as the behaviour of the mutants on hydroxyapatite suggest that the flavodehydrogenase domain is still essentially folded into an as/Ps-barrel; moreover, the visible spectra of the D282N and Y254L mutants lead us to believe that the heme-binding domains are correctly folded.

627 Figure 1 : A Blue Ultrogel A 4 R Column Allows to Separate the Holoenzyme from the Flavinfree Enzyme.

fraction number

1.25 (imol D 2 8 2 N mutant in 0.1 M N a + / K + phosphate buffer pH7 is diluted two-fold with water before loading onto the column (2.5 x 15 cm) equilibrated in 0 . 0 5 M N a + / K + phosphate buffer pH7. The elution is carried out with 0.1 M N a + / K + phosphate buffer, lactate 10 mM, pH7.

Preliminary Kinetic Characterization of the Mutants (Table 1). For the D 2 8 2 N mutant, the fact that the K m value is not significantly altered would seem to imply that, in agreement with the proposed mechanism (2), this residue is not involved in Michaelis complex formation and that the active-site is not severely disrupted. Deuterium isotope effect measurements (results not presented) showed that the a-proton abstraction is still the main rate-limiting step in overall transfer from lactate to ferricyanide, as described for the wild-type enzyme. The 70-fold lowering o f k c a t leads to the conclusion that it is this proton abstraction step that is affected by the mutation. Although a hydrogen bond is still possible, the mutation removes the D282-H373 electrostatic interaction which stabilizes the imidazolium ion formed during a-hydrogen abstraction; with the D 2 8 2 N mutation, the transition state is more difficult to reach (AAG^ = 2.80 kcal/mol). The previously published study of the Y 2 5 4 F mutant (3) showed that the major role of Y 2 5 4 in transition state stabilization (AAG* = 2.08 kcal/mol) is to freeze, with a hydrogen bond, the substrate alcohol function in a conformation such that the a-hydrogen is well oriented for abstraction by H 3 7 3 . The Phe residue could still have an orientation effect thanks to a weak electrostatic interaction between the substrate hydroxyl function and the edge o f the phenylalanine ring. This effect is lost for the Y 2 5 4 L mutant: the transition state is even more

628 Table 1: Steady-State Kinetic Parameters for the Mutant Proteins Y254L and D282N.

Enzyme

K m lactate (mM)

kcat (s- 1 )

kcat/Km (10 3 M ' V 1 )

AAG*[mut-WT] (kcal/mol)

Wild-type (5)

0.49 ± 0 . 1

270 ± 30

551.0

-

Y254F (3)

0.35 ± 0.07

6.1 ± 0 . 2 5

17.4

2.08

Y254L (this work)

0.39 ± 0.04

0.51 ± 0.02

1.31

3.64

D282N (this work)

0.73 ± 0.05

3.9 ± 0.1

5.34

2.80

The k c a t values are expressed as mol of substrate reduced s"1 (mol of subunit)" 1 , with 1 mM ferricyanide as monoelectronic acceptor. The determinations were carried out in 0.1 M phosphate buffer, EDTA 1 mM, pH7 at 30°C.

destabilized (AAG* = 3.64 kcal/mol). The absence of K m change in the Y254F and Y254L mutants seems incompatible with the postulated existence of a hydrogen bond between the substrate hydroxyl function and the Y254 phenol group in the Michaelis complex. For the Y254F mutant, an energetic compensation was postulated resulting from the weak electrostatic interaction with the Phe254 ring. This suggestion is ruled out by our studies with the Y254L mutant. Yet there remains a doubt concerning the existence or not of the hydrogen bond, since the symmetrical removal of the substrate hydroxyl group, by using propionate as an inhibitor for the wild-type enzyme, yields a Kj of 28 mM (6).

References 1.

Xia, W.X., F.S.Mathews. 1990. J. Mol. Biol. 212, 837-863

2.

Lederer, F. 1992. Protein Science 1, 540-548

3.

Dubois, J., S.K.Chapman, F.S. Mathews, G.A. Reid, F. Lederer. 1990. Biochemistry 29, 6393-6400

4.

Pompon, D„ F. Lederer. 1978. Eur. J. Biochem. 90, 563-569

5.

Lederer, F. 1991. In : Chemistry and Biochemistry of Flavoenzymes, Vol. 2 (F. Miiller; ed) CRC Press; Boca Raton, Florida, pp. 153-242

6.

Genet, R„ F. Lederer. 1990. Biochem. J. 266, 301-304

RECONSTITUTION OF FLAVIN-FREE FLAVOCYTOCHROME ¿>2 WITH 5-DEAZAFMN : A CARBANION OR A HYDRIDE MECHANISM ?

Alexis Balme and Florence Lederer CNRS URA 1461, Hôpital Necker, 75743 Paris Cedex 15

Introduction Flavocytochrome b2 (from Saccharomyces

cerevisiae catalyses the 2-electron oxidation of

L(+)-lactate to pyruvate. Its crystal structure was solved to 2.4 A resolution ( 1 ) and a catalytic role was proposed for active site side chains (2)(Fig.l). In the putative Michaelis complex, lactate interacts with Arg376, Tyrl43 and Tyr254. His373, acting as a general base, abstracts the lactate a-hydrogen to form a carbanion intermediate. This is the main rate-limiting step of the reaction. Electrons are then transferred to the FMN, either after intermediate formation of a covalent adduct between the carbanion and the cofactor, or through two monoelectronic tranfer steps in quick succession. In the normal catalytic cycle, reduced flavin yields its reducing equivalents one by one to heme b2, which in turn reduces cytochrome c. Nevertheless the flavin can also function as a transhydrogenase, by-passing the heme after the first catalytic cycle (3). The study of intermolecular deuterium and tritium transfer under transhydrogenation conditions provided the best evidence in favour of a carbanion mechanism. When [2-2h]- or [2-3H]lactate and a halogenoketoacid are incubated together with the enzyme, the isotope is in part transferred to the C2 position of the ketoacid substrate and in part lost to the solvent (Fig.2). When the halogenoketoacid is chloro- or bromopyruvate, halogen elimination can also be observed and the isotope is found in pyruvate. It was shown that the extent of tritium loss to the solvent decreases with increasing ketoacid substrate concentration. At infinite concentration, intermolecular transfer is total (4). It is believed that only the active site base, His373, is involved in intermolecular proton transfer and exchange with the solvent (5).

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

630

Figure 1: Proposed catalytic role for side chains. A: Michaelis complex. H i s 3 7 3 acts as a general base and abstracts the a-proton. B: Carbanion intermediate. Electrons are transferred one way or the other to F M N . This movement is facilitated by the positive charge o f L y s 3 4 9 . C: Reduced enzyme-product complex. *H

H C H 2 - C— CO2" II O F i g u r e 2: Intermolecular isotope transfer during a f l a v o c y t o c h r o m e b 2 - c a t a l y s e d transhydrogenation reaction between isotopically labelled lactate and a halogenopyruvate (fluoropyruvate, however, does not give rise to the halide elimination reaction).

631 S o m e twenty years ago, 5-carba-5-deazaflavin was used to replace normal flavin as a cofactor in a number of flavoproteins. It turned out to be somewhat disappointing as a mechanistic probe, since it is incapable of monoelectronic transfer and devoid of reactivity with oxygen ; it was dubbed a "flavin-shaped nicotinamide" (6,7). It was f o u n d in particular, with enzymes such as lactate oxidase (8), D-amino acid oxidase (9,10) and f l a v o c y t o c h r o m e b2 (11), that upon reduction of 5-deazaflavoenzyme with a - t r i t i a t e d substrate, the isotope was found at the C5 position of the reduced cofactor analog. Stereospecific back transfer to the ketoacid was also observed. T h e s e results can be rationalized in either of two ways: -either there occurs a direct hydride transfer between subtrate and 5-deazaflavin -or a carbanion mechanism still operates with the active site base transferring the substrate a-proton to the flavin analog C5. It should be noted that for some authors this transfer from the base to the cofactor could also occur with normal flavin (9,10,12), but the active site structure of flavocytochrome ¿>2 does not m a k e this possibility seem likely (5). It appeared to us that the study of the intermolecular tritium transfer reaction under transhydrogenation conditions should provide a way of distinguishing between the two possibilities. Indeed, direct hydrogen transfer by way of a hydride reaction should occur without any isotope loss to the solvent, as is observed with nicotinamide-dependent enzymes. But in the case of a carbanion reaction, one would expect His373 to be able to exchange its proton with solvent as it does in the normal catalytic mechanism.

Results Enzyme resolution and reconstitution The F M N was resolved from flavocytochrome b2 by a modification of a published method (13), using adsorption on PhenylSepharose in 1M ammonium sulfate and eluting flavin at acid p H (G.Fleischmann, to be published). Flavin-free enzyme was mixed with a 2-fold excess of 5-deazaFMN, incubated for 5 min at 30°C and 15 min at 0°C. Starting with a holoenzyme specific activity of 275±25s"' (lactate to ferricyanide transfer), the deflavoenzyme was obtained with an activity of 4s~ 1(1.5%). After reconstitution with 5deazaFMN, the activity rose to 7-8s~l(=2.5%). The amount of reconstitution varied f r o m 63 to 85%, depending on the preparation. The 5-deazacofactor is inactive in the transfer from

632 lactate to ferricyanide but can catalyze a transhydrogenation between L-lactate and halogenoketoacids. Using bromopyruvate the following steady-state parameters were determined: k C at=60s"l (corrected for the amount of reconstitution) and K m =2.7 mM. These figures can be compared to those obtained for the same reaction using native enzyme : kCAT=4s~L and KM=4.9 mM. The difference in kCAT was expected in view of the low E M of enzyme-bound 5-deazaflavin (11), which makes the reduction of ketoacids more favourable than with normal flavin.

Intermolecular tritium transfer Enzyme reconstituted with 5-deazaFMN was incubated in the presence of DL-[2-3H]lactate and different concentrations of fluoropyruvate. For each experiment, tritiated water, residual [^HJlactate and newly formed [^HJfluorolactate (Flac) were separated by ion exchange chromatography (4). The radioactivity of each fraction was determined as well as the specific radioactivity of fluorolactate. The amount of intermolecular transfer was taken to be dpm in Flac dpm in Flac+dpm in H2O . The radioactivity found in H2O had to be corrected first for the radioactivity of the control. Two different types of controls were used in two independent series of experiments. In the first series, the control was a mixture of DL-[2-3H]lactate and deflavoenzyme, so that the correction arose essentially from the small amount of radiolysis of the sample present already before experiment. In the second series, the control contained

DL-[2-^H]lactate,

fluoropyruvate and deflavoenzyme, so that the correction included the amount of tritium loss to solvent occurring during turnover of residual FMN. The results are shown in Fig.3 for the 5-deazaFMN enzyme and can be compared to those obtained in previous work with native enzyme (14), which are reproduced in Fig.4. It can be seen that with 5-deazaflavoenzyme tritium transfer seems to be total even at very low fluoropyruvate concentrations. In direct correlation with this phenomenon, the specific radioactivity of the fluorolactate formed is constant within experimental error, whereas with normal enzyme, as expected, it increases with the amount of intermolecular transfer.

633

5-deazaFMN

o

O •

1

1

5

10

Series 1 Series 2

[fluoropyruvate] (mM)

B

O •

5

T

T

10

15

Series 1 Series 2

20

[fluoropyruvate] (mM) Figure 3: Influence of substrate concentration on the intermolecular tritium transfer catalysed by 5 - d e a z a f l a v o c y t o c h r o m e b2- A: f r a c t i o n of i n t e r m o l e c u l a r transfer. B: specific radioactivity of the fluorolactate obtained using DL-[2-3H|lactate at 2|iCi/|J.mole. Series 1: the control s a m p l e consisted of [2-3H]lactate and d e f l a v o e n z y m e ; Series 2: the control contained in addition the s a m e fluoropyruvate concentration as the sample. In all cases, enzyme and L-lactate concentrations were 19|J.M (on a heme basis) and 30 m M , respectively. Incubations were carried out at 30°C for 90 min.

634

FMN

[fluoropyruvate] (mM)

[fluoropyruvate] (mM) Figure 4: Influence of substrate concentration on the intermolecular tritium transfer catalysed by native flavoenzyme. A: fraction of intermolecular transfer (the same data are found under the f o r m of a d o u b l e reciprocal plot in (14)). B: specific radioactivity of fluorolactate obtained using DL-[2-3H]lactate at 0.25(J.Ci/|imole. T h e control w a s a solution of [2^HJlactate chromatographed under the same conditions as the samples. Enzyme and L-lactate concentrations were 15|iM and 20mM, respectively. Incubations were carried under argon at 30°C for 30 min.

635 Discussion The experiments reported above show that no loss of tritium to the solvent occurs during the transhydrogenation

between

[2-3H]lactate

and f l u o r o p y r u v a t e c a t a l y s e d

by 5-

d e a z a f l a v o c y t o c h r o m e ¿>2 . This result supports the idea that 5-deazaflavin reduction and reoxidation occurs through a hydride mechanism. The alternative, namely proton abstraction by H i s 3 7 3 which would then, shielded f r o m solvent, transfer the proton to the nonexchangeable C5 position of the cofactor analog, is hard to defend. Analysis of the active site topology indicated that the relative orientation of the flavin and the histidine ring was highly unfavourable for the formation of a hydrogen bond between N5 and N£, and hence f o r proton transfer between the two atoms (5). It is hard to believe that the structural difference between the two cofactors is large enough to make proton transfer between 5d e a z a F M N C5 and N£ of histidine more geometrically favourable than between F M N N5 and Ne. O u r results t h e r e f o r e suggest that a mechanistic switch occurred f r o m a carbanion mechanism to a hydride transfer mechanism upon F M N substitution by 5-deazaFMN. This switch must be in large part brought about by the chemical reactivity of the "flavin-shaped nicotinamide". Nevertheless it also requires an adaptation of the substrate binding m o d e as well as of the roles of protein side chains. It has been described before how modeling studies of lactate in the active site of the crystal structure suggested the possibility of hydride transfer to the cofactor (15). T h e two proposed models are shown in Fig.5 and 6. Fig.5 presents a stereoview corresponding to the putative Michaelis complex bound as described in F i g . l A , poised for proton abstraction by His373. A displacement of the substituents at C2 by a rotation of 4 0 degrees around the C 1 - C 2 bond brings the substrate hydroxyl into hydrogen bonding distance of His373 N e (Fig.6). After this rotation the a - h y d r o g e n points directly towards the F M N N5 position and it appears reasonable to postulate a removal of the substrate hydroxyl proton by His373 in concert with hydride transfer to the 5-deazacofactor. T h u s H i s 3 7 3 would play the same role as the active site histidine in lactate and malate d e h y d r o g e n a s e s . This is a remarkable switch in catalytic role coupled to the c o f a c t o r substitution.

636

Figure 5: Stereoview of lactate modeled into the active site for a carbanion mechanism.

Figure 6: Stereoview of lactate modeled into the active site for a hydride transfer mechanism. Only the position of the substrate 0 2 and C 3 differ f r o m those of Fig.5. These positions were obtained by a rotation of 4 0 degrees around the C1-C2 axis.

References 1.

Xia, Z.X. and F.S. Mathews. 1990. J. Mol. Biol. 212,

837-863

2.

L e d e r e r F . and F.S. Mathews. 1987. In : Flavins and Flavoproteins, (D.E. E d m o n d s o n , D.B. McCormick, eds.). Walter de Gruyter, Berlin.pp. 133-142

637 3.

Urban, P., P.M. Alliel and F. Lederer .1983. Eur. J. Biochem. 134, 275-281

4.

Urban, P. and F. Lederer. 1985 J. Biol. Chem. 260, 11115-11122

5.

Lederer, F.. 1992. Protein Science 1, 540-548

6.

Hemmerich, P., V. Massey and H. Fenner.1977. FEBS Lett. 84, 5-21

7.

Walsh, C.. 1980. Acc. Chem. Res. 13, 148-155

8.

Averill.B.A., A. Schonbrunn, R.H. Abeles, L.T. Weinstock, C.C. Cheng, J. Fisher, R. Spencer and C. Walsh. 1975. J. Biol. Chem. 250, 1603-1605

9.

Fischer, J., R. Spencer and C. Walsh. 1976. Biochemistry 15, 1054-1064

10.

Hersh, L.B. and M.S. Jörns. 1975. J. Biol. Chem. 250, 8728-8734

11.

Pompon, D. and F. Lederer. 1979. Eur. J. Biochem. 96, 571-579

12.

Ghisla, S. and V. Massey. 1986. Biochem. J. 239, 1-12

13.

Van Berkel, W.J.H., W.A.M. Van Den Berg and F. Müller. 1988. Eur. J. Biochem. 178, 197-207

14.

Balme, A. and F. Lederer. 1993. Protein Science (in press).

15.

Dubois, J„ S.K. Chapman, F.S. Mathews, G.A. Reid and F. Lederer. 1990. Biochemistry 29, 6393-6400

Physical Studies on Phthalate Dioxygenase Reductase (PDR)

D. Ballou, G. Gassner, L. Wang, C. Batie, D. Gatti, W. R. Dunham, and R.H. Sands Department of Biological Chemistry and Biophysics Research Division, The University of Michigan, Ann Arbor, MI, USA Introduction Most organic oxidation-reduction reactions in biology, such as oxidations of single bonds or conversions of alcohols to aldehydes, occur in two electron steps. However, in respiration and other electron transfer systems, electrons are most frequently transmitted one at a time through the mediation of iron sulfur centers (Fe/S) and cytochromes. The two-electron to one-electron switch necessary for this is nearly always accomplished by flavoproteins that are linked either intramolecularly or intermolecularly to Fe/S centers or to cytochromes. Phthalate dioxygenase reductase (PDR) functions by transferring electrons from NADH to Phthalate Dioxygenase (PDO), an enzyme that catalyzes the dioxygenation of phthalate shown below (1,2). This dioxygenation is the first step in the biodégradation of phthalate and is representative of many bacterial systems that degrade aromatic compounds (3). PDR uses an FMN cofactor to accept the hydride from NADH and it transfers electrons one at a time via its plant type [2Fe-2S] to the oxygenase, thus serving as the 2-electron to 1-electron switch (4). PDO is a protein containing both a Rieske [2Fe-2S] center and a mononuclear iron site (1,2). The latter participates directly in the oxygenation reaction. NADH

PDR is a 36 kDa, monomeric iron-sulfurNAD*

containing flavoprotein (1).

The crystal

structure at 2 A resolution of PDR, a three OH

OH

domain protein, is discussed in this volume

(5,6). The FMN and the [2Fe-2S] center are in separate domains brought closely together at their domain interfaces, with the NADH-binding domain presenting the pyridine cofactor to the flavin. The juxtaposition of the three cofactors is appropriate for the stereochemistry of hydride transfer (pro-R-hydrogen of NADH to re-side of the FMN) and for electron transfer to the [2Fe-2S] center. PDR is now recognized to be a member of the FNR family of proteins, being a fusion of a flavoprotein with a ferredoxin (5,6).

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

640 Previous work on the kinetics of the reduction of PDR by NADH showed that there were several phases in the overall reaction (7). Michaelis complexes with NADH preceded charge-transfer (C-T) complexes characterized by long wavelength absorbance typical of other flavoproteins in their oxidized and reduced states when complexed with pyridine nucleotides (8). Intramolecular electron transfer from the reduced flavin to the [2Fe-2S] center results in the formation of a neutral flavin semiquinone and reduced [2Fe-2S]. The rates of these various processes involved were all within a factor of about four, and their clear distinction was further complicated by having only very small absorbance changes, since both semiquinones and C-T complexes have long wavelength absorbance. We have recently developed new instrumentation and methods of simulation and analysis that better permit the delineation of the complicated processes involved in the reduction by NADH. Results from this work are important for future understanding of the basis of how electron transfer from NADH becomes coupled to the PDO hydroxylation reaction. Moreover, PDR provides a prototype for understanding the mechanisms of [2Fe-2S]-flavoproteins found to participate broadly in NAD(P)H to one-electron exchange interchange reactions in biology. Results and Discussion General Properties The spectrum of PDR (Fig. IB, 0 sec spectrum) shows features consistent with those expected for the oxidized form of a [2Fe-2S] containing flavoprotein. The redox potentials for PDR have been determined by the use of dyes as indicators and either the xanthine oxidase system described by Massey (9) or dithionite as the reductant. The potentials both for reducing the flavin to the semiquinone and for reducing the [2Fe-2S] center are -174 mV at pH 7. This is a very high redox potential for a [2Fe-2S] center. The possible reasons for the unusually high potential of the [2Fe-2S] are discussed in the paper by Correll et al (5). The potential for reducing the 2-electron reduced enzyme to the 3-electron reduced state is -287 mV. Thus the first two electrons added result primarily in formation of a species of PDR containing SQ and reduced [2Fe-2S]. The third electron reduced the SQ to FNMH". Recently we have been able to produce a proteolytically clipped PDR that has lost its ferredoxin domain (it is about 8 kDa smaller). As described below, this behaves nearly identically to the holoenzyme in its reduction by NADH, except that it stops at the 2-electron reduced state. The potential for reduction of this apoenzyme by two electrons is -170 mV. The difference between this potential and the third electron potential of PDR is significant,

641 and indicates loss of interactions between the flavin and [2Fe-2S] cofactors and/or some other major environmental changes to the flavin due to removal of the ferredoxin domain.

Reduction by NADH The reduction of PDR by NADH is a hydride transfer and therefore occurs in two-electron steps. Figure 1 shows the reduction with a two-fold excess of NADH on two different time scales. In A the data were recorded in the stopped flow instrument using a diode array detector (4 ms/spectrum). In B the data were recorded with the Quick Scan feature of the HiTech/Kinetic Instruments stopped flow spectrophotometer (2.5 sec/spectrum). Figure 1 : Spectra of PDR reacting with excess NADH (pH = 8.0, T = 4 °C). (A) 4 ms diode array spectra recorded during the first 120 ms of the reaction between 20 |iM PDR and 100 |iM NADH. (B) Quick scan spectra recorded during the reaction of 20 |iM PDR with 50 H-M NADH. The times corresponding to the spectra are: 0 sec (A)1.3 s (o), 34 s, 56 s, 2 min, 7 min (•). A shows selected spectra of the events occurring in the first 120 msec of the reaction: the initial binding

of

NADH

to

form

Michaelis complexes, the hydride transfer to reduce FMN

to

FMNH", and the intramolecular electron transfer resulting in a reduced [2Fe-2S] center and the neutral blue semiquinone of the flavin (SQ). 350

400

450

500

550

600

650

700

750

There

are

three

phases readily

Wavelength (nm)

apparent in this data out to 120 msec and they can be discerned at various wavelengths in the visible spectra (Fig. 2). Blue flavin semiquinones (SQ) (see Fig IB) as well as reduced

642 flavin-NAD C-T complexes have long wavelength absorbance characteristics. However, the SQ absorbance does not extend significantly beyond 700 nm. Thus at 740 nm only the C-T complexes contribute substantially to the data. At 610 nm both SQ and C-T complexes contribute to the spectrum. The data recorded at wavelengths shorter than 500 nm (including 462 nm) have contributions from all species, with the oxidized forms of each species having higher absorbance. Thus data recorded at 462 nm, 610 nm, and 740 nm (Fig. 2) are useful for studying the dependence of the reactions on NADH and NAD, as well as for determining kinetic isotope effects. MC-1 -.-»MC-2

MC 2 >CT-

CT-—>SO

Time

B

0.09

Figure 3: (Above) Experimental data and simulations at 740 nm of 20 PDR reacting with 250 M.M NADH or 250 NADD (pH = 8.0, T = 4 OQ.

/

0.08

(sec)

0.07

Figure 2: (Left) Experimental data and simulations at 462 nm (A), 610 nm (B), and 740 nm (C) of 20 |iM PDR reacting with various concentrations of NADH (o: 100 ^M NADH; • : 20 ^M NADH; x: 10 H.M NADH) (pH = 8.0, T = 4 °C).

0.06 OOS

A study of the dependence of the reaction on NADH concentration shows that the first phase (seen as a lag in the data at 740 nm) is saturable, suggesting that another phase preceding the lag must be the 0

0.01

0.1 Time (S)

1

actual NADH binding step. Thus we propose that binding leads to a Michaelis complex, termed MC-1

(inferred from the kinetics), which is followed by formation of a second Michaelis Complex (MC-2) (Scheme 1). The increase in absorbance shown in Fig 2C (reaching a maximum at ça. 30 msec) is due to formation of CT* by hydride transfer; CT* is assigned to be a complex

643 of NAD with FMNH", with the [2Fe-2S] remaining oxidized. The subsequent loss of absorbance at 740 nm and increase at 610 nm is due to dissociation of NAD and intramolecular electron transfer to yield SQ, which is FMNH° with reduced [2Fe-2S]. Fig. 3 shows the effects of using pro-R-NADD in place of NADH. The deuterium isotope effect occurs on the hydride transfer and decreases the rate of formation of CT* by 7-fold (from 70 sec -1 to 10 sec -1 ). Thus the formation rate is changed from double that of the decay rate (35 sec -1 ) to 1/3 of the decay rate. The slowing of the formation rate of CT* without a change in the decay rate decreases the amount of CT* observed. Since the formation rate constant is actually smaller than the decay constant, the observed ti/2 values cannot be directly associated with the true rate constants (10).

E

•FMN„

PDR„

•Fe/S„

Using the model of Scheme 1 and a single set Kd = 50 |iM

NADH

of rate constants, all of the data of Figs. 2 and 3 could be fit as shown. The down phase at 740

E

-NADH FMN„ •Fe/S_

MC-1

nm is due to the intramolecular electron transfer to form the SQ and the reduced [2Fe-2S] with concommitant loss of NAD. The spectrum at

I I

I I

•NADH

* •FMN„ Fl F
kD= 10 ± 2 s"1

•NAD* N •EMNH^, H F 1 |iM). Unter these conditions 4-nitrobenzoate was reduced to 4-aminobenzoate. Irreversible inactivation was observed with 6 jxM nitrite and 30 |iM hydroxylamine. The radical scavanger hydroxyurea (5 mM), however, did not inactivate the enzymes. It has been shown that ATP is hydrolysed to ADP and Pj during "activation". No phosphorylation or adenylation of either the enzymes or the enzyme-bound riboflavin was observed (10). A two-step ionic mechanism, which considers the presence of iron-sulfur clusters and reduced flavin in the enzyme, proposes in the first step the replacement of the hydroxyl group of the substrate by the reduced flavin in an Sjsj2 reaction to yield enzyme-bound glutaryl-CoA. Thereby the nucleophile could be either the hydride or carbon 4a of the reduced flavin. In the second step, dehydrogenation by the now oxidized flavin leads to the product glutaconyl-CoA and regenerates the reduced flavin (Fig. 2) (4,10). The Sjs[2 displacement of the hydroxyl group by a hydride could be faciliated by the electron withdrawing thiolester and probably by binding the hydroxyl group to a specific iron of the iron-sulfur cluster which could act as a Lewis acid as observed in aconitase (11). The reduction of dimethyl 2-tosyl-hydroxyglutarate to dimethyl glutarate by sodium cyanoborohydride under inversion of configuration may serve as a chemical model (12). Hence, binding of a hydroxyl group to an iron-sulfur cluster might convert the hydroxyl group into a similar good leaving group as does tosylation. The second step of the overall dehydration is a reaction catalysed by the well known acyl-CoA dehydrogenases in which the unactivated hydrogen at C-3 is removed by the flavin as a hydride rather than as a proton. However, the dehydratase did not catalyse the oxidation of added glutaryl-CoA to glutaconyl-CoA by ferricenium ion, an artificial electron acceptor. In summary this very hypothetical mechanism should be regarded as working hypothesis in order to design further experiments. Moreover, the mechanism does not explain the activation of the enzyme by ATP, Mg4"1" and Ti(III)citrate which could lead to a decrease of the redox potential of the reduced riboflavin, i.e. an increase of its reductive power.

Acknowledgement This work was supported by grants from the Deutsche Forschungsgemeunschaft Fonds der Chemischen

Industrie.

and the

662 References 1. Buckel, W. 1992. FEMS Microbiol. Reviews 88: 211-232. 2. Kuchta, R.D., R.H. Abeles. 1985. J. Biol. Chem. 260: 13181-13189. 3. Brunelle, S.L., R.H. Abeles. 1993. Bioorganic Chemistry 21: 118-126. 4. Hofmeister, A., W. Buckel. 1992. Eur. J. Biochem. 206: 547-552. 5. Schweiger, G., R. Dutscho, W. Buckel. 1987. Eur. J. Biochem. 169: 441-448. 6. Dutscho, R., G. Wohlfahrt, P. Buckel, W. Buckel. 1989. Eur. J. Biochem. 181: 741-746. 7. Bendrat, K., W. Buckel. 1993. Eur. J. Biochem. 211: 697-702 8. Buckel, W., U. Dorn, R. Semmler. 1981. Eur. J. Biochem. 118: 315-321. 9. Bendrat, K., U. Müller, A.-G. Klees, W. Buckel. 1993. FEBS Lett, in press. 10. Klees, A.-G., D. Under, W. Buckel. 1992. Arch. Microbiol. 158: 294-301. 11. Beinert, H., M.C. Kennedy. 1989. Eur. J. Biochem. 186: 5-15. 12. Whitman, C.P., G. Hajipour, R.J. Watson, W.H. Johnson, Jr., M.E. Benbenek, N.J. Stolowich. 1992. J. Am. Chem. Soc. 114: 10104-10110.

H+

H,0

O

H COO"

CoAS

H

O CoAS

COO"

Fig. 2. Hypothetical m e c h a n i s m for the dehydration of (R)-2-hydroxyglutaryl-CoA to glutaconyl-CoA. All reactions a r e reversible.

Flavin and iron-sulfur containing hydroxyacyl-CoA dehydratases (II) 4-Hydroxybutyryl-CoA dehydratase from Clostridium

aminobutyricum

Uwe Scherf and Wolfgang Buckel Laboratorium fur Mikrobiologie, Fachbereich Biologie, Philipps-Universitat, D-35032 Marburg, Germany

Introduction The strictly anaerobic bacterium Clostridium aminobutyricum

ferments y-aminobutyrate to

acetate and butyrate (1). The key step in this pathway is the conversion of 4-hydroxybutyrate to crotonyl-CoA (Fig. 1), a reaction in which one of the two hydrogens at carbon-3 of 4-hydroxybutyrate is stereospecifically eliminated. The requirement for acetyl-CoA in this reaction suggested the formation of 4-hydroxybutyryl-CoA as a prerequisite to dehydration to vinylacetyl-CoA and isomerisation to crotonyl-CoA (2). This dehydration is of considerable mechanistic interest since the C-H bond to be cleaved is not activated by the thiolester.

H Fig.1. Reactions and assay of 4-hydroxybutyryl-CoA dehydratase

CoASAc AcO' O

H

O

0

OH

crotonyl-CoA

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

664 Results In order to elucidate the mechanism of the dehydration of 4-hydroxybutyryl-CoA this substrate had to be prepared. It was achieved by purification of a CoA-transferase from C. aminobutyricum which readily transfered the coenzyme from acetyl-CoA to 4-hydroxybutyrate or vinylacetate but not to crotonate (Fig. 1). The oxygen-insensitive enzyme is a homodimer with a molecular mass of 110 kDa (3). Next was the purification of an oxygen-sensitive 4-hydroxybutyryl-CoA dehydratase under strict anaerobic conditions (4). The enzyme representing approximately 15-25% of the soluble protein of C. aminobutyricum

was

obtained in an almost homogenous form by applying a single chromatography on DEAE-Sepharose. However, removal of the last traces of impurity required further ammonium sulfate fractionation as well as chromatography on phenyl-Sepharose and Superdex 200. The enzymic activity was measured in an assay in which 4-hydroxybutyryl-CoA was generated from 4-hydroxybutyrate and acetyl-CoA catalysed by purified 4-hydroxybutyrate CoA-transferase. The formation of crotonyl-CoA was measured by consecutive hydration and NAD-dependent P-oxidation (Fig. 1). The native dehydratase has an apparent molecular mass of 232 kDa and is composed of 4 apparently identical subunits (m = 56 kDa). Under anaerobic conditions the active enzyme revealed a brown colour and contained 2 + 0.2 mol FAD (64 + 5% oxidized), 16 + 0.8 mol Fe and 14.4 + 1.2 mol inorganic sulfur which probably form iron-sulfur clusters. The redox state of the FAD was determined by UV/visible spectroscopy of the supernatant obtained after heat denaturation of the enzyme under anaerobic conditions. During the whole purification procedure vinylacetyl-CoA A3-A2 -isomerase activity eluted with 4-hydroxybutyryl-CoA dehydratase. Moreover the ratio of the activities of isomerase and dehydratase as well as the purification factor of both enzymes remained constant. During conversion of either 4-hydroxybutyryl-CoA or crotonyl-CoA to an equilibrium mixture of both, K' e q = [crotonyl-CoA]/[4-hydroxybutyryl-CoA] = 4.2, no transient vinylacetyl-CoA (3butenoyl-CoA) was formed. Furthermore, starting with vinylacetyl-CoA, which was completely consumed, the same equlibrium mixture was obtained. The results suggest that enzymebound but not free vinylacetyl-CoA may be an intermediate of the dehydration (Fig. 1). Upon exposure to air both activities, the dehydratase as well as the isomerase, transiently increased by 20-40% within the first min followed by complete irreversible inactivation yielding a yellow protein within one hour. The initial activation was accompanied by oxidation of the partially reduced FAD present in the enzyme as shown by UV/visible spectroscopy. The irreversible inactivation may be due to oxidation of the iron-sulfur clusters. Attempts to reactivate the enzyme with ferrous ion under anaerobic conditions were unsuccessful. Reduction of the

665 active enzyme with dithionite yielded an inactive enzyme which was reactivated upon stoichiometric reoxidation with ferricyanide. During this cycle, the FAD of the enzyme was reduced and reoxidized again. Hence, the activity of the enzyme is directly dependent on its content of oxidized FAD. Cyclopropylcarboxyl-CoA, which might be derived by cyclisation of 4-hydroxybutyryl-CoA, was prepared by incubation of cyclopropane carboxylic acid with acetyl-CoA and 4-hydroxybutyrate CoA-transferase. However, the dehydratase did not catalyse its cleavage to crotonyl-CoA. Unlike the related 2-hydroxyacyl-CoA dehydratases, which contain reduced riboflavin and iron-sulfur clusters (5-9), 4-hydroxybutyryl-CoA dehydratase does not require activation by ATP, Mg"1-1" and Ti(III)citrate. In addition the 4-hydroxybutyryl-CoA dehydratase is not inactivated by oxidants as are aromatic nitro compounds, nitrite or hydroxylamine.

Discussion A hypothetical mechanism analogous to that proposed for the dehydration of 2-hydroxyacylCoA (7-9) accounts for the fact that FAD, present in the active enzyme, is in the oxidized state. Hence, the first step should be an oxidation followed by a reduction. Thus, 4-

0

O

4-Hydroxybutyryl-C oA

crotonyl-CoA Fig. 2. Hypothetical mechanism of 4-hydroxybutyryl-CoA dehydratase.

iL

666 hydroxybutyryl-CoA should be oxidized to yield FADH 2 and enzyme-bound 4-hydroxycrotonyl-CoA, a vinylogous 2-hydroxyacyl-CoA. The latter might be reduced by a hydride derived from FADH 2 in an Sjsf2' reaction to yield vinylacetyl-CoA which finally isomerises to crotonyl-CoA. The iron-sulfur cluster may serve as a Lewis acid facilitating the leaving of the hydroxyl group, whereas the enoyl-CoA moiety favours the nucleophilic displacement of the hydroxyl group by a hydride (Fig. 2). The salient feature of this hypothetical mechanism is the turnover of FAD in an overall nonredox reaction. However, this direct involvement of the flavin in the catalytic cycle has not been experimentally shown yet. Furthermore, there are several examples of enzymes that have a flavin requirement for activity for which no role in the catalytic mechanism is known (10); for further references see (11). On the other hand, transient absorbance changes during single turnover experiments strongly indicate that FADH 2 indeed is involved in the chorismate synthase reaction which catalyses the release of phosphate being also a mere elimination (11).

Acknowledgements This work was supported by grants from the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie.

References 1. Hardman, J.K., T.C. Stadtman. 1963. J. Biol. Chem. 238, 2088-2093. 2. Willadsen, P., W. Buckel. 1990. FEMS Lett. 70, 187-192. 3. Scherf, U., W. Buckel. 1991. Appl. Environ. Microbiol. 57, 2699-2702. 4. Scherf, U., W. Buckel. 1993. Eur. J. Biochem. in press. 5. Buckel, W. 1992. FEMS Microbiol. Reviews 88, 211-232. 6. Kuchta, R.D., R.H. Abeles. 1985. J. Biol. Chem. 260, 13181-13189. 7. Hofmeister, A., W. Buckel. 1992. Eur. J. Biochem. 206, 547-552. 8. Klees, A.-G., D. Linder, W. Buckel. 1992. Arch. Microbiol. 158,294-301. 9. Müller, U., A.-G. Klees, K. Bendrat, W. Buckel. This volume. 10. Härtel, U., W. Buckel. This volume. 11. Ramjee, M.N., S. Balasubramanian, C. A.bell, J.R. Coggins, G.M. Davies, T.R. Hawkes, D.J. Lowe, R.N.F. Thorneley. 1992. J. Am. Chem. Soc. 114, 3151-3153.

Glutamate synthase from AzospiriUum brasilense: structural and mechanistic studies.

Maria A. Vanoni Dipartimento di Fisiologia e Biochimica Generali, Università' degli Studi di Milano, Milano, Italy

Introduction Glutamate synthase catalyzes the reductive transfer of the amide group of L-glutamine to the C(2) carbon of 2-oxoglutarate to form two molecules of glutamate. GltS has been purified to homogeneity in our laboratory from AzospiriUum brasilense, a Gram negative nitrogen-fixing bacterium (1). The AzospiriUum enzyme is NADPH-dependent and it shares several properties with the pyridine nucleotide-dependent form of the enzyme found in other microorganisms (2). It is composed by two dissimilar subunits (a subunit: 162 kDa, and 13 subunit: 52.3 kDa), which form the afi protomer. The latter contains one mol FAD, one mol FMN per mol afl. In addition, equimolar amounts of non-heme iron atoms and acid-labile sulfur atoms, which may form several iron-sulfur centers are present. The characterization of AzospiriUum GltS has been carried out in our laboratory by using a combination of kinetic and spectroscopic approaches (3-6). The recent acquisition of the nucleotide sequences of the structural genes encoding AzospiriUum GltS subunits in our laboratory (7), and of the genes encoding GltS from other sources (namely : NADPHdependent GltS from E. coli, NADH-dependent GltS from Alfalfa, and Fd-dependent GltS from maize) (7, 8 and references therein) allowed us to deduce the amino acid sequences of the proteins, and to interpret mechanistic and spectroscopic data available within a structural framework. Results and Discussion Kinetic and Mechanistic Properties of GltS. Steady-state kinetic measurements on the reactions catalyzed by AzospiriUum GltS were carried out in the absence and presence of products and dead-end inhibitors (3). Both glutamine- and ammonia-dependent activities of GltS are well described by a two site uni bi bi

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

668 ping pong mechanism. The pH dependence of the kinetic parameters V and V/K for the substrates of the reactions strongly suggest that the ammonia-dependent reaction of GltS is not physiologically relevant being characterized by a high pH optimum and a low catalytic efficiency with respect to ammonia utilization (Figure 1). The order of addition of substrates to the reduced enzyme during the glutamine-dependent GltS reaction was also determined to be ordered with binding of 2-oxoglutarate followed by that of L-glutamine, thus suggesting the absence of a glutaminase function of GltS at low 2-oxoglutarate concentrations.

Figure 1. pH dependence of the kinetic parameters of GltS glutamine- and ammoniadependent reactions. The curves are drawn assuming the pK values indicated below. Panel a: The V profile of glutamine-dependent reaction (open circles) has been drawn assuming pK values of 6.85 and 9.76. The V profile for the ammonia-dependent reaction (closed circles) was drawn assuming pKa values of 8.77 and 9.96. Panel b: Calculated pK values were 7.7 and 8.4, NADPH concentration was expressed in |iM, Panel c: Calculated pK values were 8 and 8.3, L-glutamine concentration was expressed in mM; Panel d: Calculated pK values were 6 and 9.3, 2-oxoglutarate concentration was expressed in |iM, Panel e: calculated pK value was 8.7, ammonia concentration expressed in M. Velocities are expressed as |imol NADPH oxidized per min per ml of an enzyme solution exhibiting 1.44 U/ml under standard condition.

669 GltS is specific for NADPH. During catalysis the 4-proS H of the reduced pyridine nucleotide is transferred to the enzyme, and it rapidly equilibrates with the solvent. NADPH-dependent reduction of GltS is reversible, as shown by the release of tritium into the solvent, in the absence of net NADPH oxidation when the enzyme is incubated anaerobically in the presence of (4S)-[43H]-NADPH. GltS is also specific for the production of only the L- isomer of glutamate during the enzyme oxidative half reaction as demonstrated by incubating GltS with 2-[U-^C]-oxoglutarate and L-glutamine or ammonia and determining the stereochemistry of glutamate produced with L-glutamate decarboxylase. Apparent reversibility of the glutamate synthase reaction was observed at high pH values. Incubation of GltS with NADP + and Lglutamate at pH 10.3 led to reduction of the pyridine nucleotide. However, chromatographic analysis of the reaction products by using L-[U-^C]glutamate revealed production of R e labelled 2-oxoglutarate but not glutamine, thus indicating that the overall GltS physiological reaction is irreversible, despite the presence of a reversible reductive half reaction. The reversibility of GltS reactions, the presence of glutamate dehydrogenase or glutaminase activities and the mode of transfer of the ammonia unit from glutamine to 2-oxoglutarate were investigated in detail by ^N-NMR spectroscopy (4). Taking advantage of the differences in the resonances of ^N-ammonia, L-(^N-amido)-glutamine, L-(^N-amino)glutamate, ^N-NMR w a s used to monitor the transfer of ^ N from glutamine (or free ammonia) to 2-oxoglutarate (or solvent) under several conditions, at pH 7.5. By using this approach we demonstrated that the glutamine-dependent reaction of GltS is essentially irreversible, that the enzyme does not exhibit any glutamate dehydrogenase or oxidase reaction in either direction, nor any glutaminase activity. Furthermore, no exchange of ammonia with excess ammonium ion in the medium could be detected when GltS was incubated with L-glutamine, 2-oxoglutarate, NADPH and excess ^N-ammonia, leading to the conclusion that during catalysis either the glutamine nitrogen is transferred directly to 2oxoglutarate, or it is released to a site in the enzyme which is not accessible to solvent. The lack of glutaminase activity detected with Azospirillum GltS preparations appears to be peculiar to this enzyme species. The N-terminal region of Azospirillum GltS is similar to those of other GltS, and it shares extensive sequence similarities with the PurF-type L-glutamine amidotransferase domain found in a number of L-glutamine-dependent enzymes (Figure 2 of Ref. 8). In particular, the N-terminal cysteine residue of the domain is conserved, and several histidine and aspartate residues are within regions similar to those that contain these residues in the well characterized phosphoribosyl-amidotransferase from B. subtilis or E. coli (9). Indeed, the Azospirillum GltS is irreversibly inactivated by iodoacetamide, and among enzyme substrates and their analogs tested, only L-glutamine, its analog L-methionine sulfone and, to a lesser

670 extent, L-glutamate exerted complete protection from inactivation at all pH values tested. At pH 7.5, where GltS is stable, only one out of about 4 iodoacetamide modifiable residues appears to be essential for catalysis, and to be protected by L-methionine sulfone. Furthermore, N-terminal sequencing of Azospirillum GltS incubated with [2^CJiodoacetamide in the absence or presence of L-methionine sulfoneshowed release of l^C dpm above background at the first cycle in the sample treated in the absence of L-methionine sulfone. This is consistent with the hypothesis that also in GltS the N-terminal Cys residue of the a subunit is involved in glutamine amide group release during catalysis. Interestingly, the pH dependence of the rate of inactivation of GltS by iodoacetamide increases from a finite low value at low pH, to a limiting upper rate as a group with an apparent pKa of 8.7 dissociates. This result is consistent with the iodoacetamide-modified residue being a cysteine, and with the hypothesis that a Cys-His ion pair may be present in the GltS active site, as proposed for the phosphoribosylamidotransferase from E. coli (9). Characterization of the Flavin Cofactors and Iron-Sulfur Centers of GltS Anaerobic addition of NADPH to Azospirillum GltS led to partial reduction of the enzyme, consistent with the reversibility of the GltS reductive half reaction, and to formation of a long wavelength absorption band centered at approximately 650 nm, which could be ascribed to either a charge transfer complex between a flavin and the pyridine nucleotide or the stabilization of a neutral flavin semiquinone. Apparent complete reduction of the enzyme could be obtained by dithionite reduction, photochemical reduction with the light/deazaflavin system, or in the presence of a NADPH-regenerating system, formed by glucose-6phosphate/glucose-6-phosphate dehydrogenase, during NADPH reductions. In all these cases, the long wavelength absorption band was not observed, while the reduced enzyme spectrum had lower absorbance at wavelengths above 500 nm, indicating that its iron-sulfur centers could also be reduced. Unfortunately, anaerobic reduction of the enzyme with the methods above did not lead to differential changes of absorbance at different wavelengths, which could have allowed us to elucidate the reductive behaviour of each of the flavin and iron-sulfur center cofactors of GltS. In order to differentiate between the flavin cofactors, we studied the reaction between GltS and sulfite. As we have previously shown (5, 6), glutamate synthase flavins could be distinguished on the basis of their reactivity with sulfite. In fact, sulfite addition to GltS caused spectral changes consistent with the formation of a flavin N(5)-sulfite adduct with only one of the enzyme flavins, and the presence of sulfite did not affect spectral changes observed on anaerobic reduction of GltS with excess NADPH, including the appearence of a long wavelength absorption band. Finally, among the enzyme substrates and their analogs, only

671 2oxoglutarate competitively displaced sulfite from the enzyme flavin. These results, which were confirmed kinetically, led us to the proposal that in GltS, NADPH reacts with the flavin at the first enzyme sub-site. The electrons flow through at least some of the iron-sulfur centers to the second flavin, which can react with sulfite, and which reduces during catalysis, the iminoglutarate formed on addition of ammonia (from glutamine) to 2-oxoglutarate. The EPR properties of oxidized and reduced forms of GltS have been examined using different GltS preparations. The purified enzyme exhibits an EPR signal with properties consistent with the presence of one [3Fe-4S]°. 1+ cluster per afi protomer ( FeSj). Reduction of the enzyme with excess NADPH led to greater than 98 % reduction of this cluster, to formation of a free organic radical signal (0.35 spin/al3 protomer, g=2.00) which was assigned to a flavin neutral semiquinone on the basis of its linewidth of 19.5 G, and allows us to identify the long wavelength absorption band observed by absorption spectroscopy (Figure 2A). A weak g=1.96 signal accounting for 0.25 spin/afl protomer was also detected. Reduction of the enzyme by NADPH, in the presence of the NADPH-regenerating signal led to reduction of the [3Fe-4S]®>

center, only to a small amount of flavin semiquinone, and to

reduction of a second iron-sulfur center (FeSji, g=1.96) which accounts for 0.9 unpaired electrons/aB protomer (Figure 2B). The temperature-dependence and power saturation behaviour of this signal, along with circular dichroism spectra of the oxidized and reduced forms of GltS, suggest that this center is more likely a [4Fe-4S]l + >2 + center rather than a [2Fe-2S] 1+ >2 + cluster. Reduction of GltS with the light/deazaflavin system led to absorption and circular dichroism changes indistinguishable from those observed during reduction of GltS with NADPH and the NADPH-regenerating system. However, the EPR spectrum of photoreduced enzyme was more intense and different from that observed by using NADPH as reductant. Double integration of the EPR signal allowed us to calculate the presence of 1.9 unpaired electrons/aB protomer (Figure 2B). Again temperature-dependence and power saturation behaviour of the EPR signal, as well as the circular dichroism properties of the deazaflavin-reduced enzyme are consistent with the known properties of [4Fe-4S]l + >2 + centers rather than with those of [2Fe-2S] 1+ >2 + clusters. A careful re-evaluation of the non-heme iron content of GltS yielded approximately 12 gatom non-heme iron/aB protomer, rather than 8, which certainly is consistent with the presence of three distinct iron-sulfur centers in the GltS protomer. The presence of a [3Fe-4S]®>'+ center is also corroborated by the finding that the Fd-GltS from spinach contains only one center which was also identified as a [3Fe-4S]0>+l cluster (10). Analysis of the primary structure of GltS from different sources, together with the evidence for the presence of a single iron-sulfur center in Fd-GltS from plant, allowed us to propose that the single conserved cluster of cysteines in GltS (region 1102-1114 of the Azospirillum

672 Figure 2. EPR spectra of oxidized and reduced forms of Azospirillum GltS. Panel A: Comparison of the EPR spectra of GltS (30.1 (iM) in 25 mM Hepes/K + buffer, pH 7.5, 1 mMEDTA, 1 mMDTT, and 10% glycerol before and after the anaerobic addition of 33-fold molar excess NADPH. Instrument settings were as follows: 9.42 GHz; modulation amplitude, 8 G; scan speed, 200 G/min; microwave power, 0.02 mW; gain, 8 x 10^. The temperatures are indicated in the figure. Panel B: Comparison of the EPR spectra of47.85 ^M GltS before (Upper trace) and after reduction by NADPH in the presence of a NADPHregenerating system (Middle trace) or after photochemical reduction with the light/deazaflavin system (Lower trace). Istrument settings were as in panel A, except all spectra were recorded at 5 K and were normalized to 0.2 raW microwave power.

5°K

• 33-fold excess NADPH

xS

70 ° K

« 10

2900

3300

> 0.2

3700

4100

B

GltS

r ^

NAOPH-reduced GUS

12

V1

1

»23

J

1

deszaRf-reduced GltS TOO

1900

J

2S00

I

3400

Magnetic Field (Gauss)

NADPH

NADP*

Flavin 1

FeS„i

L-GIn

Fe Fe33 S 4

FeS„

\*

Flavin 2

L-Glu

NH 3 2- IG 2-OG

Scheme I

L-Glu

«300

673 GltS a subunit) provides the ligands for the formation of this center. The hypothesis is further supported by the similarity of the cysteine spacing with that found in Cluster 3 of fumarate reductase and succinate dehydrogenase iron proteins. Additional cysteine clusters in GltS, which are conserved in the enzyme from different sources, are found only in the 13 subunit of bacterial GltS, and in the C-terminal fourth of the NADH-dependent GltS from Alfalfa (7, 8). N o similarity is found between the cysteine spacing of these regions and that of other ironsulfur containing proteins whose sequences are available to date. While it is possible to rule out the presence of simple ferredoxin-type Fe-S centers in GltS, further experiments will be required to elucidate the structure and properties of the centers of bacterial GltS. The evidence of two centers ( FeSj and FeSji ), which can be reduced by NADPH, leads us to a refinement of the scheme for GltS active site ( Scheme I), which includes centers FeSj (the [3Fe-4S]°> 1+ cluster) and FeSjj as electron transporters between the enzyme flavins. As of the role of the low potential iron-sulfur center (FeSju ), more experiments are required to determine whether it is also involved in the intramolecular electron transfer process, or it serves a catalytic, structural or regulatory role in GltS.

Acknowledgements This work was supported by grants from Consiglio Nazionale delle Ricerche and Ministero per l'Università' e la Ricerca Scientifica e Tecnologica, Rome, Italy. The ^ N - N M R , EPR and circular dichroism experiments were carried out in the laboratory of Dr. Dale E. Edmondson, and were supported by grants from the National Science Foundation (U.S.-Italy Cooperative Science Program INT-8815289 and DMB-90-08173 to D E E.) and from the Consiglio Nazionale delle Ricerche (Progetti Bilaterali Italia-US. A. CT89/90.04140 to B.C. and CT92.01111 CT04 to M.A.V.),

References

1. Vanoni, M. A., G. Zanetti, B. Curti. 1991. In: Chemistry and biochemistry of flavoproteins (F. Muller ed.) Vol. III. De Gruyter p. 309. 2. Ratti, S., B. Curti, G. Zanetti, E. Galli. 1985. J. Bacteriol. 163:724. 3. Vanoni, M. A., L. Nuzzi, M. Rescigno, G. Zanetti, B. Curti. 1991. Eur. J. Biochem. 202:181.

674 4. Vanoni, M. A., D. E. Edmondson, M. Rescigno, G. Zanetti, B. Curti. 1991. Biochemistry 30:11478 5. Vanoni, M. A., G. Zanetti, B. Curti, D. E. Edmondson. 1991. In "Flavins and flavoproteins" ( Curti, B., S. Ronchi, G. Zanetti, eds.) De Gruyter, Berlin p. 749. 6. Vanoni, M. A., D. E. Edmondson, G. Zanetti, B. Curti. 1992. Biochemistry 31:4613. 17. Pelanda, R., M. A. Vanoni, M. Perego, L. Piubelli, A. Galizzi, B. Curti, G. Zanetti. 1993. J. Biol. Chem. 268:3099. 8. Pelanda, R., L. Piubelli, P. Fumagalli, A. Mazzoni, E. Verzotti, M. A. Vanoni, G. Zanetti, B. Curti, this volume. 9. Knaff, D. B., M. Hirasawa, E. Ameyibor, W. Fu, M. K. Johnson. 1991. J. Biol. Chem. 266:15080. 10. Bower, S., H. Zalkin. 1983. Biochemistry 22:1613.

Cloning and Expression of Azospirillum brasilense Glutamate Synthase

R. Pelanda, L. Piubelli, P. Fumagalli, A. Mazzoni, E. Verzotti, M A. Vanoni, G. Zanetti, B. Curti Dipartimento di Fisiologia e Biochimica Generali, Università' degli Studi di Milano, Milano, Italy

Introduction Glutamate synthase (GltS) is a complex iron-sulfur flavoprotein containing one mol FAD, one mol FMN, and three distinct iron-sulfur centers per aB protomer ( a subunit: 162 kDa; 6 subunit: 52.3 kDa) (1). The pyridine-nucleotide-dependent GltS forms with glutamine synthetase (GS) the major pathway for ammonia assimilation in microorganisms. In particular, nitrogen-fixing bacteria rely on the GS-GltS pathway for growth. Photosynthetic microorganisms and higher plants also contain a ferredoxin-dependent form of glutamate synthase (Fd-GltS), which is composed by a single polypeptide chain similar in mass to the bacterial GltS a subunit, and differs from the bacterial NAD(P)H-dependent enzyme for a lower iron-sulfur centers content. The enzyme from Azospirillum, a diazotroph, has been purified in our laboratory (2), where it is being characterized by a combination of kinetic, mechanistic, and spectroscopic approaches (3). In order to gain information on the enzyme primary structure, to obtain large quantities of Azospirillum GltS, and, eventually, to perform both site-directed mutagenesis experiments and studies on the regulation of the in vivo expression of the enzyme, the cloning of Azospirillum brasilense glutamate synthase genes has been undertaken.

Results and Discussion Cloning of Glutamate Synthase Structural Genes. A 10 kb EcoRI fragment of Azospirillum brasilense DNA containing the structural genes encoding the a and 8 subunits of GltS was identified by using a combination of heterologous

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

676 DNA probes and oligonucleotides synthesized on the basis of the N-terminal amino acid sequences of the isolated GltS subunits and of internal peptides (4). Sequencing of the 10 kb EcoRI fragment revealed the presence of gltD (1443 nucleotides) and gltB (4545 nucleotides) genes encoding the precursors of the 13 and the a subunits of Azospirillum GltS, respectively. Post-translational processing of the encoded polypeptides to yield the mature enzyme subunits occurs with removal of the N-terminal methionine residue from the 13 subunit precursor, and of a 36 residues N-terminal peptide from the a subunit precursor. The deduced amino acid sequence of GltS a subunit is very similar to that deduced from the sequence of the genes encoding the a subunit of E. coli GltS, and to that of the single polypeptide chain of maize Fd-GltS. The primary structure of Azospirillum 6 subunit is instead similar to that of the E. coli 13 subunit. The similarity of GltS a subunit extends to the Nterminal region of barley Fd-GltS (5), and both a and 13 subunits are similar to the single polypeptide chain of Alfalfa NADH-dependent GltS (6). In particular, the Azospirillum

a

subunit is similar to the N-terminal region of the Alfalfa enzyme, while the 13 subunit is similar to the C-terminal fourth of the polypeptide (6). Tentative Localization of Substrate and Cofactors Binding Regions within GltS Subunits. Comparison of the amino acid sequences of GltS from different sources with those of enzyme using the same substrates and cofactors, allowed us to tentatively identify functional regions of the protein (4, and references therein, Figure 1). The N-terminal region of GltS a subunit is similar to the PurF-type L-glutamine-dependent amidotransferase binding domain, and it contains conserved cysteine, histidine and aspartate residues which may be involved in the release of ammonia from glutamine, during catalysis. A region similar to the C-terminal part of the FMN binding domain of the flavocytochrome b2type class is also found between residues 1049 and 1100. This region is followed by a sequence (residues 1102-1114) containing three cysteines conserved among all GltS, which may be responsible for the formation of the [3Fe-4S]®>+^ cluster of GltS. This assignment was based on the similarity of this sequence to those of fumarate reductase and succinate dehydrogenase, which bind Cluster 3, the [3Fe-4S]®>1+ center of these enzymes, and on the evidence of the presence of a single iron-sulfur cluster, a [3Fe-4S]°> 1+ cluster, in plant FdGltS. A glycine-rich region, which matches the consensus sequence for an adenylate-binding fold (7) is also present in the C-terminal part of the GltS a subunit (residues 1354-1396) along with a sequence (residues 1281-1291) similar to the second FAD consensus sequence found in several flavoproteins (8). The 13 subunit of GltS shares no sequence similarity with the

677 ferredoxin-dependent plant enzyme, and it contains two distinct cysteine-rich regions (residues 47-59 and 94-108), which might provide some of the ligands for the formation of the remaining iron-sulfur centers of GltS. Two glycine-rich regions (residues 149-177 and 291339), which match the consensus sequence for the formation of adenylate binding folds are also present in the GltS 13 subunit. In addition a sequence (residues 432-442) which matches the second FAD consensus sequence (8) is present in both bacterial GltS 6 subunit and Alfalfa protein.

3Fe4S

ADP

ADP

• FMN

•v

* FAD,,

GAT 3Fe4S.

7*

FMN GAT

T

ll

3Fe4S

A.B. NADPH-GltS:

FAD,,

fill

E.C.

NADPH-GltS

Z.M.

Fd-GltS

A.A.

NADH-GltS

ADP

GAT

I

NADPH-GltS

FAD,,

FeS

T ADP r

A.B.

FeS

F

I—flTTI 11 II

ADP

V 4 ADP

AD„

II

FAD, FMN, [3Fe-4S], FeS„, FeS m

E.C. NADPH-GltS:

FAD, FMN, [3Fe-4S], FeS„, FeS M |?

Z.M. Fd-GltS:

FAD, FMN, [3Fe-4S]

A.A. NADH-GltS:

FAD?, FMN?, [3Fe-4S]?, FeS n ?, FeS,„?

Figure 1. Comparison of glutamate synthases from Azospirillum brasilense (A.B. NADPHGltS), E. coli (E.C. NADPH-GltS), maize (Z.M. Fd-GltS) and Alfalfa (A. A. NADH-GltS). The conserved regions which were tentatively identified as the PurF-type amidotransferase domain (GAT), the FMN binding region (FMN), the cysteine clusters which form the [3Fe4S] (3Fe4S), and centers FeSfj and FeSju (FeS), adenylate binding sites (ADP), and the second consensus FAD binding sequence (FADjj) are indicated, along with the cofactor content reported for each of the GltS. A question mark indicates the absence of experimental evidence for the presence of the cofactor.

678 One of the glycine-rich regions was proposed by us to bind NADPH, while the other was postulated to represent a site for allosteric regulation of GltS by a (di-)nucleotide. In fact, the presence of FAD bound to the single polypeptide chain of Fd-GltS, and the presence of a potential adenylate binding fold in the C-terminal region of GltS a subunit led us to conclude (4) that region 1354-1396 of the a subunit served for FAD binding, in spite of the presence of a potential adenylate binding fold and of a sequence similar to the second FAD consensus sequence, within the 13 subunit. It was also proposed (4) the second glycine-rich region of the GltS 13 subunit to be the candidate for the binding of NADPH in both bacterial GltS on the basis of the distinctive features of NADPH binding regions (9). The Alfalfa GltS is NADH-dependent, and also contains two glycine-rich regions in its C-terminal fourth, which is similar to the bacterial GltS 13 subunit. The region of Alfalfa GltS corresponding to residues 291-339 of Azospirillum GltS 13 subunit contains a GXGXXG motif rather than the GXGXXA sequence found in Azospirillum GltS. In addition, the amino acid sequences beyond residue 322 (the residue numbering refers to the Azospirillum protein) are poorly conserved between bacterial and plant enzymes. Arg-321 has been proposed to be the positively charged residue which characterizes the C-terminal part of the potential NADPH binding site of GltS, together with the absence of a preceeding negatively charged residue (9). The observation that in the NADH-dependent Alfalfa enzyme there is no counterpart of Arg-321, and that the residue preceeding this position is a glutamic acid, reinforces our hypothesis (Figure 2).

Figure 2. Sequence comparison of glutamate synthases from A. brasilense (A.b.), E. coli ( E.c.), maize (Z.M.), Alfalfa (AA), barley (H.V.). Amidotransferase domain: the stars indicate residues which have been demonstrated to be conserved in PurF-type amidotransferases. For comparison, the N-terminal sequence ofE. coli phosphoribosylamidotransferase (E C. Purl) is shown. Cysteine-rich regions: the cysteine spacing is indicated along with the cysteine-rich region of E. coli fumarate reductase (E.C. FrdB) which forms the [3Fe-4S] cluster of this enzyme. FMN binding domain: the residues, which in Saccharomyces cerevisiae flavocytochrome b2 (SC. B2) interact with the isoalloxazine (i) orribitylportion (r) of FMN, and those that interact with the substrate (s) are indicated. Glycine-rich regions and Second FAD consensus sequence: residues that match or do not match the consensus sequence for the formation of an adenylate binding fold (7) or the second FAD consensus sequence (8) are indicated by + or -, respectively. The dots mark positive residues within the sequence as discussed in the text. The arrow indicates the alanine residue of NADPH-dependent GltS which is substituted by a glycine in NADH-dependent GltS.

679

L-glutamine amidotransferase domain A.b.

a

lcavariA

30HRaAVDJU>GFXaDa

206rYHQRYSTHT

228LM(HSIIHTV

346DFNS

i.e.

a

43csraiiiA

72KROAIIJUX3KTGDQ

242LFHQI17STNT

2 66ZTVK5TF5FV

380DRKQ

98CSVSIVA

127KROOCOADSDSODO

304IYKFBTSTHT

326LSKH3ZXHTI

447DBN9

102CSVSTVA

lSiKRGncacxkmaDo

311IiZHSRTSTMT

3 33I/3KNQZXNTL

453DKNO

lxGvarvA

SOHKGaanDMDSGDO

207IYHRRPSTNT

229LQKHSZXNTI

350DBMO

99LAHNQNLTNA

99LAHHOKLTNA

Z.M. AA H.V.

25KR3. .. QDAAGIIT

Cysteine-rich regions A.B.

1095

ASLIAMSCMVBQCHSHTCPVGVCVQDDKIJl

1125

I.C.

1123

OPMVALGClCfLMCKLHNCATGVATQDDKLR

1153

Z.M.

1222

VMIIATOCVMMaCXTlOICPVOVASQRBnJl

1252

AA

1239

APLITLSCIMOKCXRITCiVSIAZgDFVLR

1269

CXXXXXCXXXXCP 197

SQNGVNSCTrVGXCSZV. CPKHVDPAAAIQQ

226

A.B. a

38 UUUOQllHIt.CSQCaVPrCgVH. . .CPVSNHIPEWLKLT

72

I.e. B

3 8 A X t P n U O i T A A C S A A N . F Y C B M K . . .CFVHNYIPNHL1QJI

72

AA

1 7 1 5 PLLKigSlUt.CMDCalPrCMQINSaCFLSHKIFIIIIKLV

1752

CXXXXXXXCXXXXXXCP

A.B. a

91 PKlCGRlCPQDBliCBGNCVl

B.C. £

9 1 PBVCGKVCPQDBXiCBGSCTIi

110

1771 PBrrSBVCPAP. .CBOSCVL

1788

AA

110

FMN binding domain A.B.

a

980

LAQ.LIY.D. . .LKQIHH»KVTVia.V. . . S B S G I S T I A A S V M O U a D I I L I S G N S S a

B.C. a

1008

LAQ.LXF.D. ..LKQVNPKMCSVKLV. . . SBPOVaTIATOVAXAXADLITIlUHDSa

1057

Z.M. a

1107

LAQ.Lir.O. . .LHgntPKkKVSVKLV. . . SIAOIOTVASaVSKAKADIiaiSOHDOO

1156

AA

1114

LAQ.LIH.D. . .iraONPAARISVKLV. . . SBAGVSVIASOWlCQlIABHVLISaXDaG

1174

a

1029

1 S.C. B2

a.b.

326

IDPSLTWKDIULK.KKTF1PTVXKGVQRT*DVI

10UI. B I S V . S O W L .

370

a

1030

TOASPQrsiicrA0&pwn«asBV*gvi,TunajuiKViujm>QQo u o> 11

20

Is

10

• U • •

L929 L929 + Mo 3T3 3T3 + Mo

Fig. 3 Effect of sodium molybdate on the capacity of L929 and NIH3T3 cell extracts to restore nitrate reductase enzymatic activity in extracts of mycelia obtained from the nit-1 mutant of Neurospora crassa. L929 and NIH 3T3 cells were exposed to medium alone or medium containing 5 mM sodium molybdate for 24 hours as indicated. Cells were harvested and homogenized and an aliquot of the homogenate containing the same amount of proteins (180 ng) was incubated overnight at 4°C with or without extracts of mycelia from nit-1 Neurospora crassa in the absence (-Mo) or in the presence (+Mo) of 1 mM sodium molybdate. Nitrate reductase activity was measured in reconstituted extracts and the results are normalized for the amount of protein present in the L929 and NIH3T3 cell homogenates. Values are expressed in changes of absorbance at 540 nm/min x mg. protein in the eukaryotic cell extract. N.D. = not determined because below the limit of detection of the assay. Each experimental value is the mean ± S.D. of three assays.

Therefore, L929 cells have a defect in one or more of the metabolic steps leading to the synthesis of the molybdenum cofactor.

REFERENCES 1. Johnson J.L. 1980. In: Molybdenum and Molybdenum-Containing Enzymes, ed. Coughlan, M P . (Pergamon, Oxford), pp. 345-383. 2. Terao, M., Cazzaniga, G., Ghezzi, P., Bianchi, M., Falciani, F., Perani, P. and Garattini, E. 1992 Biochem.J. 283: 863-870. 3. Della Corte, E. & Stirpe, F. 1968 Biochem.J. 108: 349-351. 4. Della Corte, E. & Stirpe, F. 1972 Biochem.J. 126, 739-745. 5. Falciani, F., Ghezzi, P., Terao, M., Cazzaniga, G. and Garattini, E. Biochem.J. 285,1001-1008.

1992

P h y s i c o c h e m i c a l P r o p e r t i e s of Retinal O x i d a s e P u r i f i e d f r o m Rabbit H e p a t i c Cytosol

Maki Tsujita, Shuhei Tomita, Yoshinori Matsuo, Shigetoshi Miura and Yoshiyuki Ichikawa Department of Biochemistry, Kagawa Medical School, Kita-gun, Kagawa 761-07, Japan

Introduction Retinoic acid is an important lipo-bioactivator for mammals. Since the 1880s, research on the abilities of retinoic acid as to cell differentiation and morphogenesis has increased and attracted attention. In 1983, Chambon et a/.(l), and Evans et al.(2) found the nuclear retinoic acid receptor (RAR), and reported that a complex of retinoic acid with its receptor controls gene regulations. Since then, it has been well known that retinoic acid plays an important role in life maintenance. However, the biosynthesis of retinoic acid has not been established. To understand the enzymatic systems in retinoic acid biosynthesis, the activity of retinoic acid synthesis from all-trans-retinal was investigated in various tissues and organs from female and male rabbits. The activity was observed in the liver, lung, kidney, ovary and pituitary body, the highest activity being found in the liver. Most of the retinoic acid synthase activity (over 99 %) was found in the cytosol and microsomes. The retinoic acid synthase localized in microsomes required NADPH as an electron donor. One atom of molecular oxygen was incorporated into a retinoic acid molecule by a cytochrome P450-linked monooxygenase system. The substrate-induced difference spectrum of the synthase with aW-ira/w-retinal was typical of type I, the high spin form being induced from a low spin cytochrome P-450. The microsomal retinoic acid synthesis was inhibited by anti-NADPH cytochrome P-450 reductase IgG, phenylisocyanide and carbon monoxide (CO). Furthermore, the activity inhibited by CO was photoreversiblly restored. These results demonstrated that this microsomal retinoic acid synthase is a cytochrome P-450-linked monooxygenase system. A cytochrome P-450-linked retinal monooxygenase has been reported (3,4). The majority of retinoic acid was also synthesized aerobically in the liver cytosol, thus, hereafter, we designate the enzyme as 'retinal oxidase'. The retinal oxidase was purified electrophoretically, as a single protein band, from rabbit liver cytosol. The characteristic properties, enzymatic reaction mechanism and substrate specificity of the retinal oxidase were investigated (5).

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

744 Results and Discussion The relative molecular mass of the purified retinal oxidase was estimated to be 270 KDa by HPLC gel filtration on a TSKgel G3000SWXL column. The minimum relative molecular mass of the retinal oxidase was determined to be 135 KDa by SDS-PAGE. This indicates that this oxidase comprises a homodimer of two subunits. However, the retinal oxidase purified from frozen rabbit livers (at -20 °C) changed easily to a heterodimer. It was found that the conversion from the homodimer to the heterodimer of the retinal oxidase was dependent on the experimental conditions for the preservation and purification of the rabbit liver (6). This will be due to endogenous proteinases. The apo-retinal oxidase was prepared from retinal oxidase by the 2.2 M potassium bromide method. The activity of the retinal oxidase was completely abolished with this treatment, due to the removal of FAD. The addition of 250 |iM FAD to the apo-enzyme solution restored 75 % of the initial activity. This was confirmed by analysis of the coenzyme of the retinal oxidase by thin layer chromatography. The amino acid residue, disulfide bond, inorganic sulfur, flavin and metal compositions of the retinal oxidase were determined. Two FADs, two molybdenums, eight irons and eight labile sulfurs were found in one molecule (270 KDa) of the retinal oxidase. This indicates that the oxidase is a metalloflavoenzyme. The retinal oxidase contains about 52.8 % hydrophobic amino acid residues, 26 cysteinyl residues and two disulfides. The optical absorption spectra of the retinal oxidase showed absorption peaks at 275 nm, 340 nm and 450 nm, and shoulders at 420 nm and 473 nm, for the oxidized form. The retinal oxidase was reduced anaerobically by the addition of retinal or sodium dithionite, and in both cases the visible absorption spectra decreased and disappeared for the reduced form. The retinal oxidase could not be reduced anaerobically by 0.2 mM NADPH or NADH. Circular dichroism spectra of the retinal oxidase, unlike the circular dichroism spectrum of a free FAD, showed a positive absorption peak at 428 nm in the visible region, and a negative absorption peak at 234 nm in the ultraviolet region. This was probably due to interaction between the irons and molybdenums, and the apoenzyme of the retinal oxidase. The activity of the oxidase was not affected by any cofactors, such as N A D P + . NAD+NADPH and NADH, and it did not occur under anaerobic conditions. Catalytic activity of the retinal oxidase was examined with respect to various substrates, such as all-trans-retinal,

9-

cr'i-retinal, 75-ci's-retinal, a//-irans-retinol, N-methylnicotinamide and hypoxanthine. The ability to catalyze the oxidation of the substrates depended on the nature of the group with which aldehyde was substituted, and the aldehyde may be replaced by an amide group. The Km value for all-trans-retinal (8 (J.M) of the retinal oxidase was much smaller than those for

745 9-cis-retinal, ii-rii-retinal and N-methylnicotinamide. No enzymatic reaction was observed with hypoxanthine or all-trans-retiml,

but JV-methylnicotinamide was an effective substrate

for the oxidase. Accordingly, the substrate specificity of the retinal oxidase is relatively broad. The effect of B O F - 4 2 7 2

(sodium-8-(3-methoxy-4-phenylsulfinylphenyl)pyrazolo[l,5-a-]-

l , 3 , 5 - t r i a z i n e - 4 - o l a t e m o n o h y d r a t e ) , a s p e c i f i c and p o t e n t i n h i b i t o r o f x a n t h i n e dehydrogenase, on the activity of the retinal oxidase was examined. The activity of bovine xanthine dehydrogenase was inhibited completely by 2 nM B O F - 4 2 7 2 , but the rabbit retinal

microsomal P-450-linked monooxygenase (7)

all-trans retinyl ester all-trans retinol

microsomal NAD(P)H-alcohol dehydrogenase (8)

ß-carotene all-trans

retinal

02* cy toso lie retinal oxidase (5)

microsomal P-450-linked monooxygenase (3,4)

all-trans retinoic acid

all-trans retinoic acid

Figure 1. Physicological biosynthases of retinoic acid in rabbit hepatocytes. Arrow marks shows the directions of the enzymatic reaction. Open arrows indicate the monooxygenase reactions. O2*, molecular oxygen; 0 # , oxygen of water.

746 oxidase activity could not be inhibited even by 10 nM BOF-4272. Although rat anti-xanthine dehydrogenase IgG could recognized rat, rabbit and bovine xanthine dehydrogenase, and apparently inhibited the activities of rabbit and bovine xanthine dehydrogenases as well as that of rat xanthine dehydrogenase, anti-xanthine dehydrogenase IgG failed to inhibit the activity of the retinal oxidase at 10 mg IgG per mg protein of retinal oxidase under the same experimental conditions. The activity of retinoic acid synthesis of retinal oxidase was inhibited competitively by 10 mM pyridoxal, a substrate of aldehyde oxidase, but not effected by acetaldehyde, a substrate of aldehyde dehydrogenase. This indicates that the retinal oxidase is neither a xanthine dehydrogenase nor an aldehyde dehydrogenase localized in the rabbit liver cytosol. The relative molecular mass, number of subunits and cofactor contents of the retinal oxidase were consistent with those of aldehyde oxidase. However, it is not clear whether retinal oxidase is an aldehyde oxidase, because the amino acid composition and sequence of aldehyde oxidase have not been reported previously. Experiments on retinoic acid synthesis in microsome and cytosol of rabbit liver under 18C>2 or H 2 1 8 0 demonstrated that the oxygen of water was incorporated into retinoic acid by the retinal o x i d a s e , and m o l e c u l a r o x y g e n into r e t i n o i c a c i d by the c y t o c h r o m e P - 4 5 0 - l i n k e d monooxygenase system (Figure 1).

References

1.

Petkovich, M „ Brande, N.J., Krust, A. and Chambon, P. 1987. Nature 330: 444-450.

2.

Gigure, V., Ong, E.S., Segui, P. and Evans, R.M. 1987. Nature 330: 624-629.

3.

Matsuo, Y„ Tomita, S. and Ichikawa, Y. 1991. Symposium of Retinoids, pp21 (Tokyo).

4.

Tomita, S„ Tsujita, M „ Matsuo, Y„ Yubisui, T. and Ichikawa, Y. Int. J. Biochem. (in press).

5.

Tsujita, M., Tomita, S„ Miura, S. and Ichikawa, Y. Biochm. Biophys. Acta, (in press).

6.

Shin, M., Tsujita, M., Tomizawa, H., Sakihama, N., Kamei, K. and Oshino, R. 1990. Arch. Biochem. Biophys. 279: 97-103.

7.

Leo, M.A. and Liever, C.S. 1985. J. Biol. Chem. 260: 5228-5231.

8.

Fidge, N.H. and Goodman, D.S. 1968. J. Biol. Chem. 243: 4372-4379.

Mutagenesis of the Cysteine Residues of the FAD Domain of Nitrate Reductase

Wilbur H. Campbell and U.N. Dwivedi Phytotechnology Research Center, Michigan Technological University, Houghton, Michigan 49945 USA

Introduction NADH:nitrate reductase (NR; EC 1.6.6.1) has three domains with each one housing one of its cofactors: FAD, heme-Fe and Mo-pterin (1). We have expressed the FAD domain in Escherichia coli in a soluble form with ferricyanide reductase activity (2). The recombinant FAD domain has been crystallized but a structural model has not yet been derived (3). It has long been thought that a key Cys residue resides in the FAD domain being involved in NADH binding. This belief derived from the inhibition of NADH:NR activity by p-chloromercuirbenzoate (pCMB) and loss of NR dehydrogenase activity concomitantly. This loss of activities could be prevented by either NADH or NAD. Sequence comparisons of NR from a number of sources and mammalian cytochrome b-5 reductase suggested that only one Cys was conserved in the FAD domain (4). To investigate if this highly conserved Cys residue is the same one which reacts with pCMB, we mutated all the Cys residues in the FAD domain to Ser residues.

Results The recombinant FAD domain derived from corn leaf NADH:NR has five Cys residues which will be called CI5, C52, C60, CI43 and C242. This numbering is based on a revised sequence for the FAD domain, which differs from the original by three added residues (2). The highly conserved Cys residue is C242. Five mutant proteins were generated by site-directed mutagenesis using PCR-based unique site elimination directly on the FAD domain insert DNA in the double-stranded pET vector (5). The presence of the desired mutation was established by nucleotide sequencing. The five mutants were less active than the wild-type, retaining from 15 to 65% of wild-type ferricyanide reductase activity (Table 1). Despite these reduced activities, all mutants were purified using Blue Sepharose (2).

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

748 Table 1. Comparison of FAD Domain Wild-Type and Cys to Ser Mutants.

Crude Extract

Mutant

Purified Protein

Specific

FAD

Specific

Visual

Thermal

Activity

Domain

Activity

Color

Instability

(uts/mg)l

Protein

(uts/mg)

(Hg/ml) Wild Type

( % Loss)

2

121.0

26

1320

Yellow

0

C15S

18.3

13

200

Colorless

26

C52S

79.0

14

974

Yellow

0

C60S

52.8

11

530

Colorless

28

C143S

25.4

19

330

Colorless

45

C242S

40.8

18

362

Yellow

4

lunit = (amoles NADH oxidized/min with ferricyanide as electron acceptor, ^ng/ml = amount of FAD protein found in Western blot by densitometry.

Since mutant FAD domains bound to blue Sepharose were eluted by NADH, the NADH binding site was intact and not disturbed to any great extent by Cys to Ser mutations. On the other hand, the typical flavoprotein visible spectrum observed for oxidized FAD domain was not found for all mutants. This is represented by a simple designation of "yellow" or "colorless" in Table 1. Furthermore, it was found that the thermal stability was markedly altered for the mutants which were found to be colorless (Table 1). Thermal inactivation was measured by incubation at 45°C for 15 min in the presence of 100 |iM FAD at pH 7.0. The sensitivity to pCMB was determined by incubation on ice with various molar ratios of pCMB to FAD domain (using either the absorbance at 460 nm or 280 nm to estimate the concentration of protein) for 5 min followed immediately by ferricyanide reductase activity assay (Fig. 1).

749 Figure 1. pCMB Inhibition of Ferricyanide Reductase Activity of Wild-Type and Mutant FAD Domains.

Activity Remaining (%)

100

80 60 40

20 0 C242S C52S C60S Wildtype

Zero One Three Five Ratio of pCMB:Protein

Discussion Based on a sequence comparison of NR and related flavoprotein reductases (4), a prediction for the secondary structure of the FAD domain of NR has been made which was guided by the tertiary structure model of ferredoxin NADP reductase (6). This secondary structure model predicts that CI 5, C52 and C60 of the FAD domain should be more closely associated with the FAD binding site, while C143 and C242 should be more closely associated with the NADH binding site. The results obtained for Cys to Ser mutations agree with this prediction for some residues, namely CI5, C60 and C242, while less so for C52 and C143. Mutants C15S and C60S were colorless, suggesting FAD is not bound with the wild-type conformation. Mutation C242S yielded a protein with a normal flavoprotein spectrum, which is consistent with this Cys being more associated with NADH binding than FAD binding. In contrast, mutation C52S yielded a protein with a normal flavoprotein spectrum, as if this Cys were not closely associated with FAD binding. Mutation C143S yielded a colorless protein, which indicates that despite the prediction this Cys is in a region involved in binding the pyrophosphate bridge of NADH (4), alteration disturbs FAD binding. CI43 is not conserved in NR sequences and is most often found to be Ala. When mutation C143 A was generated, it had a spectrum and activity very similar to the wild-type. Overall our results indicate that CI 5, C60 and CI 43 are residues buried in the interior of the

750 FAD domain conformation and substitutions lead to alteration of FAD binding and decrease of activity, which is totally dependent on the presence of FAD in the protein. The C52 residue may be near the surface as indicated both by the retention of a normal flavoprotein spectrum and the slightly decreased inhibitory effect of pCMB with the C52S mutant (Fig. 1). Most of these mutant proteins are being crystallized and will be analyzed in comparison to the wild-type. The C242S is the most interesting mutant since the protein is produced in near normal quantities (Western blot data shown in Table 1), while having a strong decrease in activity with a spectrum virtually identical to the wild-type. The C242S mutant was not inhibited at all by pCMB, even when it was in 5 fold excess (Fig. 1). These results suggest that C242 is the key Cys residue in the FAD domain which reacts with pCMB leading to inhibition of the enzyme activity. However, the C242S mutant could be purified on blue Sepharose with NADH elution suggesting the NADH binding site was normal. Kinetic analysis indicated that the C242S had virtually the same Km values for both NADH and Fe(CN) 3 " as the wildtype (data not shown). Hence, it can be concluded that the C242 is probably involved more in electron transfer from NADH to FAD than with direct binding of NADH, which is supported by the low activity of this mutant as compared to the wild-type. Acknowledgment Research supported by National Science Foundation Grant. U.N.D. is the recipient of a Biotechnology Overseas Associateship (Long Term) from the Dept. of Biotechnology, Government of India. References

1. Campbell, W.H. and J R. Kinghorn. 1989. Trends in Biochem. Sci. 15, 315-319. 2. Hyde, G.E. and W.H. Campbell. 1990. Biochem. Biophys. Res. Comm. 168, 1285-1291. 3. Lu, G., W. Campbell, Y. Lindqvist and G. Schneider. 1992. J. Mol. Biol. 224, 277-279. 4. Hyde, G.E., N. Crawford and W. Campbell. 1991. J. Biol. Chem. 266, 23542-23547. 5. Ray. F A. and J.A. Nickoloff. 1992. BioTechniques 13, 342-346. 6. Karplus, P.A., M.J. Daniels and J R. Herroit. 1991. Science 251, 60-66.

Electron Transfer and Prototropic Equilibria in Two Complex Metalloflavoproteins

Russ Hille and Ronald J. Rohlfs Department of Medical Biochemistry, The Ohio State University, Columbus, OH 4 3 2 1 0

Introduction Xanthine

of

complex

oxidoreductases possessing redox-active sites in addition to a flavin cofactor.

oxidase and trimethylamine

dehydrogenase

are examples

Xanthine

oxidase from cow's milk is a homodimer of 300,000 Da with each subunit containing a molybdenum center, two 2Fe/2S iron-sulfur centers and FAD (1,2). While a crystal structure for the enzyme is lacking, analysis of dipolar interactions between the several centers indicates that they are 11-20 A apart in the enzyme. Trimethylamine dehydrogenase from the pseudomonad isolate W3A1 possesses a 4Fe/4S iron-sulfur center in addition to a covalently linked 6-cysteinyl-FMN (3); its structure has been determined crystallographically (4) with it being found that the two redox-active centers are in close proximity (ca. 4 A edge-to-edge). Both proteins provide the opportunity to examine the factors that influence electron transfer between a flavin and an iron-sulfur center. W e have examined electron transfer within each of these proteins using a pH-jump protocol, and determined the solvent kinetic isotope effect for the equilibration of an electron between the flavin and an iron-sulfur center.

Results and Discussion The distribution of reducing equivalents within partially reduced xanthine oxidase has been shown to be dependent on the pH (5), and this pH dependence has been utilized to examine electron transfer within the enzyme using a pH jump technique (6,7). When partially reduced xanthine oxidase in dilute buffer at pH 10 is mixed with concentrated buffer at pH 6.0, a single-exponential spectral change is observed that reflects a net redistribution of reducing equivalents away from one of the iron-sulfur centers (presumably that designated Fe/S I on the basis its reduction potential; 5) to the flavin with k ^ s = 173 s _1 (Figure 1A); a pH 6-to-10 jump gives a spectral change in the opposite direction with a rate constant of 395 s-1 (Figure

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

752 IB). When the pH 10-to-6 and pH 6-to-10 jump experiments are repeated in D2O, the observed rate constants for electron transfer are 25 s _1 and 56 s -1 , respectively, indicating a pH-independent solvent kinetic isotope effect of 7.0 ± 0.3 for electron transfer between the flavin and Fe/S I. Proton inventory experiments demonstrate that for the pH jump in either direction, the observed rate constant is linearly dependent on the mole fraction of D2O (Figure 2, filled symbols), indicating that the observed isotope effect arises from the motion of a single solvent-exchangeable proton as the system traverses the transition state. Because the pH jump protocol is an equilibrium perturbation technique, the observed rate constant is the sum of the microscopic rate constants for electron transfer in the forward and reverse directions. With Keq known from the relative reduction potentials of FAD and Fe/S I, it becomes possible to calculate kETforward and kETreverse under each experimental condition. The results of such calculations (7) indicate that the solvent kinetic isotope effect on electron transfer from the flavin (semiquinone) to oxidized Fe/S I is significantly larger than in the reverse direction at both pH/D 6.0 and 10.0. We conclude that electron transfer and deprotonation of FADH- are concomitant in xanthine oxidase, and that the N5-H proton is in motion as the reaction system traverses the electron-transfer transition state. A substantial barrier to electron transfer from FADH- thus arises from the need to cleave the N5-H/D bond, making the neutral semiquinone a less effective electron donor than would otherwise be the case. The tight linkage between electron transfer and deprotonation of the neutral flavin semiquinone presumably arises from the inability of xanthine oxidase to accomodate the anionic flavin semiquinone in the active site.

0.08

0.12

time (sec)

Figure 1. pH/D jump kinetics observed with partially reduced xanthine oxidase. Left, jump from pH/D 10 to 6 (0.01 M CAPS, 0.1 N KC1 pH/D 10 mixed with 0.1 M MES, 0.1 N KC1, pH 6.0); Right, jump from pH/D 6 to 10 (0.01 M MES, 0.1 N KC1 pH/D 10 mixed with 0.1 M CAPS, 0.1 N KC1, pH 6.0). Transients obtained using H 2 0 and D2O are indicated; singleexponential fits are shown using the rate constants indicated.

753 Figure 2. Proton inventories for the pH/D jump experiments with xanthine oxidase and trimethylamine dehydrogenase. The plots are of the ratio of the observed rate constant seen at a given mole fraction D2O that observed in 100% D 2 0 for xanthine oxidase (pH/D 10 to 6, filled circles; pH/D 6 to 10, filled squares) and trimethylamine dehydrogenase (pH/D 10 to 6, open circles; pH/D 6 to 10, open squares). mole fraction D , 0

Trimethylamine dehydrogenase differs from xanthine oxidase in that it is able to accommodate either the anionic or neutral forms of the flavin semiquinone in its active site, depending on the pH (8). In order to ascertain whether this protein behaves similarly to xanthine oxidase from the standpoint of having a prototropic equilibrium tightly coupled to electron transfer, a solvent kinetic isotope effect study like that described above was undertaken. Like xanthine oxidase, the distribution of reducing equivalents within partially reduced trimethylamine dehydrogenase is markedly pH dependent: at pH 10 two-electron reduced enzyme possesses reduced iron-sulfur and cys-FMN ", while at pH 6 the enzyme possesses oxidized iron-sulfur center and cys-FMNH2 (8). When two-electron reduced trimethylamine dehydrogenase in dilute buffer at pH/D 10 is mixed with concentrated buffer at pH/D 6.0, the kinetic transients shown in Figure 3A are observed, giving rate constants for electron reequilibration between the reduced Fe/S center and FMNH- of 500 s"1 and 64 s"1 in H2O and D2O, respectively, and a solvent kinetic isotope effect of 7.8. The pH/D 6-to-10 jump experiment (Figure 3B) gives rate constants of 760 s _1 and 120 s"1 in H 2 0 and D2O, respectively, indicating a solvent kinetic isotope effect of 6.3.

-0.2

< <
iM TPB" was present during the 5 min incubation.

usually sigmoidal, whereas without TPB" they are hyperbolic (Fig. IB).

TPB" does not accelerate

development of the inhibition by long-chain, hydrophobic analogs.

A recent, unexpected observation highlights the dual residue of the binding sites for MPP + analogs (11). At catalytic concentrations TPB" potentiates the inhibition, whereas if present in molar excess over the MPP+ analog TPB" reactivates the inhibited enzyme. Fig. 2A illustrates an experiment in which ETP samples were incubated for 5 min at 30"C with 4'-decyl-MPP+, without TPB". Complete inhibition was reached at about 8 jiM inhibitor (open triangles). On addition of 10 jiM TPB" (a slight excess over the inhibitor), immediate reactivation occurred, the residual inhibition declining to 33% (Fig. 2A, solid triangle). The open circles in Fig. 2A show the same experiment but with 10 jiM TPB' present during the 5 min incubation of the inhibitor with ETP. This gave a biphasic titration curve with a plateau at - 35% inhibition (Fig. 2A, solid circle). These data have been interpreted (11) to suggest that the hydrophilic inhibitor can readily enter the membrane and thus fill both the external, the internal binding sites, the latter being in a hydrophobic environment. Hence, complete inhibition is reached rapidly. If an excess of TPB" over the inhibitor concentration is added subsequently, strong ion-pairing occurs, reducing the concentration of free analog. This, in time, results in dissociation of the 4'-decyl-MPP* from one of the inhibition sites, because TPB' binds the inhibitor more tightly than does the site vacated. In the experiment where 10 >iM TPB' is initially present, only one of the two sites , that with the lower K( , is filled at low inhibitor concentration (2 to 5 JiM), yielding partial inhibition in this range. When the concentration of the inhibitor exceeds that of the TPB", the other site is also occupied and complete inhibition results, but a second addition of TPB" once again dissociates the inhibitor from the weaker binding site.

761

5

10

15

CONCENTRATION. f M

6.0

7.0

8.0

pH

Fig. 2A. Inhibition of NADH oxidase by 4'-decyl-MPP+ and its reversal. ETP (20 jig protein/ml) was incubated at 30"C for 5 min with 4'-decy 1-MPP+in the absence (A) and presence (O) of 10 uM TPB" and then assayed. The dashed arrows indicate the addition of 10 ;jM TPB' and resulting reactivation. Fig. 2B. pH dependence of the inhibition of NADH oxidase activity by MPP* . Activity was measured after 5 min incubation of ETP (20 ^g/ml) with 1 mM MPP+ + 10 pM TPB" ; (O) and ( • ) or indicate two separate experiments.

Further evidence for the dual nature of the inhibition site has come from the studies of the effect of temperature and of pH on the inhibition. Thus, while at pH 7.5 on short incubation with hydrophilic MPP* analogs only partial inhibition is reached in the absence of TPB' , by merely raising the pH to 9.0, suppressing the ionization of a positively charged group, complete inhibition occurs in 5 min (9). The pH dependence of the inhibition with or without TPB- shows 2 inflections (Fig. 2B). The pKa values calculated by Dixon's method (12) indicate an imidazole group (pK, ~ 6.8) and a thiol group (pK, ~ 8.4). The latter may be the thiol titration of which with mercurials results in loss of one of the two binding sites for rotenone (7). Further, just as on isolation of the Complex I preparation (14) from mitochondria one of the rotenone sites is lost, concurrently one of the binding sites of MPP* analogs disappears, yielding incomplete inhibition by all analogs tested and lack of significant potentiation by TPB" .

Acknowledgement This research was supported by NIH Program Project HL-16251, NSF grant DMB-9020015 and the Department of Veterans Affairs.

References 1.

Horgan, D.J., T.P. Singer, and J.E. Casida. 1968. J.Biol. Chem. 243, 834-843.

2.

Palmer, G„ D.J. Horgan, H. Tisdale, T.P. Singer, and H. Beinert. 1968. J. Biol. Chem. 243, 844847.

762 3.

Gutman, M , T.P. Singer, H. Beinert, and J.E. Casida, 1970. Proc. Nat. Acad. Sei. U.S. p67 p hox

an

d Rac-related G-protein, the oxidase activity was assessed by measuring

superoxide generation. In addition, the effect of purified NADPH-cytochrome c reductase on superoxide formation was also investigated in a cell-free assay system.

Results and Discussion

The superoxide-generating activity of the purified cytochrome bsss was measured in a cellfree assay system consisting of arachidonate, GTPyS and cytosol in the presence or absence of phospholipids. Reconstituted assay system composed of flavinated cytochrome

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

764 b558 and either cytosol or purified cytosolic components (p47ph0x> p67 p h ox and Rac-related G-protein) gave a markedly stimulated superoxide generating activity (Table 1). In the presence of a constant amount of either cytosol or recombinat cytosolic proteins, the cytochrome demonstrated a dose-dependent superoxide producing activity. This activity was significantly stimulated upon addition of FAD, PC- and PE-dimyristoyl to the incubation and assay mixtures. In addition, flavinated cytochrome bssg also showed SODinsensitive NADPH-dependent cytochrome c reductase activity and its activity was considerably increased in the presence of FAD in the assay mixture. Half-maximal activity of NADPH concentration in the cell-free reconstituted system consisting of flavinated cytochrome bsss and cytosol was observed at approximately 58.8 |iM which was greater than the value (2.20 |iM) for NADPH of NADPH-cytochrome c reductase purified from neutrophil membrane (Fig. 1). These markedly different Km values do not support the idea that the purified reductase acts as a NADPH dehydrogenase to reduce cytochrome bsss in the respiratory burst oxidase system. The NADPH oxidase activity was not stimulated upon incubation of the purified cytochrome c reductase with either flavinated or flavin-depleted cytochrome bsss in the cell-free system, indicating that

-0.6

-0.4

-0.2

0

0.2

0.4

0.6

0$

1.0

INADPH] (nM)-I

Fig. 1: Concentration Dependence of NADPH on the Activities of Both the Cell-free NADPH Oxidase and Purified NADPH-Cytochrome c Reductase. The cell-free system composed of 0.64 jag of flavinated cytochrome bsss (10.7 pmol heme/15.9 pmol FAD) and 30 ng of cytosol. A double-reciprocal plot relating NADPH concentration to superoxide generation (|i.mol O^/min/mg cytochrome) is shown (open circles, upper scale). The cytochrome c reduction by 0.36 \ig of purified reductase (3.53 pmol FAD/2.85 pmol FMN) was also monitored when the concentration of cytochrome c was held constant at 0.2 mM (closed circles, lower scale).

765 Table 1: Reconstitution of O2 Generating Activity with Purified Cytochrome bsss and NADPH-Cytochrome c Reductase. In one ml of an assay medium, 0.36 \ig of purified reductase, 0.42 |ig of flavinated cytochrome bssg (or 0.42 p.g of flavin-free cytochrome b558*) and 8.6 (ig of cytosol (or 2.5 |Xg p47 pho x P l u s 3.5 ¡lg p67 pho x plus 3.0 ng Rac-x) were added, respectively. Flavin-free cytochrome b558 (*) was prepared by the same purification method as used for flavinated cytochrome bssg except that FAD and phospholipids were not added in its purification process.

Component

Heme

Flavin

NADPH oxidase Activity

(nmol/mg) (nmol/mg) ((imol/min/mg cytochrome) Cytochrome b558

16.7

Cytochrome b558*

16.2

Reductase Cytochrome bsss + Cytosol

0.0

24.8 (FAD) 0.02 (FAD) 9.8(FAD)/8.7(FMN)

0.25 negligible negligible 8.53

Cytochrome bsss* + Reductase

0.05

Cytochrome bsss* + Reductase + Cytosol

0.14

Cytochrome bsss* + Cytosol + 10 (iM FAD

2.65

The oxidase activities observed in a cell-free system consisting of recombinant proteins were lower (50-75%) as compared with those seen in the system containing cytosol. As mentioned earlier, cytochrome bsss purified in the absence of FAD was a flavindepleted form, suggesting that binding to the cofactor is weak. The fluorescence emission of FAD incubated with the flavin-free cytochrome is similar to that of free FAD, which peaked around 525 nm when excited at 450 nm. With few exceptions, most flavoproteins have tightly bound FAD, FMN or riboflavin. A possible reason for weak binding of FAD is that the flavin binding pocket of cytochrome b55g provides fewer stabilizing interactions between the protein and its cofactor. Since the extent of fluorescence quenching at 4 °C was very small in a titration of flavin-depleted cytochrome bsss (0.12 |iM as heme) with the free flavin (0.012-0.184 (iM), Kd for FAD was not determined correctly from this experiment. The Km value for FAD (2.58 |0.M) was derived from a double-reciprocal plot of the initial rate of a cell-free reconstituted NADPH oxidase versus the FAD concentration.

766 The NADPH oxidase activity showed a linear dependence on the FAD concentration except at lower concentrations.

In agreement with Rotrosen

et aL (2), the presence of

phospholipids in the purification process and/or in a cell-free incubation stabilizes a native structure of the cytochrome bsss and contributes the binding of FAD into a flavin-binding site.

Acknowledgements

We thank Dr. David J. Lambeth (Emory University, School of Medicine) for providing the recombinant p47ph0x, p67phox and Rae-related G-protein (Rac-x).

References

1. Haper, A.M., Dunne, M.J., and Segal, A.W. (1984) Biochem.J. 219, 519-527. 2. Rotrosen, D„ Yeung, C.L., Leto, T.L., Malech, H.L., and Kwong, C.H. (1992) Science 256,1459-1462.

FLAVOPROTEINS CONTAINING COVALENTLY-BOUND FLAVINS

Preparation and Properties of Recombinant Corynebacterial Sarcosine Oxidase

Lawrence J. Chlumsky, Lening Zhang and Marilyn S. Jörns Department of Biological Chemistry, Hahnemann University Philadelphia, Pennsylvania 19102, USA

Sarcosine oxidase catalyzes the oxidative demethylation of sarcosine (N-methylglycine) to yield glycine, formaldehyde and hydrogen peroxide.

In the presence of

tetrahydrofolate, 5,10-methylenetetrahydrofolate is formed instead of formaldehyde. The enzyme from Corynebacterium sp. P-l is a heterotetramer («, 100,000; 0,42,000; y, 20,000; 8, 6000), containing both covalent [(8a-Ns-histidyl)FAD, attached to the 0 subunit] and noncovalent (FAD) flavin.

Enzyme expression is induced when

Corynebacterium sp. P-l is grown with sarcosine as source of carbon and energy. The noncovalent flavin accepts electrons from sarcosine which are then transferred in one-electron steps to the covalent flavin where oxygen is reduced to hydrogen peroxide. In addition to heterotetramers, monomelic bacterial sarcosine oxidases (MW - 40,000) have been described which contain only a single covalent flavin (1-4).

Results and Discussion The one-step cloning and overexpression of sarcosine oxidase from Corynebacterium sp. P-l and the purification of recombinant enzyme is described elsewhere in this volume. The purified recombinant enzyme is a heterotetramer containing stoichiometric amounts of covalent and noncovalent flavin, identical to that observed for the natural enzyme produced in Corynebacterium sp. P-l.

The same specific activity is observed for

recombinant and natural enzyme preparations. However, the recombinant enzyme exhibits a pronounced lag in an NADH peroxidase-coupled assay. Also, the spectral properties of the recombinant enzyme differ significantly from the natural enzyme. This difference is due to the fact that a portion of the flavin in the recombinant enzyme is present in a modified form. The calculated absorption spectrum of the modified flavin

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

770

Fig. 1 (left) Spectra observed for recombinant and natural enzymes and the spectrum of the modified flavin in recombinant enzyme, calculated as described in the text. Fig. 2 (right) Aerobic reaction of recombinant enzyme with 2.0 mM sarcosine. 300

400 Wov«Ungth

500 (nm)

400 Wav«l«ngth

500 (nm)

shows a maximum at 383 nm (Fig. 1). The natural enzyme is induced by sarcosine and undergoes multiple turnovers prior to isolation. In contrast, the recombinant enzyme probably experiences minimal turnover during cell growth since expression in E. coli is under the control of the lac promoter. To determine whether this difference might be important, we examined the effect of turnover on the spectral properties of recombinant enzyme. Although recombinant enzyme is readily reduced by 2 mM sarcosine under aerobic conditions, the spectrum of the reoxidized enzyme did not coincide with the initial spectrum but instead closely resembled the spectrum observed for natural enzyme (Fig. 2). To determine whether any of the reaction products might be involved in this phenomenon, we incubated a fresh sample of recombinant enzyme with formaldehyde, glycine or hydrogen peroxide. An effect was observed only with hydrogen peroxide which produced spectral changes similar to that observed after turnover.

When the turnover experiment with

recombinant enzyme and sarcosine was repeated in the presence of excess catalase, we found that catalase interfered but did not completely prevent the spectral change. Therefore, the spectral change observed in the absence of catalase is only partly due to the reaction of the enzyme with free hydrogen peroxide. This suggests the spectral change may also involve reaction with hydrogen peroxide prior to its dissociation from the active site or with a precursor, like a 4a-hydroperoxyflavin. These studies suggested that the modified flavin in recombinant sarcosine oxidase might involve a reversible interaction of normal enzyme flavin with an amino acid residue.

771 0.20

Fig. 3 The reaction of 16 mM sulfite with recombinant or natural (inset) enzyme is shown in panel A. In panel B, the recombinant enzyme was first reacted with 1 mM MMTS and then sulfite was added.

u o •g 0.10 o

I o.os •(TV H 5 0

0.00 100

400 Wavalangth

500 (nm)

«00 300

400

300

Wovalangth

(nm)

600

4a-lhlalata

abduct

Hydrogen peroxide reacts with proteins mainly by oxidizing cysteine and methionine residues. To distinguish between these possibilities, we tested the effect of methyl methanethiosulfonate (MMTS) on recombinant enzyme since this reagent specifically reacts with cysteine residues. The spectral change observed with MMTS is similar to that seen with hydrogen peroxide or after turnover with sarcosine (Fig 3B). It was fully reversed upon treatment with excess dithiothreitol, consistent with the known reversibility of the MMTS reaction with cysteine residues in other proteins. The results show that the modified flavin in recombinant enzyme is due to a reversible interaction of FAD with a cysteine residue. This interaction is disrupted when the cysteine is oxidized with hydrogen peroxide or alkylated with MMTS. As suggested in Scheme I, the interaction with the cysteine residue appears to involve only the enzyme's covalent flavin. Evidence for this conclusion was obtained by studying the reaction of recombinant enzyme with sulfite. Our previous studies with natural enzyme show that sulfite reacts selectively with the covalent flavin to form a reversible N(5)-adduct, accompanied by a bleaching of 50% of the enzyme's absorbance at 450 nm (Fig. 3A, inset). The extent of bleaching observed with recombinant enzyme and sulfite is about half that observed with the natural enzyme (Fig. 3A). However, if the recombinant enzyme is first treated with MMTS, the extent of bleaching observed upon subsequent addition of sulfite is comparable to that observed with the natural enzyme (Fig 3B). The results indicate that about half of the covalent flavin in the isolated recombinant enzyme interacts with a thiol residue. Based on this information we were able to calculate extinction coefficients for the modified flavin in recombinant enzyme (Fig. 1).

The modified flavin in

recombinant sarcosine oxidase exhibits spectral properties (A^^ = 383 nm, e ^ = 7.3 mM"1 cm"1) similar to that observed for known 4a-thiolate flavin adducts observed with

772

lipoamide dehydrogenase (X nm, e^, = 7.0

mM"1

cm"1)

= 380

and mercuric

reductase

( A ^ = 382 nm, e^j = 7.5

mM"1

(5).

cm'1)

The lag in the catalytic assay observed with recombinant enzyme is due to the presence of the 4a-adduct and can be abolished

by

prior

treatment

hydrogen peroxide or MMTS.

with Our

studies with natural enzyme have shown that the rate of initial electron transfer from

sarcosine

noncovalent

to

the

enzyme's

flavin is unaffected by

complexing the enzyme's covalent flavin with sulfite (1,4). Therefore, this step is probably not affected in recombinant enzyme by the presence a 4a-thiolate adduct involving the covalent flavin. In subsequent steps, electrons from the noncovalent flavin are transferred one at a time to the covalent flavin where oxygen is reduced to hydrogen peroxide. The 4a-thiolate adduct will clearly interfere with interflavin electron transfer. The lag in the assay probably reflects adduct dissociation. Reformation of the adduct is blocked by the oxidation of the cysteine residue by hydrogen peroxide or a precursor formed during the reoxidation of the covalent flavin (Scheme I). The results suggest that newly synthesized corynebacterial sarcosine oxidase is present in an inactive form that can be activated by reaction with sarcosine in what appears to be the first example of a posttranslational modification associated with turnover. Complete activation occurs in vivo when enzyme expression in a corynebacterial cell is induced by sarcosine. The opportunity for in vivo activation is diminished when the recombinant enzyme is produced in E. coli under the control of a lac promoter. The sarcosine oxidase operon exhibits considerable economy in gene packaging. There are only 11 bases between the genes for the 0 and ft subunits and the stop codon of the 6 subunit gene overlaps with the start codon of the a subunit gene. The presence of overlapping genes or genes separated by a very short intergenic region has been

773

The Sarcoslne Oxidase Operon from Corynebactarlum sp. P-1

*

IT

a

I I I I M ^

i

associated with translational coupling, a phenomenon where the same ribosome, or component thereof, serves to translate two contiguous genes without being released from the mRNA (6). Potential ribosome binding sites are identifiable 6 to 8 bases upstream of the start of the & and a genes. These sites are positioned such that translation of the upstream gene will terminate within the ribosome binding site of the corresponding downstream gene, a feature required for the coupling effect. The y subunit gene is tentatively positioned downstream of the incompletely sequenced a subunit gene since it was not found in the sequenced region upstream of the 0 subunit gene. The deduced amino acid sequence of the 6 subunit showed no identifiable motifs or homologies with any other known sequences. The f) subunit exhibits a dinucleotide binding motif near the amino terminus (Fig. 4) which satisfies all of the 11 consensus sequence requirements, except for an aspartate in position 1, a feature observed in many other well-documented FAD-binding sites (7-9). A second potential adenylate binding site starts at LySj21 (KVALAGAGHSSVLAELAG(X)nALVSE) but this site is somewhat ambiguous since a glutamate occupies position 7 where the consensus sequence specifies a small hydrophobic residue and serine replaces the third glycine at position 6. The P subunit exhibits greater than 80% identity with peptide fragment data encompassing -43% of the P subunit in a very similar sarcosine oxidase from Corynebacterium sp. U-96 (10,11). A peptide sequence containing the covalent attachment site for flavin in the U-96 enzyme has been identified in the P subunit of our enzyme. Based on this result, Hi%76 is tentatively identified as the covalent flavin attachment site in the 0 subunit of

774 ADP-binding f o l d — Betasub Baclit Bacpat Arthro Strap

1 MADLLPEHPE FLWANPEPKK MST ST HSIKX SP

BetBBUb Baclit Bacpat Arthro Strap

101 FLFSQRgvm .IFTQTJVLV .IFTKTflVLV .IFTKTflVLV .VATLCflGVM

LAHTLGDVRI YGPK.GGSAF FGPK.GE9AF FGPK.GEAPF AGP..PDSRT

Betasub Baclit Bacpat Arthro Strap

201 FLKDGEKYTG FEVTEDLZT. FDISPDSJK. FEIAEDFJK. WEPYRDGyR.

Betaaub Baclit Bacpat Arthro Strap

Betaaub Baclit Bacpat Arthro strep

100 EQfcPEELDYD YEfcEKETHHK YEfcEKETHHK YEfcEKETHHK EEI^RATGRK

GHgi^TAYYl SHflMftAGYYL SMflM&AGYQfc SKgM^AGYYL GHflS^AAHHi

50 AKNHGITHVA AKQGVKTLLV AKQGVKTLLV SKQGVKTLLV SARGARVLGL

SVRRVEANKF VSETHEAANI VAETMEAAKE VAETMEAAKE VSGSLRSATE

NGVDAEWLTP HSLEHELFEG HSLTVDLLEG HSLOVDLLEG WDLAHEMLDA

150 EQVKEVCPII KQLTDRWAGV DEINKRHPGI SEINKRHPGV KEIRRRFPTL

200 NIGDDIRYPV HGftTYQPRAg IAKKDgVAWft FARXANEMgV DIIQNCEZTG EVPDNY EftlFEPNSg VLFSENCIQA YREL&EAHgA TVLTYTPJED TV PEN Y N J I F E P N S g VLFSENCXR& YREL&EARgA KVLTHTRVED TVPENY NftlFEKNSg VLFSEHCIRft YREIAEAHgA KVLTYTPVED APDDOE VftLFEAKAfl LLRPEHHVAft HLQI^TRQgA ELRFBEPJLR

VKITRGTIHA IKJAKGSYT1 IEJANGSYTft IQJAYGSFTft VHXGENTYTA

GKVALAGAGH NKLWSMGAW DKLIVSMGAW SKLIVSMGAH GQLVICPGAH

SSV^AE^AGF NSKJ,. .fcSKL NSKJ,. .I,SKL NSKJj. .fcSKL APQJ,..LADI

250 ELPIQSHPLQ DVEIPLQPYR NLDIPLQPYR NIEIPLQPYR GVP1TVE..R

ALVSELFEPV QWGFFECDE QWGFFESDE QWGFFECDE QIMYWFQPKG

HPTWMSNHI AKYSNNAH.Y SKYSNDID.F KKYSNTHG.Y GTGPFVPERH

HVYVSQAHKg PAFKVEVENfl PGFMVEVPNfl PAFMVEVPTg PVYIWEDADfl

ELVHg I.YYgFPSFG I . YYflFPSFG I.YYgFPSFG VQVYflFPAID

300 AGIOSY GS..GLKIGY GC..GLKLGY GC..GLKIGY GPEKGAKVAF

301 HGYgflR HSYflfiQIDPD HTFflflKIDPD HTYgfiKIDPD FRKgflHTTPE

GAFH TINREFGAYP TIHREFGVYP TIHREFGIYP TIDR..TVHA

VIEEQMAAAV EDEANLRXFL EDESNLRAFL EDEGNIRKFL HEVRAMADHM

ELFPIFARAH EQYHPGANGE EEYHPGANGE ETYMPGATGE SALIPDLPGT

350 VLRTWGGIVD LKKGAVCMYT LKKGAVCMYT LKSGAVCMYT FLKAATCMYS

TIMpASPIIS KJPfiEHFVID KTLfiEKFUD KXPQEHFVID NJPgEHFVIA

KTJ..IQHLY LHJKYSN.VA UI£EHSN.W LHEQFSN.VA RHfAHPESVT

VMCgWGTGgf IAAgFSGHgj IAAgFSGHgjf IAA9FSGH9K VACgFSGHgf

BGTPGAgFTi BFSSWgETi JFSSGVgEVfc IFSSWgETIi EFVPWgEIfc

400 AHTIANDEAH AQLATTGKTE SQLALTGKTE SQLAVTGKTE ADIALTGATA

401 AIHAPKSLEB HDISI£SLNB HDISIZSINft HDISIZSINB HPIGI£DPA£

FETGHLIDEH DALKKEAVK PALXESLQKT PALK...QKE LTAPAARGVQ

SY££VIVSGg HFfiJIWgAg HFBZIWSA2 DYEJIWgAg TYBSIVigLfl

VLEKGWLAGfl DSFDPPHTKg DAFDPPHTNfl DSFHPPHTNfl EKFGPVHNRfl

MMARNTTIIB SHflGDTRIIB SHflGDTBIIB SHgGDTRIIg SSfiGGSRITR t

SNXLWD.ESA HAJGEGREYV HAJGEGREYV HAJGEGREYV QSXFEDPAYV

GIYEKSLKfcW PFALRAQE&H PLALRSQEfcW PFALRAQEJjW PLLLRAYEIiY

dimethylgly. dehydro. flavin beta subunit flavin

427 GAAAVAH TI TI P

Fig. 4 Multiple sequence alignment of the f ) subunit from Corynebacterium sp. P-1 (Betasub) and monomeric sarcosine oxidases from Bacillus sp. NS-129 (Baclit), Bacillus sp. B-0618 (Bacpat), Arthrobacter sp. TE1826 (Arthro) and Streptomyces sp. KB210-8SY (Strep). The alignment was generated using the PILEUP program in the GCG package.

the Corynebacterium sp. P-l enzyme. The p subunit exhibits sequence homology with rat liver dimethylglycine dehydrogenase (12) and with four monomeric bacterial sarcosine oxidases (13-15). As judged by results obtained in pair-wise alignments (Table 1), there is considerable variability in the level of homology within the monomeric sarcosine oxidases themselves; percent identity values vary from 35 to 85% with similarity values ranging from 55 to 90%. The 0 subunit from coiynebacterial sarcosine oxidase shows about 25% identity and 45% similarity with the monomeric sarcosine oxidases, values which are not much different from the lower extreme within the monomerics themselves. Overall, less similarity is observed between the monomeric sarcosine oxidases and dimethylglycine dehydrogenase. However, dimethylglycine dehydrogenase gives a somewhat better fit with the 0 subunit than most of the monomeric sarcosine oxidases. The 0 subunit contains 405 amino acids and is about half the size of dimethylglycine dehydrogenase which contains 857 residues.

775 Comparisons: Corynebacterial Beta Subunit (BetaSub), Dimethylglycine Dehydrogenase (DMQDH) and Various Monomeric Sarcosine Oxidases Beta Sub

DMGDH

Bac Pat

Bac Lit

Strep

Arthro

< < < < < < < < < < Percent Similarity > > > > > > > > > > > BetaSub

-

48.0

44.8

47.4

45.6

45.4

BetaSub

-

44.6

44 JS

44.2

42J

DMGDH

-

88.1

59.8

912

BacPat

-

59.5

89.6

BacLit

-

575

Strep

DMGDH

23.2 (9)

BacPat

223 (9)

202 (12)

BacLit

245 (11)

20.1 (13)

80.4 (0)

Strep

23.4 (14)

23.4 (13)

36.7 (8)

Arthro

22J 18.4 (12) (12)

Arthro

Table 1 Values for percent similarity and identity and number of gaps required for pair-wise alignments were obtained using the GAP program in the GCG package. Abbreviations are defined in the title and in the legend to Fig. 4. As shown in a dot matrix comparison (Fig. 5), the homology with the P subunit involves the amino terminal half of dimethylglycine dehydrogenase.

An unmistakable

dinucleotide binding motif aligns near the amino terminus of all six polypeptides. The known histidine attachment site for the covalent flavin in dimethylglycine dehydrogenase is located 11 amino acids away from the carboxyl end of this motif.

A multiple

sequence alignment of the f) subunit with the monomelic sarcosine oxidases reveals that 42 residues are conserved among these 5 polypeptides (Fig. 4). The covalent flavin attachment site in dimethylglycine dehydrogenase aligns with a histidine residue that is conserved in all 4 monomelic sarcosine oxidases but not in the f) subunit. The putative attachment site for the covalent flavin in the 0 subunit (His^j) is located near the middle of the polypeptide where it aligns with a conserved asparagine in the monomelic enzymes. The results suggest that the N-terminal dinucleotide binding motif in the p subunit may actually represent a part of the binding site for the noncovalent flavin.

776

These observations clearly raise some intriguing questions regarding the role of the other three subunits in the corynebacterial enzyme.

Beta Subunit

Fig.5

Sarcosine shares certain metabolic similarities with other secondary and tertiary amino acids, like dimethylglycine, proline and pipecolic acid. Oxidative reactions assume critical importance in the initial catabolism of these amino acids because the presence of a substituted «-amino

group precludes the metabolic diversity enjoyed by other amino acids in reactions Table 2 The Big Picture Subatrate

Quaternary Structure

Subunit Sao

Yes

Hetero-

96-100 42-tSuM•l. Na> 20-23 6-14

Yaa (ß aibuoit)

Yea

Moaomar

42-45

Yea

Saicoaiae Defcydrojenaae*

Liver Yea (mitochondria)

Monomer

94

Probable

Dimetbytglycine Dehydrogenase*

Liver Yea (mitochondria)

Monomer

96

Yea

PutA Prouia

£«*, No £ typhimuHum

Ha•odimer

132

No

Praline Dehydrogenase

Yeaat Liver Droaophüa (müodMüdria)

1

Hoinnmrr

53-75

No

Pipecolic acid Ondue

Liver (peranum«)

Ya

Enzyme

Source

Saroonne Ondue*

Frakaiyotea (cytoaoi)

Sareoone Oxidaae

Prokaiyotee (cytoaci)

Covmlatf Flavin

Sfqiwuv Homology

(kDa)

X/^eooH

CH,

Q H

Tecrahydrofolate actf as a «oaubatrate k time reaction.

46

Ill

involving pyridoxal phosphate-dependent enzymes. A survey of enzymes known to catalyze oxidative reactions analogous to the sarcosine oxidase reaction is shown in Table 2. Tetrahydrofolate acts as a cosubstrate in all reactions which involve Nmethylated amino acid substrates (2). Although covalently bound flavin is relatively uncommon among flavoenzymes, it is strikingly prevalent among the enzymes listed in Table 2, with the notable exception of reactions involving proline as substrate. Mitochondrial sarcosine dehydrogenase has not yet been cloned but the enzyme exhibits many similarities with dimethylglycine dehydrogenase, including 64% identity in the amino acid sequence near the covalent flavin attachment site, a feature which has previously been observed only for enzymes which exhibit overall sequence homology, like fumarate reductase and succinate dehydrogenase (16). Recent unpublished data obtained by Dr. Stephanie Mihalik show that human pipecolic acid oxidase (17) exhibits about 35% sequence homology with the monomelic sarcosine oxidase from Bacillus sp. NS-129. In contrast, no significant homologies were detectable in a series of pair-wise comparisons involving the f) subunit from corynebacterial sarcosine oxidase, proline dehydrogenase from yeast and Drosophila (18,19) and the PutA protein from Salmonella typhimurium (20), not even between the two eukaiyotic proline dehydrogenase. The results suggest that sarcosine oxidase and the other covalent-flavin containing enzymes listed in Table 2 may constitute a new family of homologous proteins.

Acknowledgements This work was supported by NIH Grant GM 31704. We thank Dr. Stephanie Mihalik (Johns Hopkins University) for permission to cite her unpublished data.

References 1. Zeller, H.D., R. Hille and M.S. Jorns. 1989. Biochemistry 28, 5145-5154. 2.

Kvalnes-Krick, K. and M.S. Jorns. 1987. Biochemistry 26, 7391-7395.

3. Kvalnes-Krick, K. and M.S. Jorns. 1991. In: Chemistry and Biochemistry of Flavoenzymes, Vol. II (Muller, F., ed.). CRC Press, Inc., pp. 425-435.

778

4.

Ali, S.N., H.D. Zeller, M.K. Calisto and M.S. Jörns. 1991. Biochemistry. 30, 10980-10986.

5.

Williams, C.H. 1992. In: Chemistry and Biochemistry of Flavoenzymes, Vol. III Müller, F., ed.). CRC Press, Boca Raton, pp. 121-211.

6.

Schumperli, D., K. McKenney, D.A. Sobieski and M. Rosenberg. 1982. Cell 30, 865-871.

7.

Wierenga, R.K., P. Terpstra and W.GJ Hol. 1986. J. Mol. Biol. 187, 101-107.

8.

Mckie, J.H. and K.T. Douglas. 1991. FEBS. Lett. 279, 5-8.

9.

Vanbeeumen, JJ., H. Demol, B. Samyn, R.G. Bartsch, T.E. Meyer, M.M. Dolata and M.A. Cusanovich. 1991. J. Biol. Chem. 266, 12921-12931.

10. Suzuki, H. and Y. Kawamura-Konishi. 1991. J. Biochem. Tokyo. 109, 909-917. 11. Suzuki, H. and Y. Kawamura-Konishi. 1988. Biochem. Int. 17, 577-583. 12. Lang, H., M. Polster and R. Brandsch. 1991. Eur. J. Biochem. 198, 793-799. 13. Nishiya, Y. and T. Imanaka. 1993. J. Ferm. Bioe. 75, 239-244. 14. Suzuki, K, M. Ogishima, M. Sugiyama, Y. Inouye, S. Nakamura and S. Imamura. 1992. Biosci. Biotech. Biochem. 56, 432-436. 15. Koyama, Y., H. Yamamoto-Otake, M. Suzuki and E. Nakano. 1991. Agric. Biol. Chem. 55, 1259-1293. 16. Cook, RJ., K.S. Misono and C. Wagner. 1985. J. Biol. Chem. 260, 12998-13002. 17. Mihalik, SJ., M. Mcguinness and P.A. Watkins. 1991. J. Biol. Chem. 266,4822-4830. 18. Wang, S.S. and MJ. Brandriss. 1987. Mol. Cell Biol. 7, 4431-4440. 19. Adams, E. and L. Frank. 1980. Ann. Rev. Biochem. 49, 1005-1061. 20. Allen, S.W., A. Sentiwillis and S.R. Maloy. 1993. Nucleic Acids.Res. 21,1676-1676.

One-Step Cloning and Overexpression of the Sarcosine Oxidase Operon from Corynebacterium sp. P-l

Lawrence J. Chlumsky, Andrew J. Ramsey, and Marilyn S. Jorns Hahnemann University School of Medicine, Philadelphia, PA 19102, USA

Introduction Sarcosine oxidase is induced when Corynebacterium sp. P-l is grown with sarcosine as source of carbon and energy. Under these conditions, the enzyme constitutes ~3% of total soluble protein (1). The enzyme catalyzes the oxidative demethylation of sarcosine to yield glycine, formaldehyde and hydrogen peroxide (eq. 1). CH 3 N + H 2 C h 2C0 2 " +

+ H20

H 3 N + CH 2 C02" + H 2 CO + H 2 0 2

(eq. 1)

The corynebacterial enzyme is a heterotetramer containing equimolar amounts of covalent and noncovalent FAD, a single site for sarcosine and two sites for tetrahydrofolate. The covalent flavin is attached to the P subunit (1). The noncovalent flavin accepts electrons from sarcosine. These electrons are then transferred in one-electron steps to the covalent flavin, which then reduces oxygen to hydrogen peroxide (1-4). Corynebacterial sarcosine oxidase is similar to several other enzymes, including NADPH cytochrome P450 reductase, that have two flavins (FAD, FMN) per active site with different roles in catalysis; however, both flavins in these enzymes are noncovalently bound (5). The complex quaternary structure and multiple binding sites for substrate and prosthetic groups in corynebacterial sarcosine oxidase present an intriguing target for structure-function studies. In this chapter, we describe the isolation and overexpression of the operon encoding the sarcosine oxidase from Corynebacterium sp. P-l and the purification of the recombinant enzyme.

Results and Discussion Based on observations for other multisubunit bacterial enzymes (6-8), we assumed that the genes coding for the subunits of sarcosine oxidase were likely to be organized in an operon with a coding region encompassing approximately 5000 base pairs (bp). The genes comprising the sarcosine oxidase operon from Corynebacterium sp. P-l were isolated and

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

780 expressed in a single step. This was accomplished by insertion of size-fractionated genomic DNA (5.0 - 9.4 kbp), previously subjected to partial digestion by Sau3AI, into pBluescript n SK(+), an Escherichia coli vector capable of maintaining a DNA fragment large enough to code for the putative sarcosine oxidase operon. Transformants expressing the sarcosine oxidase genes were identified on indicator plates containing sarcosine, o-dianisidine and horseradish peroxidase (9). These plates were designed to screen a genomic library for colonies that generate hydrogen peroxide in a sarcosine-dependent reaction. The original isolate (pLJC305) contained a ~7300 bp corynebacterial genomic DNA insert (Figure 1). The expression of the sarcosine oxidase genes in E. coli strain XL-1 Blue/pLJC305 was shown to be under the control of the vector-encoded lac promoter, rather than the original corynebacterial promoter by (a) the loss of enzyme expression when the orientation of the genes was reversed relative to the lac promoter and (b) an increase in the level of expression of the enzyme in the original isolate in the presence of inducer (IPTG). Expression of sarcosine oxidase activity was still observed when ~1 kbp (pLJC306), but not ~4 kbp (pLJC305AClaI), of DNA was removed from the 3' end of the 7.3 kbp insert in pLJC305 (Figure 1). Recombinant sarcosine oxidase accounts for as much as -27% of the soluble cell protein when strain XL-1 Blue/pLJC305 is induced with IPTG. The enzyme was purified from XL-1 Blue/pLJC305 cultures incubated at 30 °C for 12-16 hours after induction with IPTG. The harvested cells were readily disrupted by sonication. The cell lysate was then subjected to ammonium sulfate fractionation, followed by successive column chromatography over Ultrogel ACA 34, Phenyl-Sepharose and DEAE-Sephacel. The enzyme underwent a ~ 14-fold purification with a yield of -53% (Table 1) and was >99% pure as judged by native gel electrophoresis. A total of -0.5 grams of purified enzyme was obtained from - 4 0 grams of cells. The recombinant enzyme is isolated as a heterotetramer containing equimolar amounts of covalent and noncovalent flavin, with the covalent flavin attached to the (3 subunit, identical to the enzyme isolated from Corynebacterium sp. P-l. The ability of E. coli to efficiently flavinylate the p subunit and to assemble these four foreign polypeptides into a functional enzyme is impressive, especially since in vitro refolding experiments with this enzyme have thus far been unsuccessful (Zhang and Joms, unpublished results). A description of the organization of the sarcosine oxidase operon, as well as the spectral and catalytic properties of the purified enzyme, is presented elsewhere in this volume.

782

Table 1

Purification of Recombinant Corynebacterial Sarcosine Oxidase Expressed in E. coli. Total Activity (Units)a

Protein (mg)

Specific Activity (units/mg)

cell lysate''

12600

14800

(NH4)2S04 ppt

9200

11150

Ultrogel ACA 34

7110

Phenyl-Sepharose

7050

DEAE-Sephacel

6670

Step

Yield

%

x-fold Purified

0.85

100

1.0

1.08

73.0

1.3

1340

5.32

56.4

6.3

788

8.95

56.0

10.6

545

12.2

52.9

14.4

"Units = (imol fonnaldehyde/min. 'The cell lysate was prepared from 39.1 g (wet weight) of E. coli (XL-1 Blue/pLJC305).

Acknowledgement This work was supported by NEH Grant GM 31704 (M. S. J.).

References 1. Kvalnes-Krick, K. and M. S. Jörns. 1986. Biochemistry 25, 6061-6069. 2. Jörns, M. S. 1985. Biochemistry 24, 3189-3194. 3. Zeller, H. D., R. Hille and M. S. Jörns. 1989. Biochemistry 28, 5145-5154. 4. Ali, S. N., H. D. Zeller, M. K. Calisto and M. S. Jörns. 1991. Biochemistry 30, 10980-10986. 5. Kvalnes-Krick, K. and M. S. Jörns. 1991. In: Chemistry and Biochemistry of Flavoenzymes, Vol. 2 (F. Muller, ed.). CRC Press, Boca Raton, pp. 425-435. 6. Stewart, V. 1988. Microbiol. Rev. 52,190-232. 7. Cole, S. T., C. Condon, B. D. Lemire and J. H. Weiner. 1985. Biochim. Biophys. Acta 811, 381-403. 8. Wood, D„ M. G. Darlison, R. J. Wilde and J. R. Guest. 1984. Biochem. J. 222, 519534. 9. Sagai, H., H. Masujima, S. Ikuta, and K. Suzuki, inventors; Toyo Jozo Co., Ltd., assignee. 1989. French patent. No de publication 2619395 17 February 1989, 43 p., Int. CI4 C12N15/00; Appl. 1988 August 10.

L-GULONO-Y-LACTONE OXIDASE — cDNA CLONING AND ELUCIDATION OF T H E G E N E T I C D E F E C T IN A M U T A N T R A T W I T H O S T E O G E N I C

DISORDER

K. Yagi, M. Nishikimi Institute of Applied Biochemistry, Yagi Memorial Park, Mitake, Gifu 505-01, Japan

Introduction L-Gulono-7-lactone oxidase (EC 1.1.3.8, GLO) is the enzyme that catalyzes the terminal step of the pathway of L-ascorbic acid biosynthesis and is located in the endoplasmic reticulum of the liver and/or kidney cells of most vertebrates. Interestingly, humans, monkeys, and guinea pigs are deficient in GLO (1) and, as a consequence, suffer from scurvy when their dietary intake of vitamin C is not sufficient. Another interesting feature of this enzyme is that it possesses FAD that is covalently bound through its 8a methyl group to the N(l) position of a histidyl residue of the apoprotein (2, 3). To our knowledge, this kind of covalently-bound flavin is unique among flavoproteins of eukaryotic species, although it is known to occur in a few enzymes of prokaryotes (4). As an initial step to elucidate the structure-function relationship of this flavoenzyme, we cloned a cDNA for rat GLO, and investigated the genetic defect of a mutant rat that is known to lack GLO like the above-mentioned species. In this article, we will summarize the results obtained in these studies.

Cloning of cDNA Encoding Rat GLO A rat liver cDNA library in an expression vector, Xgtl 1, was screened by use of anti-rat GLO rabbit antibody, a cDNA encoding the entire amino acid-coding region was isolated, and its nucleotide sequence was determined (5). Figure 1 shows the sequence data of the cloned cDNA. The authenticity of the clone was confirmed by the following observations: 1) The, amino-terminal sequence of 33 amino acids that had been chemically determined with purified rat GLO completely matched the amino acid sequence deduced from the nucleotide sequence. 2) The composition of amino acids determined by chemical analysis of rat GLO was in reasonable agreement with that calculated from the deduced amino acid sequence. 3) Catalytically-active GLO protein could be expressed from the cDNA that had been inserted into the eukaryotic expression vector pS VL (6), as detailed below.

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

784 GATCCTCCTGATCACTGGAATC

-1

ATGGTCCATGGGTACAAAGGGGTCCAGTTCCAAAATTGGGCAAAGACCTATGGTTGCAGTCCAGAGGTGTACTAC MetValHlsGlvTvrLvsGlvValGlnPheGlnAsnTroAlaLvsThrTvrGI vCvsSerProGluValTvrTvr

75

CAGCCCACCTCCGTGGAGGAGGTCAGAGAGGTGCTGGCCCTGGCCCGGGAGCAGAAGAAGAAAGTGAAGGTGGTG GlnProThrSerValGluGluValAraGluValLeuAlaLeuAlaAraGluGlnLYsLysLysValLvsValVal

150

GGTGGTGGCCACTCGCCTTCAGACATTGCCTGCACTGACGGTTTCATGATCCACATGGGCAAGATGAACCGGGTT GlyGlyGlyHlsSerProSecAspIleAlaCysThrAspGlyPheMetlleHlsMetGlyLysMetAsnArgVal

225

CTCCAGGTGGACAAGGAGAAGAAGCAGATAACAGTGGAAGCCGGTATCCTCCTGGCTGACCTGCACCCACAGCTG LeuGlnValAspLysGluLysLysGlnlleThrValGluAlaGlylleLeuLeuAlaAspLeuHlsProGlnLeu

300

GATGAGCATGGCCTGGCCATGTCCAATCTGGGAGCAGTGTCTGATGTGACAGTTGCTGGTGTCATTGGATCCGGA AspGluHisGlyLeuAlaMetSerAsnLeuGlyAlaValSerAspValThrValAlaGlyVallleGlySerGly

375

ACACATAACACAGGGATCAAGCACGGCATCCTGGCCACTCAGGTGGTGGCCCTGACCCTGATGACAGCTGATGGA ThrHisAsnThrGlylleLysHlsGlylleLeuAlaThrGlnValValAlaLeuThrLeuMetThrAlaAspGly

450

GAAGTTCTGGAATGTTCTGAGTCAAGAAATGCAGATGTGTTCCAGGCTGCACGGGTGCACCTGGGTTGCCTGGGC GluValLeuGluCysSerGluSerArgAsnAlaAspValPheGlnAlaAlaArgValHIsLeuGlyCysLeuGly

525

ATCATCCTCACCGTCACCCTGCAGTGTGTGCCTCAGTTTCACCTTCAGGAGACATCCTTCCCTTCGACCCTCAAA IlelleLeuThrValThrLeuGlnCysValProGlnPheHisLeuGlnGluThrSerPheProSerThrLeuLys

600

GAGGTCCTTGACAACCTAGACAGCCACCTGAAGAGGTCTGAGTACTTCCGCTTCCTCTGGTTTCCTCACACTGAG GluValLeuAspAsnLeuAspSerHlsLeuLysArgSerGluTyrPheArgPheLeuTrpPheProHisThrGlu

675

AACGTCAGCATCATCTACCAAGACCACACCAACAAGGCCCCCTCCTCTGCATCTAACTGGTTTTGGGACTATGCC AsnValSerllelleTyrGlnAspHisThrAsnLysAlaProSerSerAlaSerAsnTrpPheTrpAspTyrAla

750

ATCGGGTTCTACCTACTGGAGTTCTTGCTCTGGACCAGCACCTACCTGCCATGCCTCGTGGGCTGGATCAACCGC IleGlyPheTyrLeuLeuGluPheLeuLeuTrpThrSerThrTyrLeuProCysLeuValGlyTrpIleAsnArg

825

TTCTTCTTCTGGATGCTGTTCAACTGCAAGAAGGAGAGCAGCAACCTCAGTCACAAGATCTTCACCTACGAGTGT PhePhePheTrpMetLeuPheAsnCysLysLysGluSerSerAsnLeuSerHlsLysIlePheThrTyrGluCys

900

CGCTTCAAGCAGCATGTACAAGACTGGGCCATCCCTAGGGAGAAGACCAAGGAGGCCCTACTGGAGCTAAAGGCC ArgPheLysGlnHisValGlnAspTrpAlalleProArgGluLysThrLysGluAlaLeuLeuGluLeuLysAla

975

ATGCTGGAGGCCCACCCCAAAGTGGTAGCCCACTACCCCGTAGAGGTGCGCTTCACCCGAGGCGATGACATTCTG MetLeuGluAlaHlsProLysvalvalAlaHlsTyrProValGluValArgPheThrArgGlyAspAspIleLeu

1050

CTGAGCCCCTGCTTCCAGAGGGACAGCTGCTACATGAACATCATTATGTACAGGCCCTATGGAAAGGACGTGCCT LeuSerProCysPheGlnArgAspSerCysTyrMetAsnllelleMetTyrArgProTyrGlyLysAspValPro

1125

CGGCTAGACTACTGGCTGGCCTATGAGACCATCATGAAGAAGTTTGGAGGAAGACCCCACTGGGCAAAGGCCCAC ArgLeuAspTyrTrpLeuAlaTyrGluThrlleMetLysLysPheGlyGlyArgProHlsTrpAlaLysAlaHls

1200

AATTGCACCCAGAAGGACTTTGAGGAAATGTACCCCACCTTTCACAAGTTCTGTGACATCCGTGAGAAGCTGGAC AsnCysThrGlnLysAspPheGluGluMetTyrProThrPheHisLysPheCysAspIleArgGluLysLeuAsp

1275

CCCACTGGAATGTTCTTGAATTCGTACCTGGAGAAAGTCTTCTACTAAAGCAGGAGTGGAAACAAACCACCCTGA ProThrGlyMetPheLeuAsnSerTyrLeuGluLysValPheTyr***

1350

CCCCTCACACTTCTGCTGCCCCCGGGGGTCTGGGGAGCAGAGAAGTGCCTCACAAGCACAATGGGAACTGACCTC

1425

TCCTCCTGACCACAAAGAAAGGCTGGGCTCTGGGCGGGTCCTCTCTGCCTl'CGGCATCATTTCCCTTACATCCAG

1500

GCAAGAAGTGGCCTCTCACTCAAATTCCTGTTAGCATTTCCATGGGTCACACATAAACTGCAATCCTCTCAGGAG

1575

AAGGGGGATCCCTGATACATCATATCTATCCAGACTAAGGATGTGGTTCTTCCTAGATTCTATGGCTCCACCAGG

1650

TATAGAGAGATTCCTGGGGCCTGCAGTTCTCCATCCCTCTTCAGAAGGGAGGGATCCCTTGGCGAGAGTTTGGCT

1725

CAGAGGTGGCATGAAGCATGCTCTGCTCTCTCTTACCCTTGAAGGTCCTTCGGATGCCCAGAGATGTCTGCTGGT

1800

CCTGGGCAAGCCATCATTCAAACGGGTCCAACCTGGCCTTCTGTCTGCCATGGCCTGACCCTCGCAGTGTCTCTT

1875

CCAGAGGTGTTTAGAGTGGAACTCGCTTCAACCTCTTAACCAGTTGCTGATCCCTGTGTTTCTCTCCCTTCTCCT

1950

TGGAGACTACTCTTGGAGGGGGATCCCACCATGTCCTTGGCTTTCCCTGGGTATTGTTCTCCTCTTCCTCTTCAC

2025

AAATATGATTTCAGTTTGATTTGTGGCCTTTCTGGAGTGTTCCTTGGAGAACCAAGATGTTCCAGCTACCGAATT

2100

C

2101

Fig. 1. Nucleotide sequence and deduced amino acid sequence of rat liver GLO cDNA. The amino acid residues that were confirmed by amino acid sequencing of purified GLO are underlined. The numbering begins at the proposed initiator methionine residue. From Ref. 5.

785 In the determined nucleotide sequence, there is an open reading frame of 1320 nucleotides, which encodes a protein of 440 amino acids. The amino-terminal methionine is cleaved off in mature rat liver GLO, and the resulting polypeptide consists of 439 amino acids with a molecular weight of 50,483.

Thus the molecular weight of the holoenzyme having an

covalently-bound FAD is presumed to be 51,267, which is comparable to that of rat GLO determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (7). Hydropathy analysis by the method of Kyte and Doolittle (8) indicated the presence of several strongly hydrophobic regions, some of which may interact with the microsomal membrane. Some of the hydrophilic regions existing in the sequence must be exposed at the outside of the membrane, because anti-GLO antibody was shown to bind to intact microsomes of rat liver. Expression of GLO in Monkey Cells We constructed a minigene by inserting the 1.6-kilobase pair Bcfl-BamHl portion of the rat GLO cDNA into a eukaryotic expression vector, pS VL, and attempted to express GLO in cells of the COS-1 monkey cell line (6). The structure of the resulting construct is shown in Fig. 2A. When COS-1 cells were transfected with this minigene and tested for expression of GLO by an immunohistochemical technique, 5-10% of the cells were found to produce GLO protein (Fig. 2B). Its molecular weight was the same as that of rat liver GLO, as shown by Western blot analysis. The GLO expressed in the transfected cells was enzymatically active, and the GLO activity reached maximum at 72 h of culture.

In a cell fractionation experiment, the

A

B

• K

SV40 late

SV40 late promoter SV40 ori \

VP1 intron

polyadenylatiori site R

E

\

B /

H-Uk Amp r

pBR322 ori

V

^ I

.

M

&

* ^ ^

Fig. 2. Expression of GLO in COS-1 cells transfected with a rat GLO minigene. A, structure of the GLO minigene used. The 1.6-kilobase pair Bcll-BamUl portion of the rat GLO cDNA was placed downstream of the SV40 late promoter in the eukaryotic expression vector pSVL. The part of the c D N A is represented by the solid box. B, histochemical demonstration of expression of GLO. COS-1 cells were transfected with the construct shown in A, and the expressed protein was immunostained after incubation for 72 h. From Ref. 6.

786

Fig. 3. Gross features of ODS rats. Left, normal (ODS-+/+); right, mutant Shortness of the legs is seen in the mutant.

(ODS-od/od).

highest activity was found in the 100,000 x g pellet, indicating that the expressed GLO resided in the endoplasmic reticulum of the cell. The specific activity in the microsomal fraction (0.38 nmol/min/mg protein) was - 5 % of that in rat liver microsomes [7.1 nmol/min/mg protein (9)]. Considering that the population of the cells expressing GLO was 5-10%, this result implies that the specific activity of GLO in the microsomes of cells is comparable to that in rat liver microsomes. To our knowledge, this is the first instance of the expression of a recombinant flavoprotein possessing covalently-bound flavin in cells of a mammalian species. Elucidation of Genetic Defect in a Mutant Rat with GLO Deficiency Nearly a decade ago, Mizushima et al. (10) established a mutant strain of the Wistar rat with a hereditary osteogenic disorder (Fig. 3) and termed it the ODS rat. Since this disorder could be cured by administration of L-ascorbic acid, an abnormality of L-ascorbic acid biosynthesis was suspected and eventually traced to a deficiency of GLO in the liver. They showed by a colorimetric assay method that the ODS rat has no GLO activity.

We examined by

immunochemical techniques whether the ODS rat has immunologically cross-reacting material related to GLO (11). A dot-immunobinding assay indicated that liver microsomes of normal (ODS-+/+) rats gave a positive signal with 16 ng of microsomal protein, whereas those of ODS (ODS-od/od) rats gave no signal even with 1 ng of microsomal protein, indicating that the amount of immunologically detectable protein in liver microsomes of ODS rats is, if it is present at all, less than 1/64 of that present in liver microsomes of normal rats.

The

microsomes of the heterozygote (ODS-+/od rat) contained about half the amount of the protein

787 present in normal rat liver microsomes. The results of the measurement of the enzymatic activity and the immunochemical analysis pointed to the conclusion that the mutation in the ODS rat produces a 'null' phenotype for GLO deficiency. But this conclusion became doutbful in a strict sense, when we performed an in vitro translation experiment with poly(A) + RNA from ODS rat liver and found that a detectable amount of GLO-specific protein was produced with a reticulocyte lysate cell-free system, although it was considerably less than the amount produced with poly(A) + RNA from normal rat liver (9). Knowing this result and expecting that GLO may exist at a very low level in the liver of the ODS rat, we measured GLO activity again by a more sensitive method using highperformance liquid chromatography for determination of L-ascorbic acid (12), and found that there existed a marginal level of GLO in ODS rat liver microsomes. The level of activity was approximately 1/200 of that in normal ones, i.e., 0.037 ± 0.010 vs. 7.12 ± 1.15 nmol/min/mg protein (9). Moreover, Northern blot analysis showed that the liver of the ODS-od/od rat contained GLO-specific mRNA of the same size as that of the ODS-+/+ rat and that the levels of the mRNAs detected in both genotypes were comparable to each other (9). The liver of the ODS-+/od rat also contained a comparable level of GLO-specific mRNA. Thus it is clear that the mutation in the ODS-od/od GLO gene does not affect the transcriptional efficiency,

Fig. 4. Determination of a mutation in the GLO gene of the ODS-odllod rat. GLO cDNA was synthesized from ODS-+/+ or -od/od rat liver poly(A) + RNA and amplified by the reverse transcriptase-polymerase chain reaction, subcloned, and sequenced. The sequences from nuclecotide 172 (bottom) to 192 (top) are shown. Arrow denotes the position of nucleotide 182, where the mutation occurred. From Ref. 13.

788 the processing of the primary transcript, and the stability of the mRNA. These results also indicated that the defect in the GLO gene of the ODS-od/od

rat is a point mutation or other

minor alteration of nucleotide sequence in its exon regions, because the sizes of both the mutant mRNA and the mutant translation product are the same as those of normal rats. To elucidate the genetic defect in the mutant GLO gene at the nucleotide level, we carried out the reverse transcriptase-polymerase chain reaction with poly(A) + RNAs from livers of ODS+/+ and -odlod rats and determined the nucleotide sequences of the polymerase chain reactionamplified cDNAs for the GLOs of the respective rats (13). Comparison of the two sequences revealed that the mutant GLO cDNA has a single base mutation from G to A at nucleotide 182, which mutation alters the 61st amino acid residue from Cys to Tyr (Fig. 4). 1

2

3

4

A

Fig. 5. Western blot analysis of GLO in COS-1 cells transfected with a minigene encoding GLO of ODS-+/+ or -odlod rat. Total extract (60 ng) from transfected or non-transfected COS-1 cells, or normal rat liver microsomes (10 |ig) was subjected to Western blot analysis. A, the blotted membrane was imaged by chemiluminescence with an X-ray film. B, for densitometry, the film was scanned with a chromatoscanner. Lane 1, extract from nontransfected COS-1 cells; lane 2, extract from the ODS-od/od GLO minigene-transfected COS1 cells; lane 3, extract from the ODS-+/+ GLO minigene-transfected COS-1 cells; lane 4, rat liver microsomes. From Ref. 13.

789 A transient expression experiment was done to examine whether the determined mutation actually caused a deficiency in GLO (13). We constructed minigenes containing the entire amino acid-coding region of the GLO cDNA of the ODS-+/+ rat or that of the ODS-od/od rat and used them to transfect COS-1 cells. Northern blot analysis indicated that the amounts of GLO-specific mRNA produced with both minigenes were comparable to each other, as would be expected from the fact that the expression of GLO is controlled by the same promoter. However, the amount of immunologically detectable protein produced in the ODS-od/od GLO minigene-transfected cells was decreased to about one-tenth of the normal level, as demonstrated by Western blot analysis (Fig. 5). Measurement of GLO activity showed that the level of the enzymatic activity in the ODS-od/od GLO minigene-transfected cells was onefourteenth that in the cells transfected with the ODS-+/+ GLO minigene. Thus it is clear that the Cys->Tyr substitution leads to almost the same degree of reduction in both levels of GLO protein and enzymatic activity. This situation is similar overall to what was observed for the liver of the ODS rat. Since the amount of GLO-specific mRNA was not affected in either case, the molecular mechanism for the reduced GLO expression should be a low translation^ efficiency or instability of the mutant GLO protein. The former possibility may be less likely, since the position of the point mutation (at nucleotide 182) is considerably away from the mRNA leader sequence, the part of mRNA that is generally known to modulate the level of translational initiation (14).

References 1. Burns, J. J. 1959. Am. J. Med. 26, 740-748 2. Kenney, W. C., D. E. Edmondson, T. P. Singer, H. Nakagawa, A. Asano and R. Sato. 1976. Biochem. Biophys. Res. Commun. 71, 1194-1200 3. Kiuchi, K„ M. Nishikimi and K. Yagi. 1982. Biochemistry 21, 5076-5082 4. Singer, T. P. and W. S. Mclntire. 1984. Methods Enzymol. 106, 369-378 5. Koshizaka, T„ M. Nishikimi, T. Ozawa and K. Yagi. 1988. J. Biol. Chem. 263, 16191621 6. Yagi, K., T. Koshizaka, M. Kito, T. Ozawa and M. Nishikimi. 1991. Biochem. Biophys. Res. Commun. 177, 659-663 7. Nishikimi, M., B. Tolbert and S. Udenfriend. 1976. Arch. Biochem. Biophys. 175, 427-435 8. Kyte, J. and R. F. Doolittle. 1982. J. Mol. Biol. 157, 105-132

9. Nishikimi, M., T. Koshizaka, K. Kondo, T. Ozawa and K. Yagi. 1989. Experientia 45, 126-129 10. Mizushima, Y., T. Hirauchi, T. Yoshizaki and S. Makino. 1984. Experientia 40, 359361 11. Nishikimi, M., T. Koshizaka, H. Mochizuki, H. Iwata, S. Makino, Y. Hayashi, T. Ozawa and K. Yagi. 1988. Biochem. Int. 16, 615-621 12. Kito, M., N. Ohishi and K. Yagi. 1991. Biochem. Int. 24, 131-135 13. Kawai, T., M. Nishikimi, T. Ozawa and K. Yagi. 1992. J. Biol. Chem. 267, 2197321976 14. Kozak, M. 1991. J. Biol. Chem. 266, 19867-19870

HIGH-LEVEL EXPRESSION OF RAT L-GULONO-y-LACTONE OXIDASE IN SILKWORM CELLS WITH A BACULOVIRUS VECTOR

M. Nishikimi, K. Yagi Institute of Applied Biochemistry, Yagi Memorial Park, Mitake, Gifu 505-01, Japan J. Kobayashi Department of Chemistry for Materials, Faculty of Engineering, Mie University, Tsu 514, Japan

Introduction L-Gulono-y-lactone oxidase (EC 1.1.3.8, GLO) is a microsomal enzyme that catalyzes the terminal step of L-ascorbic acid biosynthesis in phylogenetically higher animals, and possesses the coenzyme FAD covalently bound through its 8 a methyl group to the N ( l ) position of a histidyl residue of the apoprotein (1,2). We previously isolated a cDNA clone encoding rat liver GLO (3), and attempted to express this enzyme by transfecting COS-1 cells with a minigene constructed from the cDNA and a eukaryotic expression vector, pSVL (4). However, the expressed GLO level was not very high, although the expressed enzyme was catalytically active.

In the present study, we were successful in achieving high-level

expression of GLO by placing its cDNA in a baculovirus vector. In addition we could obtain production of its apoprotein by expressing GLO under riboflavin-deficient conditions.

Results and Discussion We integrated the 1.6-kilobase pair Bcl\-BamH\

fragment of the rat GLO cDNA into the

genome of Bombyx mori nuclear polyhedrosis virus using the transfer vector pBM030 as described by Maeda (5). With the resulting recombinant virus, high-level expression from the cDNA could be achieved under the control of the strong polyhedrin promoter. When cells of a silkworm cell line, BmN4, were infected with the recombinant virus and cultured in a normal medium [MGM-448 medium (6) supplemented with 10% fetal bovine serum (FBS)], the total cell extract obtained on the fifth day after the start of infection gave a specific activity of 16-20 nmol/min/mg protein (Fig. 1).

This level of activity was of the same order as found

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • New York - Printed in Germany

792

Fig. 1 Time course of G L O expression in B m N 4 cells infected with the recombinant baculovirus. Monolayers of B m N 4 cells (~10 6 cells) were infected with the recombinant virus at a multiplicity of infection of 10 plaque-forming units per cell and cultured at 2 7 ° C in M G M - 4 4 8 medium suppplemented with 10% F B S . Cells were harvested every day till the sixth day after the start of infection. The cell extracts, prepared by sonication in phosphatebuffered saline, were assayed for G L O as described previously (7) except for omission of dithiothreitol from the incubation mixture. in rat liver microsomes (14 nmol/min/mg protein) of male S D rats, the same strain used for the construction of the c D N A library from which the G L O c D N A was cloned. Western blot analysis showed that the G L O expressed in the silkworm cells had the same molecular weight ( - 5 1 , 0 0 0 ) as rat liver G L O . F A D was covalently bound to the recombinant G L O protein, as demonstrated by the appearance of yellowish green fluorescence at the position of G L O on a sodium dodecyl sulfate gel upon soaking in 10% acetic acid. These results indicate that G L O of a normal molecular weight is synthesized and is covalently bound to F A D in silkworm cells. To test the conditions for production of the apoprotein of G L O , we first cultured B m N 4 cells in M G M - 4 4 8 medium minus riboflavin but supplemented with 10% F B S for a week and then infected the cells with the recombinant virus (Fig. 2A). The G L O activity in the cells on the fifth day after the start of infection decreased considerably. When F A D (10 (J.M in final concentration) was added to the assay mixture, the activity almost doubled, indicating the production of the apoprotein of G L O . To make the culture medium more strictly deficient in riboflavin,

we used MGM-448 medium minus riboflavin without supplementation of F B S for

culturing the recombinant virus-infected cells. The G L O activity in the cells on the fifth day after the start of virus infection decreased to a greater degree; the activity was increased about

793

Fig. 2 Expression of GLO in BmN4 cells infected with the recombinant virus under riboflavin-deficient conditions. Cells were cultured for a week (A) or three weeks (B) in MGM-448 medium minus riboflavin but supplemented with 10% FBS, and then infected with the recombinant virus and cultured for five days in the same medium (+) or in MGM-448 medium minus riboflavin without supplementation of FBS (-). The cells cultured under normal conditions (N) were obtained on the fifth day after the start of virus infection. The GLO activity in the cell extract prepared from each culture was measured in the absence (open bars) or presence (solid bars) of FAD. fivefold by the addition of FAD to the assay mixture (Fig. 2A). On the other hand, the addition of FAD to the assay mixture had essentially no effect on GLO activity for the extract of recombinant virus-infected cells cultured under normal conditions, indicating that the GLO produced under such conditions was totally the holoenzyme. The cells that had been cultured in the riboflavin-deficient medium supplemented with 10% FBS for three weeks before virus infection gave essentially similar results, as shown in Fig. 2B.

It may be that flavins

contained in the supplemented FBS were sufficient in concentration to sustain the growth of BmN4 cells. Using the apoprotein of recombinant GLO, we examined whether FAD could bind covalently to the apoprotein. The extract from recombinant virus-infected cells cultured under the strictly riboflavin-deficient conditions was incubated with FAD (0.2 mM) at 37°C for 5 min and then subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The intensity of fluorescence at the position of GLO on the gel soaked in 10% acetic acid was not increased compared with that for the cell extract alone, indicating that covalent binding of FAD to the apoprotein did not occur. Thus, it is clear that noncovalent interaction between FAD and the apoprotein can elicit catalytic activity of the enzyme. This feature is in contrast with the observation that the formation of a covalent bond between FAD and the apoprotein paralleled

794 the appearance of enzymatic activity in the case of 6-hydroxy-D-nicotine oxidase (8).

References 1. Kenney, W. C., D. E. Edmondson, T. P. Singer, H. Nakagawa, A. Asano and R. Sato. 1976. Biochem. Biophys. Res. Commun. 71, 1194-1200 2. Kiuchi, K., M. Nishikimi and K. Yagi. 1982. Biochemistry 21, 5076-5082 3. Koshizaka, T„ M. Nishikimi, T. Ozawa and K. Yagi. 1988. J. Biol. Chem. 263, 16191621 4. Yagi, K., T. Koshizaka, M. Kito, T. Ozawa and M. Nishikimi. 1991. Biochem. Biophys. Res. Commun. 177, 659-663 5. Maeda, S. 1989. In: Invertebrate cell system applications (J. Mitsuhashi ed.). CRC Press, Boca Raton, FL. pp. 167-181 6. Mitsuhashi, J. 1984. Zool. Sci. 1, 415-419 7. Kito, M„ N. Ohishi and K. Yagi. 1991. Biochem. Int. 24, 131-135 8. Brandsch, R. and V. Bichler. 1991. J. Biol. Chem. 266, 19056-19062

Structures Responsible for FAD Binding and Substrate Recognition of Rat Liver Monoamine Oxidase

F. Ogata, Y. Tsugeno, I. Hirasiki, J. Mitoma, and A. Ito Department of Chemistry, Faculty of Science, Kyushu University 33, Hakozaki, Higashiku, Fukuoka 812, Japan

Introduction Monoamine oxidase (MAO), which catalyzes the oxidative deamination of biogenic and xenobiotic amines, is an integral membrane protein in the outer membrane of mitochondria and has a FAD bound to a cysteine residue covalently (1). There are two forms of enzyme (MAO A and MAO B) which have different substrate and inhibitor specificities and which are coded by different genes (2,3,4,5). We intended to elucidate the structures responsible for FAD binding by using point mutation and substrate specificity by the analysis of chimera enzymes constructed from MAO A and MAO B.

Results and Discussion To determine the structural requirement of the FAD binding site, we altered the cysteine residue(397Cys) which participates covalent attachment of FAD in MAO to Ala, Ser, or His residue by site directed mutagenesis and expressed the mutants in mammalian (COS cell) and yeast cells. All the mutant proteins were localized in mitochondria and were expressed in almost equal amount. The C397S and C397H proteins exhibited no enzyme activity, whereas the C397A proteins had a little activity in yeast, although the mutant showed little activity in the expression of COS cells (Table I and II). These activities could not restore by the addition of FAD or riboflavin either during or after the culture. These findings indicate that the covalent linkage of FAD and MAO polypeptide plays an important role in the formation of enzymatically active structure. The deletion mutant (MAOAC28) which lacks the targeting signal located in carboxyl terminal of MAO B (6) was also expressed and the enzyme activity was investigated. MAOAC28 had no activity, although it has the cysteine residue that is the FAD-binding amino acid. In yeast cell, MAOAC28 was localized in mitochondrial and cytosol fractions, however, the enzyme activity was not detected in both

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter & Co., Berlin • N e w York - Printed in Germany

796 fractions. The correct targeting or insertion of the protein into mitochondrial outer membrane seems to be closely concerned to the expression of the enzyme activity. These findings also suggest the close correlation between the covalent binding of FAD to MAO B protein and the insertion of the protein into the membrane, as a functionally competent form.

Table 1 Specific Activity of Native and Mutated MAO B Expressed in COS Cells. The activity was determined by the method of Kraml (7) using kynuramine as a substrate. Substrate(Kynuramine) (|iU/mg of protein) Control MAOB(wild-type)

(|xU/ng of MAO B)* ND**

1.6 24.9

0.983

1.1

0.048

MAOBC397A

1.5

0.030

MAOBC397H MAOBAC28

0.6 0.04

0.013 0.001

MAOBC397S

* The MAO content was determined by densitometric analysis after western blotting ** ND : not determined

Table 2 Specific Activities of Native and Mutated MAO B Expressed in Yeast. The activity assay was carried out by the methods of Kraml (7) and Wurtman and Axelrod (8) using [14C]-phenylethylamine (PEA) as a substrate. Substrate [14C]-Phenylethylamine

Kynuramine (|lU/mg of protein) Control MAOB(wild-type)

2.1 17.9

(M-U/mg of protein) (nU/ng of MAO B)* 6.2

103

120 7.0

MAOBC397S

1.8

MAOBC397A

4.0

MAOBC397H MAOBAC28(Mt)

1.9

4.0

9.0

0.12

6.3

2

MAOBAC28(Sup)

0.11

0.12

3

* and **; same as Table 1

8.0

ND**

14

24

797 To elucidate the region responsible for substrate recognition of MAO, the cDNAs of MAO A and MAO B were cut into three portions of similar size and six kinds of chimera enzymes were constructed from the cDNA fragments and were expressed in yeast. The kinetic constants (Vmax and Km) of the native and six chimera enzymes in the mitochondrial fractions were determined by using serotonin and phenylethylamine (PEA) as substrates (Table 3). Serotonin and PEA have been shown to be preferable substrates for MAO A and MAO B, respectively. In this table, the ratio of Vmax for serotonin to PEA was calculated and used as a criteria to determine which type of substrate specificity the chimeric enzyme has, because the extent of expression of the enzymes was not identical with each preparation. MAO A showed the ratio of Vmax for serotonin to PEA of 9.3, whereas that of MAOB was less than 0.16. The values were the representatives of the MAO A and MAO B type activity. MAOAAB had the ratio of 7.1 indicating that this chimeric enzyme has a substrate specificity of MAO A type. MAOABA, MAOABB, and MAOBBA had relatively small values of the ratio (serotonin/PEA). It means that all of them belong to the enzymes with MAO B type specificity. These results showed that the middle portion of MAO molecule plays an important role to recognize the substrate specificity.

Table 3 Kinetic Parameters and Substrate Specificity of Native and Chimeric MAO for Serotonin and Phenylethylamine (PEA). Vmax Native and Chimeric MAO Serotonin PEA

Km Serotonin/PEA

nmol/min/mg of protein

Serotonin PEA

Type

mM

A

5.0

0.54

9.3

0.30

0.22

A

B

Pi process was estimated from stopped-flow CD measurements. The estimates of these rate constants were then varied from the measured values to allow an optimal fit to the lag observed in the experimental data for recovery of activity following dilution from urea (Fig. 1) (9). The final values giving the best fits to the experimental data were very close to the values determined from the CD kinetic data. The first order rate constant for the a(ij—>afi isomerization was initially estimated from the kinetics of the shift that occurred in the rate of formation of a p upon dilution from 50 ng/ml to 5 |ig/ml (9). The rate constant was then varied to obtain the optimum fit to the experimental data, including the secondary dilution experiments. The second order rate constant for the homodimerization of the p subunit was measured using two approaches. First, the rate with which refolding P subunit became heterodimerization-incompetent was determined over a range of concentrations and the data fit to a second order mechanism. Second, the rate with which refolding P subunit formed the 5 M urea-insensitive structure was determined over a range of concentrations and the data fit to a second order mechanism. Both approaches gave similar values for the second order rate constant. The heterodimerization rate constant was determined by simulation. When we attempted to fit the data presented in Figure 1 using these 5 rate constants, we were able to simulate the early and intermediate portions of the curves quite satisfactorily, but at later times, the simulations invariably continued to give a slow increase in activity that was not demonstrated by the data. To account for the flattening of the time courses of activity recovery at later times of refolding, we have introduced a first order conversion of Pi—>P', a monomelic form of p subunit that is incompetent to heterodimerize. With the addition of this step, we have been able to fit the experimental data quite well (Fig. 1). It is this proposed first-order step that results in reduced yield of active enzyme at lower protein concentrations. The variance of the 50 |ig/ml data from the simulation is due, we believe, to aggregation of folding intermediates that occurs at the higher protein concentrations, which has not been incorporated into Fig. 2. Sugihara and Baldwin (8) have described p subunit termination mutants that appear to fold and assemble correctly at lower temperatures into proteins that have normal activity and stability, but at higher temperatures fail to assemble into the heterodimer. Based on the properties of these mutants, it was proposed that the carboxyl-terminal region of the P subunit must play a critical role in the folding and assembly reaction, but have little or no effect on the activity or stability of the successfully folded product (8). We have designed a series of mutants at position P313 based on the original termination mutants. The mutants, PD313A, pD313N, PD313G, and pD313P, all exhibit kinetic defects in the refolding reaction. However, they display the same conformational stability as the wild-type protein; in fact, the asparaginyl and alanyl mutants are slightly more stable than the wild-type protein. The prolyl mutant has the strongest kinetic defect of the four mutant enzymes. The lag phase in recovery of activity is the same as for the wild type, indicating that the process Pu~»Pi is the same. It appears that the heterodimerization rate constant is much lower for the mutant than for the wild-type protein; the time courses of activity recovery for the PD313P mutant can be satisfactorily fit to the model in Fig. 2 by changing only the heterodimerization rate constant. Likewise, the homodimerization rate constant of the prolyl mutant appears to be extremely low or non-existent. Examination of the prolyl mutant P subunit by analytical ultracentrifugation showed it to be monomelic. The PD313P mutant P subunit does not fold into a 5 M urea hyperstable structure, but rather folds into a structure without significant near-

828

ultraviolet circular dichroism, suggestive of a molten globule-like structure. It appears that the proposed Pi—>p' reaction for the wild-type protein also occurs for the PD313P mutant P subunit, whereas the homodimerization reaction does not occur. Conclusions It appears that the folding of luciferase subunits into the biologically active aP structure is a kinetically-determined process. The slow formation of PP leads to a hyperstable structure that does not catalyze the high quantum yield reaction. The observation of mutant proteins exhibiting kinetic defects in the folding reaction is entirely consistent with this hypothesis. It thus appears that the native structure of a protein must (a) be kinetically accessible and (b) have sufficient conformational stability to exist on a biological time scale. Alterations in the amino acid sequence may alter the kinetic pathway such that alternative structures become kinetically accessible. Clearly, in a folded protein there is substantial conformational flexibility, but it is unlikely that all conformations are in equilibrium under native conditions. Acknowledgements Research in this laboratory is supported in part by grants from the National Science Foundation (DMB 87-16262), the Office of Naval Research (N00014-91-J-4079 and N00014-92-J-1900), and the Robert A. Welch Foundation (A865). AFC was a Visiting Scientist with the Center for Macromolecular Design. His permanent address is Unité de Biochimie Cellulaire, Institut Pasteur, 28 rue du Dr. Roux, Paris 75724 (CEDEX 15), France. TOB and MMZ are deeply indebted to Prof. Michel Goldberg of the Pasteur Institute for his hospitality and involvement in the initial phases of this work. REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Ziegler, M. M. and T. O. Baldwin. 1981. Curr. Top. Bioenerg. 12, 65-113 Baldwin, T. O. and M. M. Ziegler. 1992. In: Chemistry and Biochemistry of Flavoenzymes, Vol. III. (F. Müller, ed.). CRC Press, Boca Raton, FL. pp. 467-530 Waddle, J. and T. O. Baldwin. 1991. Biochem. Biophys. Res. Commun. 178, 11881193 Sinclair, J. F., J. J. Waddle, E. F. Waddill and T. O. Baldwin. 1993. Biochemistry 32, 5036-5044 Friedland, J. and J. W. Hastings. 1967. Proc. Nat. Acad. Sei. U.S.A. 58, 2336-2342 Gunsalus-Miguel, A., E. A. Meighen, M. Z. Nicoli, K. H. Nealson and J. W. Hastings. 1972. J. Biol. Chem. 247, 398-404 Waddle, J. J., T. C. Johnston and T. O. Baldwin. 1987. Biochemistry 26, 4917-4921 Sugihara, J. and T. O. Baldwin. 1988. Biochemistry 27, 2872-2880 Ziegler, M. M., M. E. Goldberg, A.-F. Chaffotte and T. O. Baldwin. 1993. J. Biol. Chem. 268,10760-10765 Baldwin, T. O., M. M. Ziegler, A.-F. Chaffotte and M. E. Goldberg. 1993. J. Biol. Chem. 268, 10766-10772 Clark, A. C„ J. F. Sinclair and T.O. Baldwin. 1993. J. Biol. Chem. 268, 10773-10779 Cohn, D. H., A. J. Mileham, M. I. Simon, K. H. Nealson, S. K. Rausch, D. Bonam and T. O. Baldwin. 1985. J. Biol. Chem. 260, 6139-6146 Johnston, T. C„ R. B. Thompson and T. O. Baldwin. 1986. J. Biol. Chem. 261, 4805-4811

KINETIC MECHANISM OF THE BACTERIAL LUCIFERASE REACTION

W. A. Francisco, H. M. Abu-Soud, A. C. Clark, F. M. Raushel and T. O. Baldwin Center for Macromolecular Design and Departments of Chemistry and of Biochemistry and Biophysics, Texas A&M University, College Station, Texas 77843-2128

INTRODUCTION We have undertaken a detailed, multidimensional investigation of the kinetic mechanism of the bacterial luciferase-catalyzed reaction (1-3). Luciferase is a heterodimeric enzyme with a single active center on the a subunit. While the individual subunits exhibit low but authentic bioluminescence activity (4, 5), the active form of the enzyme is the heterodimer. The P subunit is required for the high quantum yield reaction, but its precise function is unknown (6). Light emission from the enzyme involves reaction of FMNH2, an aliphatic aldehyde and O2 on the surface of the enzyme to yield an excited state flavin and the carboxylic acid (6). One atom of the oxygen is found in the product carboxylate (7). It is assumed that the other atom from molecular oxygen is converted to water. FMN is the flavin product that is released following bioluminescence (8). It is known that the reaction proceeds through the intermediacy of the C4a-peroxydihydroflavin (9, 10) which can be distinguished from FMNH2 by the characteristic absorbance at 380 nm (10). The formation of FMN can be monitored by absorbance at 445 nm. Bioluminescence resulting from formation of the excited flavin species can likewise be monitored. The lifetimes of singlet excited states are typically in the nanosecond range so that the intensity of light emission at any time is proportional to the rate of formation of the excited state. It has been proposed that the emitter in the bioluminescence reaction is the C4a-hydroxyflavin (11); the FMN product is produced by dehydration of the C4a-hydroxyflavin. Several chemical mechanisms for the reaction of FMNH2, O2 and aldehyde have been proposed (6,12). We proposed a mechanism by which the proposed tetrahedral intermediate formed by reaction of the C4a-peroxyflavin with the aldehyde collapses to form the dioxirane and the C4a-hydroxyflavin (13; Fig. 1). The primary excited state suggested by this mechanism would be formed on the carboxylic acid product by collapse of the dioxirane. The C4a-hydroxyflavin would become excited by energy transfer from the primary excited state. In the presence of lumazine protein (14) or yellow fluorescence protein (15), the secondary emitter would likewise be excited by energy transfer. The experiments reported here comprise a detailed investigation of the kinetic mechanism of the luciferase catalyzed reaction (1-3). All measurements were made under conditions of 25°, 50 mM Bis-Tris HC1, pH 7.0. The enzyme concentration was maintained at 75 |iM for most

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • N e w York - Printed in Germany

830

C4a-peroxydihydroflavin

Tetrahedral Intermediate

Pseudobase

R

R

H Y O

Dioxirane Figure 1. Schematic representation of the reaction of the flavin C4a peroxide with the aldehyde substrate to yield the proposed dioxirane intermediate and the C4a hydroxyflavin from a tetrahedral intermediate (E'-FMNHOOR in Scheme I) (13). experiments, and the FMNH2» aldehyde and O2 concentrations were varied. The highest flavin concentration used was 15 (O.M. Experimental data were collected with a stopped flow spectrophotometer. Rate constants were determined either by fitting of the data to a specific equation or by simulation using KINSIM (16). The enzyme used in these experiments was purified from Escherichia coli carrying the luxAB genes from Vibrio harveyi on a pUC-derived plasmid. From this recombinant plasmid, we have been able to isolate about 1 gram of luciferase per liter of culture (17). The high level overproduction of luciferase was essential to the completion of this project, since the complete analysis required over 75 grams of enzyme. In some experiments, mutant forms of luciferase having mutations at position 106 of the a subunit were used. These mutant luciferases, aC106A, aC106S and aC106V, have been described previously (17-19). E-X

E'-FM NH,-X

kS3 2

° >- E' -FM NHOOH -X — — • FMN+H 2 0 2 +X k3x

k502 E-FMNH,

E'-FM NH,

E'-FM NHOOH

k25RCHO

k19RCHO

FMN

E'-FM NHOH

• FMN+HjOj

k,RCHO

E'-FMNH2-RCH0J^

E-RCHO

k, 7

E'-FMNHOOH-RCHO

— E'-FMNHOOR

k15RCHO

Scheme I representing the bacterial luciferase-catalyzed reaction.

831 RESULTS AND DISCUSSION

Three spectroscopic signals were utilized to determine the kinetic mechanism of the bacterial luciferase reaction. Absorbance measurements at 380 nm allowed determination of the formation of EFMNHOOH (10). Emission of visible light allowed measurement of processes occurring following addition of the aldehyde substrate, and absorbance measurements at 445 nm allowed detection of FMN formation from decay of E-FMNHOOH or from dehydration of the pseudobase, E-FMNHOH (8, 11). The time courses for the various transformations were determined as a function of the concentration of FMNH2, O2, aldehyde and enzyme. The minimal model that satisfies the complete data set is presented in Scheme I. The rate constants presented in Table I were progressively determined by fitting of the data to rate equations and by simulation of more complex reactions (1-3). The reaction of FMNH2 with O2 to yield FMN and H2O2 in the absence of enzyme was monitored at 380 nm and at 445 nm. The data were fit to the sum of two consecutive first-order reactions (A-»B-»C) where the two rate constants arc 4.7 s' 1 and 11.5 s"1; the order of the two rate constants, k2i and k23, is arbitrary. Formation and Decay of the Peroxydihydroflavin Intermediate

Table I: Rate Constants and Equilibrium Constants for the Model in Scheme I a 1.7 x 107 M' 1 s*1 ki 1200 s*1 k2 200 s"1 k3 14 s"1 k4 2.4 x 106 M"1 s- 1 k5 b 1.9 x 107 M"1 s"1 k7 b 120 s"1 kg b k9 1.6 s- 1 b 1.2 s"1 klO b 1.1 S"1 kn 0.60 s*1 kl3 b 3.0 x 103 M' 1 s"1 ki5 b 0.06 s*1 ki6 0.10 s- 1 kn 9.1 x 105 M"1 s"1 ki9 b b 5.8 s- 1 k2o 4.7 s- 1 k2i 11.5 s"1 k23 b 1.2 x 106 M -1 s*1 k25 37 s- 1 k26b b 5.1 x 104 M"1 s"1 k27 c 7.7 x 104 M-1 s"1 k33 k 37 c 0.004 s"1

The second-order rate constant (ks) for the M 3.9 x 103 M' 1 formation of E-FMNHOOH was determined by mixing K 2 d 6.1 x 103 M"1 E-FMNH2 with varying concentrations of O2. The K3100 fold) rate of decay of the E-FMNHOOH intermediate to yield FMN and H2O2 (kn) (2). The instability of the C4a-hydroperoxyflavin intermediate for the valine mutant (2) probably accounts for the results of Xi et al. (19). CONCLUSIONS

The results of these studies (1-3) comprise a set of rate constants defining the primary reactions catalyzed by the bacterial luciferase from Vibrio harveyi. These rate constants were determined under a single set of well-defined experimental conditions. It is clear from the complexity of the reaction that few valid conclusions can be drawn about the effects of inhibitors, mutations, buffer conditions, etc., on the reaction without performing a detailed kinetic analysis. The discovery of an isomerization of the E-FMNH2 complex to yield the O2reactive E'-FMNH2 was unexpected, but is consistent with reports of a two step mechanism for binding of FMNH2 to the enzyme of Photobacterium phosphoreum (27) and a conformational change that occurs in the Vibrio harveyi enzyme during the catalytic cycle (28). The mechanism of aldehyde substrate inhibition appears to reside simply in the ordered binding of substrates (1-3). If enzyme and aldehyde are mixed prior to addition of FMNH2, FMNH2 binding cannot occur until after aldehyde release. The inhibition is due to loss of the free FMNH2 to reaction with O2 prior to binding to the enzyme. Formation of the ternary complex E-FMNH2-RCHO reduces the rate of formation of E-FMNHOOH, but does not greatly reduce the bioluminescence quantum yield. Oxygen can react directly with the complex, albeit at a reduced rate, and if the aldehyde temporarily dissociates, O2 can react with the E'-FMNH2 very rapidly (1). The rate constants shown in Table I allow simulation with high precision of the various reaction time courses that occur on the V. harveyi enzyme. These results should serve as a foundation for investigations into the details of the chemical mechanism of bacterial luciferase. ACKNOWLEDGEMENTS

The research in the laboratories of FMR and TOB is supported in part by grants from the National Institutes of Health (GM 33894), the National Science Foundation (DMB 87-16262), the Office of Naval Research (N00014-91-J-4079 and N00014-92-J-1900), and the Robert A. Welch Foundation (A865). The enzyme used in these studies was purified by Vicki Green. We are indebted to Dr. Miriam M. Ziegler for advice and criticism of the manuscript, and to Nancy Harvey for preparation of the manuscript.

836 REFERENCES 1.

Abu-Soud, H„ L. S. Mullins, T. O. Baldwin and F. M. Raushel. 1992. Biochemistry. 31, 3807-3813

2.

Abu-Soud, H., A. C. Clark, T. O. Baldwin and F. M. Raushel. 1993. J. Biol. Chem. 268, 7699-7706

3.

Francisco, W. A., H. M. Abu-Soud, T. O. Baldwin and F. M. Raushel. 1993. J. Biol. Chem., in press

4.

Waddle, J. and T. O. Baldwin. 1991. Biochem. Biophys. Res. Commun. 178, 11881193

5.

Sinclair, J. F., J. J. Waddle, E. F. Waddill and T. O. Baldwin. 1993. Biochemistry. 32, 5036-5044

6.

Baldwin, T. O. and M. M. Ziegler. 1992. In: Chemistry and Biochemistry of Flavoenzymes, Vol. III. (F. Müller, ed.). CRC Critical Reviews in Biochemistry, CRC Press, Boca Raton, FL. pp. 467-530

7.

Suzuki, K., T. Kaidoh, M. Katagiri and T. Tsuchiya. 1983. Biochim. Biophys. Acta. 722, 297-301

8.

Hastings, J. W. and Q. H. Gibson. 1963. J. Biol. Chem. 238, 2537-2554

9.

Vervoort, J., F. Müller, J. Lee, W. A. M. van Den Berg and C. T. W. Moonen. 1986. Biochemistry. 25, 8062-8067

10. Hastings, J. W„ C. Balny, C. LePeuch and P. Douzou. 1973. Proc. Natl. Acad. Sei. U.S.A. 70, 3468-3472 11. Kurfuerst, M„ P. Macheroux, S. Ghisla and J. W. Hastings. 1987. Biochim. Biophys. Acta. 924, 104-110 12. Ziegler, M. M. and T. O. Baldwin. 1981. Curr. Top. Bioenerg. 12, 65-113 13. Raushel, F. and T. O. Baldwin. 1989. Biochem. Biophys. Res. Commun. 164, 11371142 14. Lee, J., I. B. C. Matheson, F. Müller, D. J. O'Kane, J. Vervoort. and A. J. W. G. Visser. 1990. In: Chemistry and Biochemistry of Flavoenzymes, Vol. II (F. Müller, ed.). CRC Press, Boca Raton, FL. 109-151 15. Daubner, S. Colette and T. O. Baldwin. 1989. Biochem. Biophys. Res. Commun. 161, 1191-1198 16. Barshop, B. A., R. F. Wrenn. and C. Frieden. 1983. Anal. Biochem. 130, 134-145

837 17. Baldwin, T. O., L. H. Chen, L. J. Chlumsky, J. H. Devine and M. M. Ziegler. 1989. J. Biolumin. Chemilumin. 4,40-48 18. Baldwin, T. 0 . , L. H. Chen, L. J. Chlumsky, J. H. Devine, T. C. Johnston, J.-W. Lin, J. Sugihara, J. J. Waddle and M. M. Ziegler. 1987. In: Flavins and Flavoproteins, (D. B. McCormick and D. E. Edmondson, eds.). Walter de Gruyter, Berlin, pp. 621-631 19. Xi, L„ K.-W. Cho, M. E. Herndon and S.-C. Tu. 1990. J. Biol. Chem. 265, 4200-4203 20. Holzman, T. F. and T. O. Baldwin. 1983. Biochemistry. 22, 2838-2846 21. Spudich, J. and J. W. Hastings. 1963. J. Biol. Chem. 238, 3106-3108 22. Tu, S. C. 1979. Biochemistry. 18, 5940-5945 23. Makemson, J. C„ J. W. Hastings and J. M. E. Quirke. 1992. Arch. Biochem. Biophys. 294, 361-366 24. Nicoli, M. Z„ E. A. Meighen and J. W. Hastings. 1974. J. Biol. Chem. 249, 2385-2392 25. Cohn, D. H., A. J. Mileham, M. I. Simon, K. H. Nealson, S. K. Rausch, D. Bonam and T. O. Baldwin. 1985. J. Biol. Chem. 260, 6139-6146 26. Ziegler, M. M. and T. O. Baldwin. 1981. In: Bioluminescence and Chemiluminescence: Basic Chemistry and Analytical Applications (M. A. DeLuca and W. D. McElroy, eds.). Academic Press, New York. pp. 155-160 27. Watanabe, T., K. Yoshida, M. Takahashi, G. Tomita and T. Nakamura. 1976. In: Flavins and Flavoprotein (T. P. Singer, ed.). Elsevier, Amsterdam, pp. 62-67 28. AbouKhair, N. K„ M. M. Ziegler and T. O. Baldwin. 1985. Biochemistry. 24, 39423947

BACTERIAL LUCIFERASE: BIOLUMINESCENCE EMISSION USING LUMAZINES AS SUBSTRATES

Peter Macheroux and Sandro Ghisla University of Konstanz, Faculty of Biology, P. O. Box 5560, D-78434 Konstanz, Germany J. Woodland Hastings Harvard University, The Biological Laboratories, 16 Divinity Avenue, Cambridge, MA 02138

Introduction In the bacterial luciferase reaction reduced flavin mononucleotide (FMNH2) serves as a reactant to form the flavin-4a-hydroperoxide [1,2], which, in turn, reacts with a long-chain aldehyde to yield the excited state [3]. Luciferase can use various flavin analogs in the reaction; such analogs may have effects on the spectrum, kinetics and yield of light emission [4,5,6]. However, no molecules other than flavin derivatives have been found to catalyze the emission of light. We report here that the reduced forms of tetrahydro-FMN (4H-FMN), which is structurally analogous to FMN but chemically a lumazine (see structures in scheme 1), and lumazine 8-ribityl phosphate, are also capable of acting as substrates in the light emitting reaction.

Results and Discussion There is a major difference between reduced flavin, which is the 1,5-dihydro-tautomer (Scheme 1, III), and the products obtained upon 2 e- reduction of 8-substituted lumazines (IV, V). With the latter the 7,8-tautomer (V) is the more stable form thermodynamically [7], but luciferase from Vibrio harveyi evidently binds both, since the protein fraction after G-25 gel filtration has spectral characteristics indicative of a mixture of the two (Figure 1, curve A). Upon standing at 4 °C (and final warming at 25 °C), the spectrum changes to that shown in curve B, with isosbestic points at 358 and 505 nra. Further spectral changes occur upon treatment of the sample with 2% sodium dodecyl sulfate (curve C). The existence of a luciferase-bound hydroperoxy intermediate is indicated by the fact that bioluminescence occurs upon the addition of long chain aldehyde to the protein fraction

Flavins and Flavoproteins 1993 © 1994 Walter de Gruyter& Co., Berlin • New York - Printed in Germany

840 from gel filtration. Its spectral identity is not known, but only the 5,8 dihydro form (IV) would be expected to form such a hydroperoxide, and it would be expected to be metastable [7]. Light emission can also be obtained by initiating the reaction with 4H-FMNH 2 in the presence of aldehyde.

Its intensity is about 10% that observed with FMNH^

an

d the

emission maximum is shifted slightly to the blue (Figure 2). A dependence of the emission intensity on the 4H- FMNH 2 concentration yields an apparent Km of 8 |i.M which compares to 0.3 |J.M for FMNH2. A similar response was obtained with reduced lumazine 8-ribityl phosphate. These results indicate that 4H-FMNH 2

can

indeed replace the normal substrate of the V.

harveyi luciferase reaction, and that the isoalloxazine ring system is not obligatory for the chemistry of the steps leading to light emission. With regard to its oxygen reactivity, the dihydrolumazine can be assumed to be similar to that of reduced flavin, and indeed the corresponding tetrahydropteridines have been shown to form hydroperoxides which are necessary intermediates during the hydroxylation of aromatic amino acids catalyzed by enzymes such a phenylalanine hydroxylase. We thus assume that the 5,8-dihydrolumazine chromophore bound to luciferase forms a hydroperoxide, which can react with long chain aldehyde in a way similar to that occurring in the normal reaction (Scheme 2). The primary function of the flavin/lumazine might thus be to provide a hydroperoxide of appropriate reactivity.

Scheme 1. Structures of FMN (I);

tetrahydro-FMN (II);

1,5-dihydro-FMN (III); 5,8-

dihydro-8-R-lumazine, obtained by the 2e- reduction of tetrahydro-FMN (IV), and the corresponding, tautomeric 7,8-dihydro form (V).

841 0,6

LU

< m cc S

m