Chemistry and Biochemistry of Flavoenzymes: Volume III [1 ed.] 9781315891460, 9781351070560, 9781351087469, 9781351095914, 9781351079013

Table of contents :

1. Pyridoxine-5-P Oxidase 2. Xanthine Oxidase, Xanthine Dehydrogenase and Aldehyde Oxidase 3. D- and L-Amino Acid Oxidases 4. Methanol Oxidase 5. Lipoamide Dehydrogenase, Glutathione Reductase, Thioredoxin Reductase and Mercuric Ion Reductase - Family of Flavoenzyme Transhydrogenases 6. Refined Three-Dimensional Structure of Glutathione Reductase 7. Structure and Function of Succinate Dehydrogenase and Fumarate Reductase 8. Three-Dimensional Structure of Medium-Chain Acyl-CoA Dehydrogenase 9. Glutamate Synthase 10. Assimilatory Nitrate Reductase 11. Biological Reduction and Formation of Sulfate; The Role of APS Reductase, an Iron-Sulfur- Containing Protein 12. The Stereochemistry of the Prosthetic Groups of Flavoproteins 13. Structure and Mechanism of Spinach Glycolate Oxidase 14. General Properties of Flavodoxins 15. Structure and Redox Properties of Clostridial Flavodoxin 16. Biochemistry and Molecular Biology of Bacterial Bioluminescence 17. Acetolactate Synthase 18. Phthalate Dioxygenase Reductase and Related Flavin-Iron-Sulfur-Containing Electron Transferases 19. Nuclear Magnetic Resonance Studies on Flavoproteins 20. Acyl-Coenzyme A Dehydrogenases

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Chemistry and Biochemistry of Flavoenzymes Volume III Editor

Franz Muller, Ph.D, Head Central Development Toxicology Sandoz Agro, Ltd. Basle, Switzerland and Adjunct Professor Department of Biochemistry University of Georgia Athens, Georgia

Boca Raton London New York

CRC Press is anCRC imprint Press of the Taylor Raton & Francis Group, an informaLondon business Boca Ann Arbor

First published 1992 by CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 Reissued 2018 by CRC Press © 1992 by CRC Press, Inc. CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright. com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a notfor-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data   (Revised for volume 3) Chemistry and biochemistry of flavoenzymes. Includes bibliographical references and index. 1.  Flavoproteins.  2. Flavins.  I. Müller, Franz, 1934– QP552.F54C44  1990   574.19’258    90-40024 ISBN 0-8493-4393-3 (v. 1) ISBN 0-8493-4394-1 (v. 2) ISBN 0-8493-4395-x (v. 3) A Library of Congress record exists under LC control number: 90040024 Publisher’s Note The publisher has gone to great lengths to ensure the quality of this reprint but points out that some imperfections in the original copies may be apparent. Disclaimer The publisher has made every effort to trace copyright holders and welcomes correspondence from those they have been unable to contact. ISBN 13: 978-1-315-89146-0 (hbk) ISBN 13: 978-1-351-07056-0 (ebk) Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

PREFACE Flavin was discovered about a century ago. Since its discovery, the field of flavin and flavoproteins has developed into a mature science. Although a few flavoproteins had already been discovered during the first three decades of the 20th century, a profound understanding of the enzymic reaction mechanisms of flavoproteins became possible only after full development of the chemical and physical properties of free flavin. This development was initiated about 60 years ago and was mainly concerned with the chemical synthesis of the flavin molecule. Research in the field of flavins and flavoproteins was discontinued during the Second World War, but revived in the 1950s. The postwar period brought increased progress in the chemistry of free flavin, which knowledge was instrumental in understanding the biochemical properties of flavoproteins. Flavoproteins are involved in a variety of biological reactions including electron transfer, oxidation, dehydrogenation, and monooxygenation reactions. Many flavoproteins contain flavin as the sole prosthetic group, but the reaction mechanisms can still be complex. Other flavoproteins are composed of flavin and heme and non-heme iron, molybdenum and other metal ions, or pteridin. Although they represent a minority, flavoproteins resulting from covalent bondings of the prosthetic group to amino acid residues of proteins have enhanced the variety of flavoproteins. Present knowledge in the field discussed in this work would not be possible without advancements in biophysical equipment. New techniques have contributed a great deal to the elucidation of primary, secondary, tertiary, and quaternary structures of flavoproteins. In recent years the three-dimensional structures of a few flavoproteins have become available, and it can be expected that this research will continue to advance. These data will provide the basis for new research on flavoproteins — namely, cloning of genes, encoding for flavoproteins, followed by site-directed mutagenesis of flavoproteins. This research is presently in its infancy, but is expected to develop and provide even deeper insight into the enzymic mechanisms of flavoproteins and the biochemical principles governing the catalytic action of particular flavoenzymes. Given the wealth of data available on flavins and flavoproteins, it seems appropriate to report on the progress of a century of research in the field. In the present work an attempt has been made to summarize this knowledge as comprehensively as possible. Despite limitations of space and unavailability of some material, this and succeeding volumes of the Chemistry and Biochemistry of Flavoenzymes should be a valuable reference for researchers in the field and those entering it, as well as a source for lecturers in biochemistry, biophysics, chemistry, pharmacology, toxicology, and medicine. In conclusion I wish to thank all contributors for their cooperation and their efforts to meet our deadlines, and I would also like to express personal appreciation to my wife, Rita, and our daughters, Sandra and Kirsten, for their patience over many weekends in the development of this series. Franz Miiller

THE EDITOR Franz Muller, Ph.D., is Head, Central Development of Toxicology, Sandoz Agro Ltd., Basle, Switzerland and Adjunct Professor of Biochemistry, University of Georgia, Athens, Georgia. Dr. Muller obtained his training at the University of Basle, and received his M.S. degree in 1961 and Ph.D. degree in 1964 from the Departments of Organic and Inorganic Chemistry. After doing postdoctoral work at the Medical Nobel Institute, Department of Biochemistry, Stockholm, Sweden, he was first a Research Associate in the Department of Biological Chemistry at the University of Michigan in Ann Arbor, and later Instructor, and a recipient of a Career Research Development Award from the National Institutes of Health. In 1971 he was appointed an Associate Professor of Biochemistry at the Agricultural University, Wageningen, The Netherlands, and Research Professor in 1977, where he served as Chairman of the Department of Biochemistry and chaired the university's Science Committee and the Committee for Biotechnology. In 1987 he assumed his present position. Dr. Muller is a member of the Swiss Chemical Society, the Swiss Association of Chemists, the Federation of European Biochemical Societies and the New York Academy of Sciences. He received the Venia Legendi (Habilitation) from the University of Constance, Federal Republic of Germany, in 1970. Dr. Muller has been the recipient of research grants from The Netherlands Science Foundation. He was also a member of the Committee for Natural Sciences of the grant agency and chaired the Section for Protein Chemistry. He has published more than 200 papers. His current research interests are in biochemical transformation of pesticides.

CONTRIBUTORS Brian A. C. Ackrell, Ph.D. Molecular Biology Division Veterans Administration Medical Center San Francisco, California Thomas O. Baldwin, Ph.D. Department of Biochemistry and Biophysics Texas A&M University College Station, Texas David P. Ballou, Ph.D. Department of Biological Chemistry University of Michigan Ann Arbor, Michigan Michael J. Barber, D. Phil. Department of Biochemistry and Molecular Biology College of Medicine University of South Florida Tampa, Florida P. I. H. Bastiaens, M.Sc. Department of Biochemistry Agricultural University Wageningen, The Netherlands Christopher J. Bade, Ph.D. Department of Biochemistry and Molecular Biology University of Louisiana Medical School New Orleans, Louisiana

Bruno Curti, Ph.D. Department of General Physiology and Biochemistry University of Milano Milano, Italy Paul C. Engel, D. Phil. Department of Molecular Biology and Biotechnology Krebs Institute University of Sheffield Sheffield, England Robert P. Gunsalus, Ph.D. Department of Microbiology and Molecular Genetics and Molecular Biology Institute University of California Los Angeles, California Russ Hille, Ph.D. Department of Medical Biochemistry Ohio State University Columbus, Ohio T. R. Hopkins, Ph.D. Biospec Products, Inc. Bartlesville, Oklahoma Michael K. Johnson, Ph.D. Department of Chemistry University of Georgia Athens, Georgia

Gary Cecchini, Ph.D. Molecular Biology Division Veterans Administration Medical Center San Francisco, California

Andrew Karplus, Ph.D. Biochemistry, Molecular and Cell Biology Section Cornell University Ithaca, New York

Jorge E. Churchich, Ph.D. Department of Biochemistry University of Tennessee Knoxville, Tennessee

Jung-Ja P. Kim, Ph.D. Department of Biochemistry Medical College of Wisconsin Milwaukee, Wisconsin

Carl C. Correll, M.Sc. Department of Biological Chemistry and Biophysics Research Division University of Michigan Ann Arbor, Michigan

Francis Kwok, Ph.D. Department of Applied Biology and Chemical Technology Hong Kong Polytechnic Hong Kong

Jorge Lampreia, Ph. D, Center of Chemical and Biological Technology (CTQB) Oeiras, Portugal

Emil F, Pai, Dr. rer. nat. Department of Biophysics Max-Planck-Institute for Medical Research Heidelberg, Germany

John Lee, Ph.D. Department of Biochemistry University of Georgia Athens, Georgia

Harry D. Peck, Jr., Ph.D. Department of Biochemistry University of Georgia Athens, Georgia

Jean LeGall, Ph.D. Department of Biochemistry University of Georgia Athens, Georgia Ylva Lindquist, Ph.D. Department of Molecular Biology Swedish University of Agricultural Science Uppsala, Sweden Martha L. Ludwig, Ph.D. Biophysics Research Division and Department of Biological Chemistry University of Michigan Ann Arbor, Michigan Catherine L. Luschinsky, M. S. Department of Biological Chemistry University of Michigan Ann Arbor, Michigan Stephen G. Mayhew, Ph.D. Department of Biochemistry University College Belfield Dublin, Ireland Isabel Moura, Ph. D. Center of Chemical and Biological Technology (CTQB) Oeiras, Portugal Jose J. G. Moura, Ph.D. Center of Chemical and Biological Technology (CTQB) Oeiras, Portugal Franz Muller, Ph.D. Department of Toxicology Sandoz, Ltd. Basle. SwitvfM-i^rn-i

Severino Ronchi, Ph.D. Institute of Veterinary Physiology and Biochemistry University of Milano Milano, Italy John V. Schloss, Ph.D. Department of Medicinal Chemistry University of Kansas Lawrence, Kansas G. E. Schulz, Dr. Rer. Nat. Institute of Organic and Biochemistry Albert-Ludwigs University Freiburg, Germany Mirella Pilone Simonetta, D. Biol. S., Department of General Physiology and Biochemistry University of Milano Milano, Italy Larry P. Solomonson, Ph.D. Department of Biochemistry and Molecular Biology College of Medicine University of South Florida Tampa, Florida Gordon Tollin, Ph.D. Department of Biochemistry University of Arizona Tucson, Arizona Maria A. Vanoni, Ph.D. Department of General Physiology and Biochemistry University of Milano \s:\~~~. T+~U.

Charles H. Williams, Jr., Ph.D. Research Service Department of Veterans Affairs Medical Center Ann Arbor, Michigan

Jin Wu, B.S. Department of Chemistry Syracuse University Syracuse, New York

Antonio V. Xavier, Ph.D. Center of Chemical and Biological Technology (CTQB) Oeiras, Portugal G. Zanetti, Ph.D. Department of General Physiology and Biochemistry University of Milano Milano, Italy

Miriam M. Ziegler, Ph.D. Department of Biochemistry and Biophysics Texas A&M University College Station, Texas

TABLE OF CONTENTS Chapter 1 Pyridoxine-5-P Oxidase F, Kwok and J. E. Churchich

1

Chapter 2 Xanthine Oxidase, Xanthine Dehydrogenase, and Aldehyde Oxidase R. Hille

21

Chapter 3 D- and L-Amino Acid Oxidases B. Curti, S. Ronchi, and M. P. Simonetta

69

Chapter 4 Methanol Oxidase F. Miiller, T. R. Hopkins, J. Lee, and P. I. H. Bastiaens

95

Chapter 5 Lipoamide Dehydrogenase, Glutathione Reductase, Thioredoxin Reductase, and Mercuric Ion Reductase — A Family of Flavoenzyme Transhydrogenases C. H. Williams, Jr.

121

Chapter 6 Refined Three-Dimensional Structure of Glutathione Reductase P. A. Karplus and G. E. Schulz

213

Chapter 7 Structure and Function of Succinate Dehydrogenase and Fumarate Reductase B. A. C. Ackrell, M. K. Johnson, R. P. Gunsalus, and G. Cecchini

229

Chapter 8 Three-Dimensional Structure of Medium-Chain Acyl-CoA Dehydrogenase J.-J. P. Kim and J. Wu

299

Chapter 9 Glutamate Synthase M. A. Vanoni, B. Curti, and G. Zanetti

309

Chapter 10 Assimilatory Nitrate Reductase M. J. Barber and L. P. Solomonson

319

Chapter 11 Biological Reduction and Formation of Sulfate; the Role of APS Reductase, and FAD(Iron-Sulfur)-Containing Protein J. Lampreia, L Moura, A, V. Xavier, J. Le Gall, H. D. Peck, Jr., and J. J. G. Moura

333

Chapter 12 The Stereochemistry of the Prosthetic Groups of Flavoproteins E. F. Pai

357

Chapter 13 Structure and Mechanism of Spinach Glycolate Oxidase Y. Lindqvist

367

Chapter 14 General Properties of Flavodoxins S. G. Mayhew and G. Tollin

389

Chapter 15 Structure and Redox Properties of Clostridial Flavodoxin M. L. Ludwig and C. L. Luschinsky

427

Chapter 16 Biochemistry and Molecular Biology of Bacterial Bioluminescence T. O. Baldwin and M. M. Ziegler

467

Chapter 17 Acetolactate Synthase J. V. Schloss

531

Chapter 18 Phthalate Dioxygenase Reductase and Related Flavin-Iron-Sulfur-Containing Electron Transferases C. J. Bade, D. P. Ballou, and C. C. Correll

543

Chapter 19 Nuclear Magnetic Resonance Studies on Flavoproteins F. Miiller

557

Chapter 20 Acyl-Coenzyme A Dehydrogenases P. C. Engel

597

Index

657

Volume III

1

Chapter 1

PYRIDOXINE-5-P OXIDASE Francis Kwok and Jorge E. Churchich TABLE OF CONTENTS I.

Introduction

2

II.

Assay A. Colorimetric Method B. Spectrophotometric Method C. Other Methods

3 3 3 3

III.

Purification of Pyridoxine 5'-P Oxidase A. Sheep Brain Pyridoxine 5'-P Oxidase B. Aggregation of the Oxidase

3 4 5

IV.

Resolution of Pyridoxine 5'-P Oxidase A. The KBr Method B. The Ammonium Sulfate Method

6 6 6

V.

Substrate Specificities

6

VI.

Properties of the FMN-Binding Site A. Studies Using FMN Analogues B. Spectroscopic Studies C. Emission Anisotrophy

7 7 10 12

VII.

Properties of the Substrate-Binding Site A. Chemical Modification Studies B. Kinetic Studies C. Studies on Stereospecificity

12 12 14 16

VIII.

Regulation A. Interaction with Pyridoxal Kinase B. Pyridoxal 5'-P Oxidase in Tumors

17 17 17

References

18

2

Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION Pyridoxal 5'-phosphate, the metabolically active form of vitamin B6, is the coenzyme required by numerous enzymes involved in transamination, and racemization reactions. Two reaction steps are included in the conversion of pyridoxine and pyridoxamine to pyridoxal 5'-phosphate: (1) phosphorylation catalyzed by pyridoxal kinase and (2) oxidation of the phosphorylated vitamins catalyzed by the FMN-dependent pyridoxine (pyridoxamine) 5'phosphate oxidase. The reactions are shown as in the following:

Pyridoxine (pyridoxamine) 5'-P oxidase (EC1.4.3.5) was first found in rabbit liver by Pogell1 and later studied by many other laboratories including the laboratory of Wada and Snell.2 The localization of this enzyme in mammals covers a wide range of tissues including liver, kidney, and brain with high activities; and heart, skeletal muscle, pancreas, and bone marrow with relatively lower activities.3 These differences in oxidase activities among different tissues led to the establishment of a complicated network for the pyridoxal 5'-P distribution because tissues with high oxidase activities produce pyridoxal 5'-P not only for internal consumption but also for external supply to other tissues with low oxidase activities. An example of the distribution network suggested by Lumeng et al.4 is that the synthesis of pyridoxal 5'-P in muscle is not adequate for its own tissue consumption and as a result, additional supply of pyridoxal 5'-P has to come from either the liver cells or erythrocytes via the circulation. Each tissue maintains an independent pool of vitamin B6 which includes pyridoxine, pyridoxamine, and pyridoxal. The content of different chemical forms of vitamin B6 in this pool is regulated by a combination of enzymes, such as pyridoxine 5'-P oxidase, pyridoxal kinase, different species of phosphatases, and various pyridoxal 5'-binding proteins. Then, the activity of any one of the above enzymes or proteins is also regulated by metabolites from other metabolic pathways. This is demonstrated by findings such as decreases in pyridoxine 5'-P oxidase activity as a result of riboflavin deficiency5 and the activation of rat liver oxidase activity by 3-hydroxykynurenine and 3-hydroxyanthranilate both of which are metabolites of tryptophan metabolism.6 Apart from mammalian tissues, pyridoxine 5'-P oxidase activities have also been found in other eukaryotic systems, including yeast7 and wheat seedlings.8 The yeast enzyme has been found to be activated by various aliphatic amines7 and no isoenzymes from yeast have been found. Three isoenzyme forms of pyridoxine 5'-P oxidase have been identified from wheat seedlings and two of them have been partially purified.8 Although much evidence has shown that pyridoxine 5'-P oxidase is an enzyme widely distributed in tissues because of its importance in biochemical reactions in both mammalian and plant tissues, it appears that tumor tissue utilizes a different pathway in the synthesis of pyridoxal 5'-P apart from the pathway utilized by normal tissues. Nutter et al.9 reported that no pyridoxine 5'-P oxidase activity was detected in Morris hepatoma cells suggesting

Volume III

3

that tumor tissues do not require the oxidase activity for pyridoxal 5'-P synthesis. The possibilities of acquiring the vitamin from other normal tissues by tumor tissues or synthesizing it via a nonconventional pathway have been suggested.

II. ASSAY The optimum pH of pyridoxine (pyridoxamine) 5'-P oxidase was recorded at 9 for the rabbit liver enzyme and 8.4 for the brain enzyme. Several assay systems have been designed for the oxidase. A. COLORIMETRIC METHOD The colorimetric method as developed by Wada and Snell2 utilizes the development of a color adduct from the reaction between phenylhydrazine in sulfuric acid with pyridoxal 5'-P. The formation of a yellowish complex can be monitored spectrophotometrically at 412 nm. An extinction coefficient of 23,000 cm~ * M~l is used for calculating the concentration of the complex which is directly proportional to enzymatic activity. This method is more appropriately adopted for the assay of oxidase activity in crude homogenate because of interference created by turbidity of the assay mixture. Prior to the addition of phenylhydrazine and further colorimetric measurement, the proteins in the assay mixture should have been precipitated using \M trichloroacetic acid and then removed by centrifugation. B. SPECTROPHOTOMETRIC METHOD This spectrophotometric method is found to be more convenient for routine procedures. It measures the oxidase activity by monitoring the formation of pyridoxal 5'-P which is proportional to the increase in absorbance at 388 nm. Pyridoxal 5'-P is known to have an extinction coefficient of 4900 cm~ l M~l at pH 7. Initial rate measurement is carried out by monitoring the change in absorbance at 388 nm for at least 3 min in a spectrophotometer. This method is only recommended for the assay of pyridoxine 5'-P oxidase activity in nonturbid solution. C. OTHER METHODS Other less commonly used methods include continuous monitoring using polarographic techniques for measurement for O2 consumption or using substrates containing either a fluorescent or radioactive group that is released by the oxidase reaction.10 For the assay of rabbit liver pyridoxamine 5'-P oxidase, Kazarinoff and McCormick11 use 0.2 M Tris-HCl buffer at pH 8 as the assay medium. In addition, pyridoxamine 5'-P was the preferred substrate used although no difference in the maximum velocity between pyridoxamine 5'-P or pyridoxine 5'-P was observed. However, a difference of tenfold in oxidase activity was observed between using the two phosphorylated vitamins as substrates in the assay of the brain enzyme. Therefore, Kwok and Churchich12 recommended the use of pyridoxine 5'-P as the sensitive substrate in the assay of pyridoxine 5'-P oxidase activity in brain extracts.

III. PURIFICATION OF PYRIDOXINE 5'-P OXIDASE Pyridoxine 5'-P oxidase was purified from rabbit liver by Kazarinoff and McCormick11 and from porcine brain by Kwok and Churchich.12 The inclusion of a step using phosphopyridoxyl-Sepharose by Churchich13 led to a higher efficiency achieved in the purification of pig brain oxidase and later, this step of affinity chromatographic procedure was adapted successfully in the purification of other mammalian oxidases.14'15

4

Chemistry and Biochemistry of Flavoenzymes TABLE 1 Purification of Pyridoxine-5-Phosphate Oxidase from Sheep Brain Volume (ml)

Protein (mg)

28379 2277 2152 2150

752040 81070 10440 3250

500 22

25

Homogenate (NH4)2SO4 (40—60%) DEAE-cellulose fraction pH 5.0 treatment Phosphopyridoxyl-Sepharose Sephadex G-100

Specific activity (U/mg)

Total activity (U)

0.06 0.155 1.29

4.2 312 341

15.3

50,019 12,614 13,560 13,500 7,800 5,226

Note: The purification of pyridoxine 5'-P oxidase was made from 20 kg of sheep brain.

TABLE 2 Purification of Pyridoxal Kinase from a Sheep Brain Treatment Homogenate 40— 60% (NH4)2SO4 fraction DEAE-cellulose fraction Pyridoxal-agarose fraction Sephadex G-100

Volume (ml)

Protein (mg)

10500 1680 1500

278250 59808 9450

60 40

64 25.7

Protein (rag/ml)

26.5 35.6

6.3 1.07 0.643

Specific activity (units/ing)

Total activity (units)

0.125 0.668 5.81 5.32 1104

34781 39951 54904 34048 28373

Note: The purification of pyridoxal kinase was made from 20 kg of sheep brain tissue wet weight.

A. SHEEP BRAIN PYRIDOXINE 5'-P OXIDASE Purification of pyridoxine 5'-P oxidase from sheep brain is adapted from the procedure for the purification of pyridoxal kinase.16 Both procedures share the common steps of homogenization, ammonium sulfate fractionation, and DEAE-cellulose chromatography. During the elution of DEAE-cellulose chromatography, the profile exhibits partially overlapping peaks of kinase and oxidase activities. Fractions containing both enzymatic activities are allowed to pass through a pyridoxal-Sepharose column which retains only the kinase activity, and the straight-through material is combined with other fractions containing the oxidase activity. Acid precipitation using 0.2 N acetic acid to adjust the pH from 7.4 to 3.5 and phosphopyridoxyl-Sepharose chromatography eluted with 10 ~ 2 M pyridoxal 5'-P at pH 5 are introduced as further steps of the purification procedure after DEAE-cellulose chromatography. The last step of gel filtration using Sephadex G-100 provides the separation of the oxidase from both contaminating proteins and also the eluting ligand of the affinity chromatography pyridoxal 5'-P. After the last step of gel filtration, the oxidase preparation is observed as a single protein band in SDS-polyacrylamide gel electrophoresis. Comparison of summarized procedures for the purification of pyridoxine 5'-P oxidase and pyridoxal kinase are shown in Table 1 and 2, respectively. In 10% SDS-polyacrylamide gel electrophoresis, pyridoxine 5'-P oxidase was reported as a protein component of 30 kDa in molecular mass. However, in 10 to 20% gradient polyacrylamide gel electrophoresis, protein components characterized by molecular masses of 60 kDa, 90 kDa, and 120 kDa, were detected by staining with Coomassie dye for enzymatic activity (Figure 1). The 60 kDa protein band has appeared consistently as the major species among various molecular mass components but the band intensities of other higher molecular mass components vary from preparation to preparation. Results suggest that the existence of higher molecular mass species pyridoxine 5'-P oxidase may be due to aggregation.

Volume III

5

FIGURE 1. 10 to 20% gradient poly aery lamide gel electrophoresis, Lane 1 represents molecular weight markers = IgG, 160,000; aspartate aminotransferase, 110,000; bovine serum albumin, 68,000; and ovalalbumin, 43,000. Lane 2 and 3 represent pyridoxine 5'-P oxidase stained with Coomassie Blue and lane 4, pyridoxine 5'-P oxidase stained for activity.

B. AGGREGATION OF THE OXIDASE Visser et al.12 analyzed the multiple forms of pyridoxine 5'-P oxidase in different pH mediums. At pH 8.4 the high performance liquid chromatography (HPLC) system adapted with the TSK 3000 SW column separated the 60 kDa, and 120 kDa species as separate peaks. As the pH of the elution buffer was reduced from 8.4 to 7.4, the size of the 120 kDa peak became smaller and the 60 kDA peak became larger. Using elution buffer at pH 5.5, the 120 kDa peak completely disappeared from the profile and the 60 kDa peak was the sole peak detected. By quantitating the integrated peak size of the 120 kDa and 60 kDa component at various concentrations of proteins a dissociation constant of 4 X 10~5 M was calculated for the association process: 2 dimer ^± tetramer at pH 7.4. The above results obtained by Visser et al.17 clearly indicate that the aggregation process undergoing pyridoxine 5'-P oxidase is reversible and predominantly pH dependent. This implies that at pH 7.4,

6

Chemistry and Biochemistry of Flavoenzymes

which is the physiological pH of mammalian body system, the oxidase is able to aggregate if the concentration of the enzyme is above 4 x 10~ 5 M. Whether pyridoxine 5'-P oxidase can reach the concentration of 4 x 10~ 5 M under in vivo conditions depends on the existence of compartmentalization within organelles in the cell; and whether or not the aggregated species of the oxidase play a significant role in the metabolism of vitamin B6 under in vivo conditions requires further scientific investigation.

IV. RESOLUTION OF PYRIDOXINE 5'-P OXIDASE Pyridoxine 5'-P oxidase is a flavin mononucleotide (FMN)-dependent enzyme. The FMN group acts as a coenzyme and is absolutely required for catalytic activity. Purified enzymes from different sources11-13'15 have shown that one molecule of FMN binds to one molecule of dimeric enzyme and the coenzyme is not covalently bound to be apoenzyme. Therefore, FMN can be removed from the holoenzyme using various methods of resolution. A. THE KBr METHOD For rabbit liver oxidase, Kazarinoff and McCormick11 reported an 85 to 100% yield of apoenzyme using KBr in the resolution procedure. The first step of the procedure was to dialyze the holoenzyme against four changes of 2 M KBr, 0.1 mM EDTA in 0.1 M potassium acetate buffer at pH 4.0 over a period of 48 h. After dialysis against 2 M KBr, the pH was then readjusted to 7 and KBr was removed by further dialysis against four changes of 0.02 M potassium phosphate, pH 7. Denatured protein was removed by centrifugation at 18,000 Xg for 10 min. The oxidase in the supernatant was in its apo-form. B. THE AMMONIUM SULFATE METHOD For the oxidase from brain, the method using KBr was reported to be noneffective since the enzyme lost more than 90% of its activity within the 48-h dialysis against 2 M KBr.18 Choi et al.15 developed a procedure suitable for the resolution of pyridoxine 5'-P oxidase from both porcine and sheep brains. It began with the addition of solid ammonium sulfate to a solution of holo-oxidase (4 mg/ml) in 0.01 M potassium phosphate at pH 5.5. A final concentration of 1.8 M ammonium sulfate was reached. The mixture was subsequently dialyzed against 100 ml of 1.8 M ammonium sulfate solution with the pH adjusted to 3.5 using HC1. After 4 h of dialysis at 4°C, the dialysis tubing was switched to a beaker containing 2 1 of 0.01 M potassium phosphate, pH 5.5. Dialysis was continued overnight at 4°C with one change of buffer. The end result of this procedure accounted for 95% resolution and 1% loss of the initial oxidase activity after reconstitution with FMN.

V. SUBSTRATE SPECIFICITIES Pyridoxine (pyridoxamine) 5'-P catalyzes the oxidation of pyridoxine 5'-P, pyridoxamine 5'-P and W-Cphosphopyridoxyl) amines. As a substrate of the oxidase, a compound requires the presence of a phosphate group. No oxidation of the unphosphorylated form of substrates or substrate analogues catalyzed by pyridoxine 5'-P oxidase has been observed. Kazarinoff et al.11 summarized the substrate specificities of pyridoxine 5'-P oxidase from rabbit liver in Table 3. Relative maximum velocities using pyridoxine 5'-P and pyridoxamine 5'-P as substrates were found to be the same for the liver enzyme. However, pyridoxine 5'-P was found to be tenfold more active than pyridoxamine 5'-P for the brain oxidase. For phosphopyridoxyl derivatives, the insertion of an -NO2 group into the para-position of the aromatic carboxylic acids attached to the phosphopyridoxyl moiety yields a good substrate for the oxidase whereas an OH inserted into the para-position diminishes the Kcat value (Table 4). It appears that

Volume HI

7

TABLE 3 Activity of Substrate Analogues in Pyridoxamine-P Oxidase System Compound Pyridoxamine 5'-phosphate Pyridoxine 5 '-phosphate NPP-glycineb Pyridoxamine 5'-sulphatec Pyridoxal-P-oximed Pyridoxal-P-O-carboxymethyloximed S'-Homopyridoxine-F* 5 ' -Methy lpyridoxine-Pe a-HPP-ornithinef ct-NPP-lysinef NPP-B-alanineg NPP-L-alanineg NPP-D-alanines NPP- L-ot - aminobuty rate8 NPP-D-a-aminobutyrateg NPP-ct-aminobutyrate8 NPP-L-serine* NPP-L-leucineg NPP-D-leucineg NPP-benzylamineg NPP-L-phenylalanineg NPP-L-tyrosineg NPP-D-tyrosineg NPP-L-tyrptophang a-NPP-diaminodecaneh a

b c d

e f

g h

Apparent KM (xlOA/)

Relative V^ax (%)

1.0 3.0 6.8

100

too

100

Not active

2.1 2.5 0.59

3.1 53 20 11 22 77 9.1 29 77 13 7.5 12.5

3.3 9.5 3.1 160 12 40

10 60 100 20 85 140 130 96 120 130 39 86 120 120 54 57 55 18 20

Apparent KM nd Vmax values determined from Lineweaver-Burk plots. Vmax values given relative to pyridoxamine-P as substrate. Synthesized by the method given in Reference 4. Synthesized by the method given in Reference 19. Synthesized by the method given in Reference 20; the value given is the inhibition constant Kif Data taken from Reference 3. These compounds were kindly provided by Dr, James K. Coward, Department of Pharmacology, Yale University School of Medicine, New Haven, Conn. Data taken from Reference 4. Synthesized by the method given in Reference 21.

From Kazarinoff, M. N. and McCormick, D. B., J, Biol. Chem., 250, 3436, 1975. With permission.

substituents affect the reactivity by their ability to withdraw electrons from the reaction site. Churchich13 showed that substituents with positive CT values (-COOH and -NO2) increase the reactivity of the substrate and substituents with negative a values (-OH) tend to decrease the reactivity of phosphopyridoxyl moiety with the oxidase. Gregory19 reported that Af-acetylphosphopyridoxyl-lysine was the best substrate for the liver oxidase. Using it as a substrate, the oxidase exhibited 75% higher maximum activity than pyridoxamine 5'-P (Table 5). However, its usage as a substrate for the brain enzyme has never been tested.

VI. PROPERTIES OF THE FMN-BINDING SITE A. STUDIES USING FMN ANALOGUES Various analogues have been used by Kazarinoff and McCormick20 to study the coenzyme

Chemistry and Biochemistry of Flavoenzymes TABLE 4 Enzymatic Assays Conducted at pH 8.4 in 0.1 M Pyrophosphate Buffer at 25°C Substrate Pyridoxamine-5-P P-Pyridoxyl-3-alanine P-Pyridoxyl-4-aminobutyrate P-Pyridoxyl-5-aminovalerate P-Pyridoxyl-aniline P-Pyridoxyl-m-aminobenzoate P-Pyridoxyl-p-aminobenzoate Pyridoxine-5-P P-Pyridoxyl-p-aminonitrobenzene P-Pyridoxyl-p-aminophenol

kcat/KM

kca, (s-1)

KM (V-M)

(M~l) (s-1)

0.009 0.04 0.04 0.035 0.01 0.10 0.10 0.09 0.10 0.004

100 35 30 30 1 1 1 13 1

1.1 1.3 1.2 1 1 1 6.9 1

90 x x x x x x x x

103 103 103 104 105 105 103 105

TABLE 5 Substrate Activity of Vitamin B6 Vitamers and Analogues for Pyridoxamine-P Oxidase Compound Pyridoxamine-P Pyridoxine-P Ac-P-Pyx-lysine

(MJW)

Relative Vmax (%)

13 20 65

100 138 175

KM

Note: All reactions were run in 0.2 M Tris-HCl, pH 8.0, at 37°C. Vnym and KM values were calculated from Line weaver- Burk plots. Vmax values are relative to that observed using pyridoxamine-P as a substrate. Reaction mixtures (3.5 ml) contained 0.31 mg of pyridoxamine-P oxidase protein (1.94 units). From Gregory, J. F., III,/. Biol. Chem., 255, 2355, 1980. With permission.

site of rabbit liver pyridoxine 5'-P oxidase. The analogues used were aimed at assessing the coenzyme specificity with respect to its 5'-terminus group, secondary hydroxyl moieties on the No. 10 side chain, and the isoalloxazine ring system. A phosphate group or a similar chemical group attached at the 5'-terminal end of the FMN molecule is definitely required for coenzyme catalysis. Riboflavin and FAD cannot replace FMN as the coenzyme for the oxidase. In addition, compounds such as riboflavin 5'-sulfate or riboflavin 5'-methylphosphate are candidates of coenzyme for pyridoxine 5'P oxidase. (Table 6.) Another interesting aspect of coenzyme function is that the deletion of the hydroxyl group of FMN abolishes its function as coenzyme. Analogues with modification of the hydroxyl groups were tested for coenzyme function and results were also summarized in Table 7. Addition of halogen groups to the isolalloxazine ring of the FMN retains the functionality as coenzyme to a certain limit. Various analogues with modifications on the ring system were shown to be inhibitors of pyridoxine 5'-P oxidase. Results are summarized in Table 8.

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TABLE 6 Coenzymatic Properties of 7,8-Dimethyl-10(l'-D-Ribityl) Isoalloxazines Modified on the 5'-Position3 Flavin Riboflavin 5 '-phosphate Riboflavin 5f-0-methylphosphate Riboflavin 5'-sulfate a

KM X 10*

4.0 12 220

Relative V (%)

100 55 25

Another interesting aspect of coenzyme function is that the deletion of the hydroxyl group of FMN abolishes its function as a coenzyme. Analogues with modification of the hydroxyl groups were tested for coenzyme function and results are summarized in Table 7.

From Kazarinoff, M. N. and McCormick, D. B., Biochim. Biophys. Acta, 359, 282, 1974. With permission.

TABLE 7 Coenzymatic Properties of 7,8-DimethyIIsoalloxa/ine co-Phosphates with Different Side Chains at Position 10 Flavin phosphate

FMN Diacetyl-FMN 3'- to 6'-Hydroxyalkyl-FMNs D, L-Glyceryl-FMN D-Arabityl-FMN D-Lyxityl-FMN L-Lyxityl-FMN D-Dulcityl-FMN

KM X 108

4.0 — — — — — 160 —

Relative V (%)

100 * * * ** ** 78 **

Note: Single asterisk (*) indicates less than 3% activity and no inhibition with respect to FMN at 5.10~ 5 M, Double asterisk (**) indicates less than 3% activity and no inhibition with respect to FMN at 5.10~6 M. From Kazarinoff, M. N. and McCormick, D. B., Biochim. Biophys. Acta, 359, 282, 1974. With permission.

It has been documented in the literature that tryptophan residues play an important role in the binding of flavin molecules to many flavin-binding proteins.21'24 Previous examples have shown that many flavoproteins exhibit low yield of fluorescence due to mutual quenching of flavin associated with tryptophan residue(s). As in pig liver amino acid oxidase,21 a tryptophan residue, is found to be close to its FAD-binding site. Results from chemical modification studies on glucose oxidase from Aspergillus niger22 have also supported the fact that a tryptophan residue quenches the fluorescence of the coenzyme, FAD. Evidence from the analysis of the crystal structures of two flavodoxins has shown that in the protein from Desulfovibrio vulgaris, a tryptophan and a tyrosine residue are located in close proximity to the ring23 and in Clostridium MP flavodoxin,24 a tryptophan and a methionine residue are situated at the side of the flavin molecule. Earlier work of McCormick25 suggested that one of six tryptophan residues in rabbit liver pyridoxine 5'-P oxidase interacts with FMN

10

Chemistry and Biochemistry of Flavoenzymes

FIGURE 2. Time course of inactivation of pyridoxine-5-P oxidase by /V-bromosuccinimide. Experiments conducted at pH 7 in 0.1 m phosphate buffer. Mixing molar ratio of inhibitor to protein 5:1.

at the active site, speculating that an essential tryptophan residue was present at the coenzyme site of the liver oxidase. This proposal had not been thoroughly studied until chemical modification studies were conducted recently in our laboratory. We observed that pretreatment of the apo-oxidase with a fivefold molar excess of A^-bromosuccinimide abolishes its ability to bind FMN and to regain catalytic activity. No inhibition of the catalytic activity of the holoenzyme was observed by addition of fourfold molar excess of A^-bromosuccinimide (Figure 2). The inability of yV-bromosuccinimide to inhibit the holoenzyme provides evidence that FMN prevents the modification of an accessible tryptophan residue. In comparison with a control sample of apo-oxidase, the fluorescence yield of A^-bromosuccinimide-treated apoenzyme was reduced. This reduction of the protein fluorescence was the result of the modification of tryptophan residues in the enzyme. The modification followed the pattern of a pseudo-first order reaction. A second-order rate constant of 8.5 x 104 A/" 1 sec"1 was determined, indicating an exposed tryptophan residue easily accessible to N-bromosuccinimide attack. The attack of the essential tryptophan residue can only be prevented by the presence of FMN. Amino acid analysis of the N-bromosuccinimide-treated apo-oxidase revealed that a tryptophan residue was lost in comparison with the nontreated enzyme (Table 9). B. SPECTROSCOPIC STUDIES Pyridoxine 5'-P oxidase, a dimeric enzyme with two identical subunits, has molecular weights ranging between 54 to 60 kDa. The enzyme binds FMN at a ratio of 1 mol FMN/ 1 mol dimer. After binding to the apoenzyme, the absorption bands of FMN are shifted from 327 and 445 nm maximum to 380 and 448 nm, respectively, and the ratio of A380:A448 for the holoenzyme is significantly higher than the ratio of A372:A445 for free FMN. This phenomenon has been observed in both the liver and the brain enzyme. Since the oxidase utilizes oxygen in the reaction, dithionite can be used to reduce the holoenzyme. In the presence of dithionite, the absorption bands at 380 and 448 nm of the oxidase gradually decrease as a function of dithionite concentrations. The absorption spectrum of the holo-oxidase shows an isosbetic point at 350 nm. Phosphopyridoxaloxime, a potent inhibitor of pyridoxine 5'-P oxidase, was used to probe the active site of this enzyme.13 Upon binding to the holoenzyme, the fluorescence

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TABLE 8 Coenzymatic Properties of lO-(l'-D-Ribitol) Isoalloxazine 5'-Phosphates Modified on the Ring System Flavin phosphate

FMN Iso-FMN 7,8-Dichloro-FMN 3-Methyl-FMN 3-Carboxymethyl-FMN 2-Thio-FMN 2-p-Hydroxyethylimino-FMN 2-AniIino-FMN 20 Morpholino-FMN

K M X 10"

Relative V (%)

4.0 6.0

100 77

4.1*

— 62 — 25 25 — 63

22 4.0*

25 600 ** 300

Note: Single asterisk (*) indicates the value given is k, for these competitive inhibitors. At concentrations of FMN much less than K M , the inhibition appears to be uncompetitive. (Lineweaver-Burk plots for different inhibitor concentrations appear parallel). The double asterisk (**) indicates no coenzyme activity, but FMN activity is inhibited approximately 5% at 5.10"5 M From Kazarinoff, M. N. and McCormick, D. B., Biochim. Biophys. Acta, 359, 282, 1974. With permission.

of this probe was quenched. However, this effect was not observed with apooxidase. The explanation for this is that phosphopyridoxaloxime interacts with the coenzyme at the active site. This interaction can be accounted for either through dipole-dipole coupling process (resonance energy transfer) or intermolecular energy exchange, as both quenching mechanism are operative over a distance of separation of 8 A between the donor/acceptor pair. Another fluorescent probe, to-phosphopyridoxal, can also be used to probe the active site of the oxidase effectively. This competitive inhibitor binds to pyridoxine 5'-P oxidase. Through its interaction with the lysyl residue at the active site, 6/s-phosphopyridoxal covalently attaches to the enzyme after reduction with sodium borohydride. Similar quenching effect of the to-phosphopyridoxyl moieties by the FMN, as previously observed with phosphopyridoxaloxime, was also detected in the holo-oxidase. Using the method of resonance energy transfer to deduce the proximity relationship between the to-phosphopyridoxyl chromophore attached to a specific lysyl residue and the cofactor, FMN, the distance of separation between the donor (to-P-pyridoxal) and the acceptor (FMN) was determined. The estimated distance of 9 A is comparable to the distance of separation between Ppyridoxaloxime and the prosthetic group FMN (8 A). 15 Spectroscopic properties of several FMN analogues bound to pyridoxine 5'-P oxidase were investigated by Merrill et al.26 When 3-deaza-FMN binds to the enzyme at pH 4.7, it exhibits a split of the absorption band at 425 nm. This splitting effect is explained to be due to lower general polarity of the coenzyme site environment. As 3-deazaflavin exists as enol tautomers,24 upon dissociation of a proton (pKa = 5.8), it yields a resonance stabilized anion which is bound preferentially because the protein accepts a cationic group or strong dipole that stabilizes the negative charge. Merrill et al.26 also proposed that the primary association of the protein to the pyrimodinoid edge of the flavin ring may occur proximal to N(l) and oxygen at position 2 of the ring. This is simulated by the binding of 8-hydroxyFMN to the oxidase and the resulting sensitivity of the dissociation constant to alternations

12

Chemistry and Biochemistry of Flavoenzymes TABLE 9 Amino Acid Composition of 60,000Molecular Weight Dimer Amino acid

Apoxidase

Apoxidase with NBS

Histidine Tyrosine Cysteine Tryptophan

8 5 6 4

7.8 4.8 6 3.2

Note: The main amino acid affected by NBS treatment is tryptophan. The decrease of tyrosine and histidine is 2.5%, compared with tryptophan 20%.

at N(l) and oxygen at position 2. Because pyridoxine 5'-P oxidase forms no semiquinone, it is predicted that an ionic interaction will take place at N(l) rather than N(5) as in other flavoproteins.23'24 The presence of a strong dipolar or electrostatic interaction at N(l) which serves to delocalize the dihydroflavin anion leads to the observation of pH-dependent reduced spectrum of oxidase. This pH-dependent reduced spectrum closely resembles a neutral dihydroflavin. The observation of 3-methyl-FMN binding to the oxidase with 30% of its fluorescence quenched, other than 99% for FMN, is consistent with alteration of the flavin-aromatic amino acid complex responsible for quenching. Further support for this proposal is the loss of an 8 nm bathchromatic shift in the 372 nm flavin absorption band upon protein binding of 3-methylflavin compared to FMN. Previous model studies of McCormick25 on flavinaromatic amino acid complexes have established that one spectroscopic result of complex formation is a batchchromatic shift of the shortest absorption band of FMN. The spectra for 3-(carboxymethyl)-FMN rule out the possibility that these effects are due to deprotonation of the N(3)-hydrogen by protein base. 3-(Carboxymethyl)-FMN compensates for loss of the 3-hydrogen by alternate interactions involving the carboxyl group, since its Kd is near that for FMN (Table 10). Most importantly, while the enzyme is completely inactive with this analogue, both the quenching of flavin fluorescence and bathchromatic shift in the 362-nm band are similar to FMN. C. EMISSION ANISOTROPY Visser et al.17 investigated the rotational dynamics of the FMN in pyridoxine 5 '-P oxidase using time-correlated photon counting technique in time-dependent emission anisotropy measurements. Analysis of the anisotropy decay function revealed the presence of two rotational correlation time components shown in Table 11. The predominant contribution of a very short component (0—0.2 ns) was observed regardless of protein concentration. It was concluded that subnanosecond motions of the protein influence the orientation of the prosthetic group at the catalytic site. In addition, subnanosecond motions of the enzyme were suggested to be used to accommodate bulky substrates such as secondary amines carrying aromatic carboxylic acids covalently linked to the phosphopyridoxyl.

VII. PROPERTIES OF THE SUBSTRATE-BINDING SITE A. CHEMICAL MODIFICATION STUDIES Pyridoxine 5'-P oxidase not only catalyzes the oxidation of phosphopyridoxine or phosphopyridoxamine, it also catalyzes the oxidation of various AHphosphopyridoxyl) amines. Various substrate analogues used as competitive inhibitors have been tested by Kazarinoff and McCormick.11

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TABLE 10 Equilibrium Dissociation Constants, K M , and/or Ki for Flavin 5'Phosphates and Apopyridoxamine-5' -Phosphate Oxidase at 25°C Flavin

PH

Riboflavin 5 '-phosphate Riboflavin 5 '-phosphate Riboflavin 5 '-phosphate Riboflavin 5 '-phosphate Riboflavin 5 '-phosphate 1-Deaza-FMN 2-Thio-FMN 2-(methylthio)-FMN 2-(hydroxyethylimino)-FMN 3-Methyl-FMN 3-Deaza-FMN 3-(carboxymethyl)-FMN 5-Deaza-FMN FMN 5 -oxide Iso-FMN 7,8-Dichloro-FMN 8-Hydroxy-FMN 8-Hydroxy-FMN 8-(DimethyIamino)-FMN 8-(Diethylamino)-FMN 8a-S-(7V-acetylcysteinyl)-FMN 8a-S-(mercaptopropionic acid)-FMN 8a-^-(Af-acetylhistidyl)-FMN 8a-7V-( 1 ,6-diamionohexyl)-FMN a b c

d e f

5 6 7 8 9 8 7, 8 7, 8

8 8 8 8 8 8 8 8 5 8 8 8 8 8 8 8

Kd» (nM)

30 16 11 13 21 190 520e

900* 260 180 21 20 260f

AF 0 c (kcal/mol)

K M or

V

-10.3 -10.6 -10.9 -10.8 -10.5 -9.2 -8.6 -8.3

20, 40d

250"

(-7.1) -9.0 -9.2 -10.5 -10.5 -9.0 (-9.9) (-10.1) -9.8 -10.3 -9.7 -9.5 (-9.8) (-9.6) (-8.2) (-8.3)

6000d 220d

40d 20d 60d 41d

62 29 76 120 70 90 1000

800

Determined by fluorescence titration as described in the text. Determined by Line weaver- Burk plots. The standard, molar, free energy of binding was calculated from AF° = -RT In Ka; Ka = K d - 1 . Unitary free energies can be obtained by subtracting 2.4 kcal/mol for the cratic entropy value of 8 eu. Estimates for AP based on KM or Kj are indicated by parentheses. Values from Kazarinoff and McCormick (1975). Measured at pH 7 to minimize hydrolysis to FMN. An approximate value; the analogue was contaminated with lumichrome.

TABLE 11 Emission Anisotrophy Decay of Bound FMN

o,

Protein cone. (\*M)

Emission wavelengths (nm)

Pi

2.8 2.8 28 28 28

498 544 544 544 498

0.42 0.36 0.31 0.32 0.36

f*2

0.05

0 0.03 0.02 0.04

(ns)

(ns)

02

Temp. (°C)

0.15 0.19 0.21 0.36 0.21

0.49

19 19 19 4 19

0 4.79 5.70 1.68

Chemical modification studies have shown that amino acids, such as histidine, arginine and lysine are present at the active site of pyridoxine 5'-P oxidase. All three amino acid residues are implicated to be essential for catalytic activity. In the studies involved in the elucidation of essential histidyl residue in pyridoxine 5'P oxidase, Horiike et al.27 found that 1.29 mM diethylpyrocarbonate reacted with a total of

14

Chemistry and Biochemistry of Flavoenzymes

4 histidyl residues of the enzyme. One of the four histidyl residues was kinetically shown to be related with enzymatic activity. This finding has been consistent with results obtained with other oxidases. 2833 This critical histidyl residue in the oxidase may participate in the binding of substrate and product or as a base to abstract a proton from the substrate or both.34 Phenylglyoxal, 2,3-butadione and 2,4-pentadione inhibit pyridoxine 5'-P oxidase activity.35 The inhibition of phenylglyoxal follows pseudo-first-order kinetics, the rate of which is a function of reagent concentration. The second-order rate constants for apoenzyme and holoenzyme are 3.7 and 11.1 A/" 1 min" 1 respectively. The presence of FMN enhances the inactivation by threefold indicating that its presence causes the arginyl residue to be more accessible to the modifying reagent. This finding implicates that binding of the FMN to the protein results in some sort of conformational change required for substrate binding as bulky phosphopyridoxyl derivatives are also substrates of the oxidase. 2,3-Butadione also inhibits pyridoxine 5'-P oxidase. The inhibition is observed to be augmented by borate buffer and reversible by the removal of borate. Pentadione inactivates the oxidase. Treatment of pentadione inactivated enzyme with hydroxylamine does not restore enzymatic activity, indicating that the inactivation is due to modification of arginyl but not lysyl residues. Choi et al.35 showed that phenylglyoxal-treated oxidase retained the affinity for FMN, and that the coenzyme did not provide any protection from the phenylglyoxal inactivation. Apparently, the product/inhibitor, pyridoxal 5'-P provides effective protection for the oxidase from phenylglyoxal inactivation and also the modification of one of the six arginyl residues in the enzyme, indicating the presence of an essential arginyl residue in the substrate-binding site. Besides the presence of histidyl and arginyl residues, lysine is another likely candidate at the active site of pyridoxine 5'-P oxidase. Pyridoxal 5'-P has been shown to be an inhibitor of the enzyme with a K{ of 6 |mM. Churchich13 observed that inhibition of the oxidase activity by pyridoxal 5'-P required a preincubation period of approximately 15 min at 30°C. Bispyridoxal 5'-P (K; = 2 (jiM) is a more potent inhibitor than pyridoxal 5'-P. Similar to pyridoxal 5'-P, b/s-phosphopyridoxyl reacts covalently with one lysyl residue of the oxidase; after sodium borohydride reduction, one molecule of to-phosphopyridoxyl is incorporated per dimeric molecule of pyridoxine 5'-P oxidase. Choi et al.15 utilized 6/s-phosphopyridoxyl to probe the active site of the oxidase. After tryptic digestion of the oxidase modified with fc-phosphopyridoxyl, a single peptide marked with the probe was isolated and sequenced. The amino acid sequence of this active site fragment is shown. Leu-Tyr-Val-Gly-Thr-Asp-Glu-Ala-Phe-Glu-Ser-Lys-Lys 1 Pyr The modified lysyl residue has been suggested to be involved in the regulation of enzyme activity. Product inhibition by pyridoxal 5'-P has been observed in pyridoxine 5'-P oxidase and has been shown to be a key mechanism in the regulation of this enzymatic activity.37 B. KINETIC STUDIES Oxidation catalyzed by pyridoxine 5'~P (PNP) oxidase proceeds via a binary complex (ping-pong) mechanism with pyridoxine 5'-P and a ternary (sequential) mechanism with pyridoxamine 5'-P (PMP). In both cases, oxygen molecules function as the electron acceptor. Results from steady-state kinetics38 showed that parallel Lineweaver Burk plots were obtained with oxygen and pyridoxine 5'-P. Choi et al.38 indicated that the oxidase catalyzed oxidation of pyridoxine 5'-P via "ping-pong" mechanism because of the observation of competitive inhibition as further evidence. Kinetic data were applied to Equation 1 for the reaction

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FIGURE 3. Double reciprocal plot for reaction of oxidase with PNP. Oxygen concentrations (millimolar) were 0.13 (•), 0.26 (•), 0.78 (A), and 1.3 O- The inset is a secondary plot of intercepts vs. the reciprocals of the oxygen concentration; TN is turnover number. (From Choi, J. D., Bowers-Kromro, D. M., Davis, M. D., Edmonson, D. E., and McCormick, D. B., J. BioL Chem., 258, 840, 1983. With permission.)

reflected by the parallel line pattern:

where Ks and K02 are the Michaelis constants for substrate and oxygen, respectively. Equation 1 can be modified for competitive substrate inhibition as Equation 2.

where K{ is the dissociation constant for the nonproductive complex. The presence of the [SJ/Kj term in this equation is reflected in the curvature of the double reciprocal plots (Figure 3). At a low level of substrate, the substrate inhibition term [S]/Kt is negligible and linear parallel lines are expressed in the double reciprocal plots. In the region of excess substrate inhibition, the KS/[S] and [S]/Kj terms to reciprocal velocity are dependent on O2 concentration.

16

Chemistry and Biochemistry of Flavoenzymes

FIGURE 4. A scheme representing the mechanistic pathway of oxidation catalyzed by pyridoxine 5'-P oxidase.

Plots of reciprocal velocity vs. pyridoxine 5'-P concentration are linear at high substrate levels and cross at a point which gives a dissociation constant (K;) of 50 jxM and maximum turnover of 40 min" 1 . A secondary plot of the slope vs. 1/[O2] is apparently linear which is consistent with pyridoxine 5'-P exerting its inhibitory effect at high substrate levels and probably reflects the formation of a nonproductive complex between substrate and reduced enzyme. A similar interpretation has been previously suggested for the reaction of L-amino acid oxidase with phenylalanine.34 The studies with PMP also showed a family of parallel lines but no significant substrate inhibition. Choi et al.38 explained that the release of product from its complex with reduced enzyme was significantly slower (15-fold) than maximum turnover number suggesting that oxygen probably reacts with the reduced enzyme-product complex before product release. Since O2 reaction with the enzyme-imine complex is much faster than catalytic turnover, the dissociation of the imine (pyridoxal) from the oxidized enzyme is suggested to constitute the rate-limiting step in catalysis. The following scheme formulated by Choi et al.38 (Figure 4) showed the relative magnitude of k4[O2] vs. k'5 dictated the choice of pathways. It was suggested if k4[O2] — k'5, both paths then might operate, but double reciprocal plots would be curved as found for L-amino acid oxidase with phenylalanine.34 Deuterium isotope effect and anaerobic stopped-flow data have suggested that the oxidase reduction is rate-limiting for the reaction with pyridoxine 5'-P but not with pyridoxamine 5'-P. Enzyme reduction with PMP is about 100-fold faster than that with pyridoxine 5'-P and shows a similar magnitude of kinetic isotope effect in the reduction half-reaction. The 100-fold increase in rate of flavin reduction by pyridoxamine 5'-P as compared with pyridoxine 5'-P shows that electron transfer from the primary amine is a more facile process than from a primary alcohol. C. STUDIES ON STEREOSPECIFICITY Pyridoxine 5'-phosphate oxidase is nonstereospecific with respect to the hydrogen removal from the substrate, pyridoxamine. Bowers-Kromro and McCormick39 demonstrated the nonstereospecificity of the oxidase by preparing tritium labeled [4'-3HR] pyridoxamine5'-phosphate from either aspartate aminotransferase or glutamate decarboxylase. Results showed that about half (47%) of the initial tritium racemically labeled and pro-R labeled pyridoxamine 5'-phosphate was lost as a result of oxidase catalyzed removal of one of the 4'-methylene hydrogens. Since either of the 4'-methylene hydrogens on the aminoethyl of pyridoxamine 5'-phosphate can be removed during oxidation, Bowers-Kromro and McCormick39 proposed that the cleft within which FMN and the substrate fit must be quite

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open to solvent and may allow for differential approach of substrate. It was suggested that the abstracting base may be equidistant from the two hydrogens. Although only pyridoxamine 5'-phosphate was used as substrate for the demonstration of stereospecifity of pyridoxamine 5'-phosphate oxidase, it is assumed that the removal of hydrogen from pyridoxine 5-phosphate may follow the same pattern. Hypothetically, pyridoxine 5'-phosphate oxidase is an enzyme which exhibits overall lack of stereospecificity for hydrogen abstraction from substrate only because of "randomization of label" in the microenvironment of the active site during oxidation.

VIII. REGULATION A. INTERACTION WITH PYRIDOXAL KINASE The synthesis of pyridoxal-P (PLP) from pyridoxine (PN) and pyridoxamine (PM) requires the joint action of pyridoxal (PL) kinase and pyridoxine-P (pyridoxamine-P) oxidase. Since pyridoxine-P oxidase contributes to the formation of pyridoxal-P, it is evident that some regulatory mechanisms have to exist for adequate formation of pyridoxal-P in cells. The inhibitory effect exerted by pyridoxal-P on the oxidase suggests that the accumulation of pyridoxal-5-P could, in principle, prevent its further formation from pyridoxine and pyridoxamine. In addition, it is possible that accumulation of pyridoxal-5-P is facilitated by macromolecular aggregates which restrict the out diffusion of the metabolites. Kwok and Churchich12 used resonance energy transfer and transient kinetic experiments to elucidate the existence of this macromolecular aggregate between pyridoxal kinase and pyridoxine-P oxidase from the brain. It was demonstrated that the relationship between the reciprocal transient time (I/ y) and the molarity of pyridoxine-5-P oxidase is linear until pyridoxine-P oxidase concentration has been increased up to 6 (xM (Figure 5). The transient times (y) observed at oxidase concentration of 6, 7, 8, and 9 |JiM are considerably shorter than the values calculated from Equation 3 developed by Hess and Wurster.40

The regulation of vitamin B6 levels in tissues is dependent on a number of factors. They include vitamin uptake and metabolism by the liver,41 uptake mechanisms,42 binding to proteins, and activities of enzyme involved in the metabolism of the vitamin.43"45 It is apparent that pyridoxal kinase, not only converts PN, PL, and PM to the phosphorylated forms but also that this enzyme plays a trapping role whereby diffusible nonphosphorylated vitamin forms become intracellularly phosphorylated and poorly diffusible across the cell membrane.4 In addition, the conversion of PNP and PMP to PLP has been reported to be regulated by product inhibition of PNP oxidase.37 It is apparent that this is the only role played by PNP oxidase since the uptake of pyridoxine in liver cells is dependent on the activity of pyridoxal kinase, but not of the oxidase. B. PYRIDOXAL 5'-P OXIDASE IN TUMORS Experimental evidence has shown that pyridoxine 5-P oxidase is a key enzyme in the formation of pyridoxal 5'-P. Against this background, Thanassi et al.46 have found that oxidase activity in Morris hepatoma 7777 approaches zero, relative to the host and control liver. The finding is against the initial speculation that pyridoxine 5'-P oxidase activity should be higher in tumor than in normal tissue due to a faster growth rate in tumor tissue. Experiments conducted by Thanassi et al.46 and Nutter et al.47 have shown that the absence of oxidase activity in a tumor is not caused by insufficient FMN, presence of an inhibitor

18

Chemistry and Biochemistry of Flavoenzymes

FIGURE 5. Relation between the reciprocal transient time (T) and the molarity of pyridoxine-5-P oxidase. Enzymatic assays performed in 0.1 M potassium phosphate (pH 7) containing ATP (0.1 mM), pyridoxine (0.1 mA/), and ZnCl2 (0.015 mM). The concentration of pyridoxal kinase(0.25 p,M) was kept constant, whereas pyridoxine-5-P oxidase was varied from 2 to 9 \iM. (From Kwok, F. and Churchich, J. E., J. Biol. Chem., 255, 882, 1980. With permission.)

in the tumor, a missing or defective enzyme or relocation of the enzyme. It appears that the absence of the oxidase activity is a common feature in tumor tissue because Thanassi et al.46 have also found the absence of oxidase in other tumors; such as, 9618 A2 and 7288 C Morris hepatomas, a large bowel tumor from a Fisher rat treated with dimethylhydrazine, a spontaneous B16 melanoma from BDF1 mouse, a Walker 256 mammary carcinoma from a Sprague-Dawley rat, and a spontaneous liver epithelial cell tumor from a Wistar rat. The lack of pyridoxine 5'-P oxidase in solid tumors suggests that tumors are incapable of metabolizing nutritionally active forms of vitamin B I6 to pyridoxal 5'-P and may be dependent on efficient uptake and binding of pyridoxal and pyridoxal 5'-P. The possibility of direct pyridoxal 5'-P uptake by hepatoma ascitic cells has been proposed by Ito et al.48

REFERENCES 1. Pogell, B. M., Enzymatic oxidation of pyridoxamine-5-P to pyridoxal-5-P in rabbit liver, /. Biol. Chem., 232, 761, 1958. 2. Wada, H. and Snell, E. E., The enzymatic oxidation of pyridoxine and pyridoxamine-phosphates, J. Biol. Chem., 236, 2089, 1961. 3. Black, A. L., Guirard, B. M., and Snell, E. E., Increased muscle phosphorylase in rats fed high levels of vitamin B6, J. Nutr., 107, 1962, 1977.

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4. Lumeng, L., Lui, A., and Lim, T. K., Plasma content of B6 vitamers and its relationship to hepatic vitamin B6 metabolism, J. Clin. Invest., 66, 688, 1980. 5. Rasmussen, K., Barsa, P., and McCormick, D. B., Pyridoxamine-5-P oxidase activity in rat tissues during development of riboflavin, Proc. Soc. Exp, Biol, Med., 161, 527, 1979. 6. Takeuchi, F., Tsubouchi, Ryoko, and Shibata, Y., Effect of tryptophan metabolites on the activities of rat liver pyridoxal kinase and pyridoxamine-5-P oxidase in vitro, Biochem. J., 227, 537, 1985. 7. Tsuge, H., Itoh, K., Akatsuka, F., Okada, T., and Ohashi, K., Inactivation of pyridoxamine-5-P oxidase by aliphatic primary amines, Biochem. Intern., 6, 743, 1983. 8. Tsuge, H., Kuroda, Y., Iwamoto, A., and Ohashi, K., Partial purification and property of pyridoxaimine5-P oxidase from wheat seedlings, Arch. Biochem. Biophys., 217, 479, 1982. 9. Nutter, L. M., Meister, N. T., and Thanassi, J. W., Absence of pyridoxine-5-P oxidase in Morris hepatomma 7777, Biochemistry, 22, 1599, 1983. 10. Merrill, A. H., Ka/arinoff, M. N., Tsuge, H., Horiike, K., and McCormick, D. B., Pyridoxamine5-P oxidase from rabbit liver, Methods EnzymoL, 62, 568, 1979. 11. Kazarinoff, M. N. and McCormick, D. B., Rabbit liver pyridoxamine-5-P oxidase, J. Biol. Chem., 250, 3436, 1975. 12. Kwok, F. and Churchich, J. E., Interaction between pyridoxal kinase and pyridoxine-5-P oxidase, J. Biol. Chem., 255, 882, 1980. 13. Churchich, J. E., Brain pyridoxine-5-P oxidase, dimeric enzyme containing one FMN site, Eur. J. Biochem., 138,327, 1984. 14. Bowers-Komro, D. M., Hagen, T. M., and McCormick, D. B., Modified purification of pyridoxamine5-P oxidase from rabbit liver, Methods EnzymoL, 122, 116, 1986. 15. Choi, S. Y., Churchich, J. E., Zaiden, E., and Kwok, F., Brain pyridoxine-5-P oxidase, modulation of its catalytic activity by pyridoxal-5-P, J. Biol Chem., 262, 12013, 1987. 16. Kerry, J. A., Rhode, M., and Kwok, F., Brain pyridoxal kinase purification and characterization, Eur. J. Biochem., 158, 581, 1986. 17. Visser, A. J. W., Kwok, F., and Churchich, J. E., Time-resolved flavin fluorescence of pyridoxine-5P oxidase from sheep brain, in Flavins and Flavoproteins, Edmonson, D. E. and McCormick, D. B., Eds., New York, Walter de Gruyter, Berlin, 1987, 217. 18. Zaiden, E., Choi, S. Y., and Kwok, F., Pyridoxine-5-P oxidase isolated from sheep brain, Proc. Aust. Biochem. Soc. V., 19, 23, 1987. 19. Gregory, J. F., Ill, Effects of e-pyridoxylysine and related compounds on liver and brain pyridoxal kinase and liver pyridoxamine-5-P oxidase, /. Biol. Chem., 255, 2355, 1980. 20. Kazarinoff, M. N. and McCormick, D. B., Specificity of pyridoxamine-5-P oxidase for flavin phosphates, Biochim. Biophys. Acta, 359, 282, 1974. 21. Tu, S. C. and McCormick, D. B., Conformation of D-amino acid oxidase, Biochemistry, 13, 893, 1974. 22. Tsuge, H. and Mitsuda, H., Studies on the molecular complex of flavins, J. Biochem. (Tokyo), 73, 109, 1973. 23. Waterpaugh, K. D., Seiker, L. C., Jensen, L. H., LeGall, J., and Dubourdieu, M., Structure of the oxidized form of a flavoprotein at 2.5 A resolution, Proc. Natl Acad. Sci. U.S.A., 69, 3185, 1972. 24. Anderson, R. D., Apgar, P. A., Burnett, R. M., Darling, G. D., LeQuese, M. E., Mayhew, S. G., and Ludwig, M. L., Structure of the radical form of Clostridial flavodoxin, Proc. Natl. Acad. Sci. U.S.A., 69,3189, 1972. 25. McCormick, D. B., Photochemistry of flavins, Photochem. Photobiol., 26, 169, 1977. 26. Merrill, A. H., Kasai, S., Matsui, K., Tsuge, H., and McCormick, D. B., FMN binding by pyridoxamine-5-P oxidase, Biochemistry, 18, 3635, 1979. 27. Horiike, H., Tsuge, H., and McCormick, D. B., Evidence for an essential histidyl residue at the active site of pyridoxamine-5-P oxidase from rabbit liver, J. Biol. Chem., 254, 6638, 1979. 28. Thome-Beau, F., Le-Thi-Lan, Olomucki, A., and Thoai, N., Essential histidyl residues in arginine oxygenase, Eur. J. Biochem., 19, 270, 1971. 29. Hiramatsu, A., Tsurushiin, S., and Yasunobu, P. A., Evidence for essential histidyl residues in bovine liver monoamine oxidase, Eur. J. Biochem., 57, 587, 1975. 30. Choong, Y. S., Shepherd, M. G., and Sullivan, P. A., Modification of lactate oxidase with diethyl pyrocarbonate, Biochem. J., 165, 385, 1977. 31. Steenkamp, D. J., Schabort, J. C., Holzapfel, C. W., and Ferreira, N. P., The role of essential histidyl residues in the mechanism of catalysis, Biochim. Biophys. Acta, 358, 126, 1974. 32. Hitwatshi, A., Ichikawa, Y., Yamano, T., and Maruya, N., Essential histdyl and cysteinyl residues of NADPH ADRENODOXIN reductase, Biochemistry, 15, 3091, 1976. 33. Boggaram, V. and Mannervik, B., An essential histidyl residue in the catalytic mechanism of glutathione reductase, Biochem. Biophys. Res. Commun., 83, 558, 1978. 34. Bright, H. J. and Porter, D. J. T., In Enzymes, 12, 421, 1975 35. Choi, D. J. and McCormick, D. B., Roles of arginyl residues in pyridoxcamine-5-P oxidase, Biochemistry, 20, 3722, 1981.

20

Chemistry and Biochemistry of Flavoenzymes 36. Arnone, A., Bier, C. J., Cotton, F. A., Day, V. W., and Hazen, A., A high resolution structure of an inhibitor complex of nuclease, J. Biol. Chem., 246, 2302, 1971. 37. Merrill A. H., Horiike, K., and McCormick, D. B., Evidence for the regulation of pyridoxal-5-P formation in liver by pyridoxamine-5-P oxidase, Biochem. Biophys. Res. Commun., 83, 984, 1983. 38. Choi, J. D., Bowers-Kromro, D. M., Davis, M. D., Edmonson, D. E., and McCormick, D. B., Kinetic properties of pyridoxamine-5-P oxidase from rabbit liver, J. Biol. Chem., 258, 840, 1983. 39. Bowers-Kromro, D. M. and McCormick, D. B., Pyridoxamine-5-P oxidase exhibits no specificity in prochiral hydrogen abstraction from substrate, J. Biol. Chem., 260, 9580, 1985. 40. Hess, B. and Wursterm, B., Transient time of the pyruvate kinase lactate dehydrogenase system of rabbit muscle in vitro, FEBS Lett., 9, 73, 1970. 41. Mehansho, H., Buss, D. D., Ha mm, M. W., and Henderson, L. H., Transport and metabolism of pyridoxine in rat, Biochim. Biophys. Acta, 631, 112, 1980. 42. Spector, R. and Greenwald, L. L., Transport and metabolism of vitamin B6 in rabbit brain and choroid plexus, J. Biol. Chem., 253, 2373, 1978. 43. Snell, E. E. and Haskell, B. E., Function in transamination and decarboxylation reactions, in Comprehensive Biochemistry, Florkin, M. and Stoltz, E. H., Eds., Vol. 15, 138, 1971. 44. Lumeng, L. and Li, T. K., Characterization of the pyridoxal-5-P and pyridoxamine-5-P hydrolase activity in rat liver, J. Biol. Chem., 250, 8126, 1975. 45. Meister, N. T. and Thanassi, J. W., Pyridoxine-kinase, pyridoxine-5-P phosphate and oxidase in B6 deficient rat liver and brain, J. Nutr., 110, 1965, 1980. 46. Thanassim, J. W., Nutter, L. M., Meister, N. T., Commers, P., and Chui, J. F., Vitamin B6 metabolism in Morris hepatomas, J. Biol. Chem., 256, 3370, 1981. 47. Nutter, L. M., Mesister, N. T., and Thanassi, J. W., Absence of pyridoxine-5-P oxidase in Morris hepatoma, 7777, Biochemistry, 22, 1599, 1983. 48. Ito, K., Nakahasa, K., and Lakomoto, Y., Gann., 55, 379, 1964.

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Chapter 2

XANTHINE OXIDASE, XANTHINE DEHYDROGENASE, AND ALDEHYDE OXIDASE Russ Hille

TABLE OF CONTENTS I.

Introduction

22

II.

Xanthine Oxidase A. Physicochemical Properties B. Thermodynamics and Kinetics of Internal Electron Transfer C. The Flavin Site and the Oxidative Half-Reaction D. The Molybdenum Center and the Reductive Half-Reaction

22 23 28 37 41

III.

Xanthine Dehydrogenase A. Physicochemical Properties B. Kinetic Properties C. The Dehydrogenase/Oxidase Interconversion D. Molecular Genetics

51 52 53 55 57

IV.

Aldehyde Oxidase A. Physicochemical Properties B. Kinetic Properties

58 59 60

V.

Concluding Remarks

61

Acknowledgments

62

References

62

22

Chemistry and Biochemistry of Flavoenzymes

L INTRODUCTION Among those enzymes capable of carrying out the hydroxylation of either aliphatic or aromatic substrates, the molybdenum hydroxylases possess a unique reaction stoichiometry. Unlike systems utilizing heme, flavin or biopterin cofactors, the molybdenum enzymes use water as the source of oxygen to be incorporated into substrate rather than molecular oxygen. In the former cases the thermodynamic favorability of the overall reaction that is derived from the reduction of one of the atoms of dioxygen to water (utilizing external reducing equivalents, generally in the form of NAD(P)H) is used to generate a highly activated intermediate in the reaction cycle: a Fe(IV) = O in the case of the heme systems, 4a peroxides in the flavin (see Miiller, Vol. I of this series), and biopterin systems. By contrast, the thermodynamic driving force for the molybdenum-catalyzed reactions is derived entirely from the oxidation of substrate itself and there is at present no evidence for an activated intermediate of the type encountered in the other systems. Thus, it is clear that the molybdenum hydroxylases represent a unique solution to the general problem of oxygen activation in biology. This review concerns itself with those complex molybdoflavoproteins catalyzing the hydroxylation of xanthine and related heterocycles as well as a variety of aldehydes. In addition to those aspects of protein function relevant to the flavin centers of the enzymes, the structure and function of the molybdenum center are also discussed. An effort has been made in the present review to provide on the one hand an appreciation of the extensive body of work dealing with the physicochemical properties of the enzymes, and at the same time give a critical overview of much of the exciting work that has taken place in the past 5 years. The reader is referred to other recent reviews for different perspectives of the field. 1 - 2 Xanthine oxidase, xanthine dehydrogenase, and aldehyde oxidase each contain a molybdenum center, two iron-sulfur centers of the 2Fe/2S variety, and noncovalently bound flavin adenine dinucleotide (FAD). Associated with the molybdenum center of each enzyme is a pterin cofactor that appears to be directly coordinated to the active-site molybdenum atom via side-chain thiol groups. 3 It is to be emphasized that these enzymes differ principally in their substrate specificities (which overlap appreciably), and the conclusions drawn with regard to the function of one may reasonably be expected to apply to the others. A further common aspect of these enzymes (which they share with all other molybdenum-containing hydroxylases) is that the reductive and oxidative half-reactions of the catalytic cycle are spatially separated in the enzymes, the former taking place at the molybdenum center and the latter at the flavin. For this reason, internal electron transfer between the molybdenum and flavin (and undoubtedly involving the iron-sulfur centers as well) is an integral aspect of the overall catalytic sequence. A total of six reducing equivalents are required to bring about full reduction of these enzymes: two each at the flavin and molybdenum (in cycling between the VI and IV oxidation states), and one each at the iron-sulfur centers. As a consequence of the considerable capacity of the enzymes for reducing equivalents, both reductive and oxidative half-reactions are complicated by the fact that they necessarily involve the reaction of multiple substrate molecules with a given enzyme molecule to go to completion.

II. XANTHINE OXIDASE Historically, the most extensively studied of the proteins under consideration has been xanthine oxidase. First purified in 1924,4 this enzyme has been the focus of much attention for the succeeding interval of time, and a wealth of physicochemical information with regard to it has accumulated.1-5-6 The enzyme catalyzes the hydroxylation of xanthine at the C(8) position to form uric acid, utilizing molecular oxygen as the acceptor of the reducing equivalents thus generated (see Scheme 1).

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SCHEME 1 . The overall stoichiometry of the reaction catalyzed by xanthine oxidase.

Work in the past ten years has focused on the structure and function of the several centers in the protein, and their respective roles in overall catalysis. The discussion below begins with a description of the physicochemical properties of xanthine oxidase, particularly those aspects which have been used to examine the function of the enzyme, and proceeds to an account of our present understanding of electron transfer within the enzyme and the mechanism of action of both the oxidative and reductive half- reactions. A. PHYSICOCHEMICAL PROPERTIES Xanthine oxidase as isolated from cow's milk is a homodimer of Mr 300,000.5 Each subunit is catalytically independent of the other and contains the full complement of redoxactive sites: molybdenum,7 two 2Fe/2S centers,8 and FAD.9 The two iron-sulfur centers are designated I and II,8 and are distinguishable on the basis of the g values and temperaturedependence of their EPR signals (see below). The two most commonly employed enzyme preparations utilize either selective precipitation of enzyme by salicylate (the product of enzymatic action on salicylaldehyde, which is empirically found to stabilize enzyme activity) and gel exclusion chromatography10 or selective binding to calcium phosphate11 (both batch and column procedures being used). Both preparations begin with unpasteurized buttermilk (i.e., the liquid remaining when cream is churned to make butter) and yield an enzyme of comparable purity and activity. The latter preparation is used routinely in the author's laboratory with the modification that raw milk rather than buttermilk is used. A typical yield from 60 1 of unpasteurized milk is 2.5 g of enzyme, having a specific activity of 70 to 80% of the maximum theoretical value (see below) . When raw milk is used as the starting material , it is typically necessary to include a carboxymethyl-cellulose chromatography step in the purification to remove lactoperoxidase which otherwise contaminates the xanthine oxidase product.12 In preparations from unpasteurized raw milk (as opposed to pasteurized cream), it is not necessary to incubate with pancreatin as has been done routinely in preparations using buttermilk as the source.10-11 This bovine pancreatic extract possesses a lipase activity that facilitates the dissociation of the xanthine oxidase from lipid vesicles in cream, but also has protease activity which must be inhibited by the addition of phenylmethylsulfonylfluoride to the incubation. Despite the presence of the inhibitor, selected proteolysis of xanthine oxidase in the course of the incubation has been reported to give rise to specific artifacts in the preparation.13 Omitting the pancreatin incubation has no adverse effect on the yield or quality of enzyme, although the physical properties of the crude enzyme are such that the preparation is substantially more tedious owing to the presence of a persistent opalescent turbidity (R. Hille, R. Stewart, V. Massey, unpublished). All preparations of xanthine oxidase as conventionally isolated contain varying amounts (typically 20 to 30%) of enzyme existing in a specific inactive form which lacks a labile sulfur atom at the molybdenum center.14 This so-called desulfo form of xanthine oxidase can be obtained from the active enzyme by reaction with cyanide to release one equivalent of thiocyanate. This reaction that is partially reversible by incubation of desulfo xanthine

24

Chemistry and Biochemistry of Flavoenzymes

oxidase with sulfide, although no more than 50% of maximal activity is recovered by this procedure.14 It has not yet proven possible to quantitatively and reproducibly reactivate desulfo xanthine oxidase by reaction with any of several thiol-containing compounds under a variety of procedures. Two different affinity procedures for the separation of the functional and nonfunctional, desulfo forms of xanthine oxidase have been described, however. The first is based on the very tight binding of reduced enzyme to an immobilized pyrazolopyrimidine affinity column, 15 while the second relies on the ability of the pyrazolopyrimidine alloxanthine16 to preferentially bind to reduced, functional xanthine oxidase and thereby block its binding to an immobilized folate affinity column17 (which otherwise binds both functional and nonfunctional enzyme). The latter procedure is the more convenient and reproducible, even though it is necessary to reactivate the functional, alloxanthine-blocked enzyme eluted from the folate affinity column by incubation with potassium ferricyanide in order to recover enzyme activity. This reactivation is best carried out in the dark to prevent the photodecomposition of the ferricyanide, which would reasonably be expected to release cyanide into the enzyme solution. The steady-state kinetics of xanthine oxidase give a parallel pattern of lines in a Lineweaver-Burk plot of l/V max vs. l/[xanthine] at varying oxygen concentrations.18'19 Given that the reductive and oxidative half-reactions of the catalytic cycle take place at different sites on the enzyme, however, this pattern is to be expected20 and does not necessarily reflect the absence of a ternary complex of enzyme, xanthine, and oxygen in the catalytic cycle. The optimal pH for enzyme activity is 8.5, 21 but the effect of pH on the KM for xanthine is such that kcat/KM achieves its maximum value of 107 M"^" 1 at pH 7.0 (derived from data in reference 19; since the rate-limiting step in catalysis is thought to be in the reductive half-reaction of the catalytic cycle,22 it is the K M for xanthine rather than oxygen that is the relevant parameter in calculating kcat/KM). It is to be noted that the pH-dependence study was carried out in air-equilibrated buffers (approximately 200 jxM in oxygen) and was not saturated with respect to oxygen (the KM for oxygen being 125 (o-M at pH 8.5). The xanthine KM values used above are thus only apparent KM values. For xanthine oxidase, kcat/KM exhibits a modest kinetic isotope effect of 1.7 with 8-deuteroxanthine.22 This figure is independent of pH above 8.5 and its low value has been interpreted to indicate that product release is predominantly rate-limiting in the reductive half-reaction, and therefore also in the overall catalytic cycle.23 D'Ardenne and Edmondson23a have recently examined the kinetic isotope effect of xanthine oxidase and xanthine dehydrogenase using both deuterated and tritiated substrate, finding a tritium isotope on kcat/KM of 3.6 for both enzymes. The intrinsic deuterium isotope effect for C8-H bond cleavage was determined to be 7.4 for xanthine oxidase and 4.2 for xanthine dehydrogenase. From this data, it was estimated that the C-H bond cleavage step was approximately 75 times faster than kcat in the case of the oxidase and 10 times greater in the case of the dehydrogenase. These results are consistent with rate-limiting product dissociation in the overall reaction.23 Given the number of redox-active centers in xanthine oxidase, it is hardly surprising that a variety of spectroscopic probes have been used to advantage in studying the enzyme. Figure 1A shows the visible absorption spectra for oxidized and dithionite-reduced enzyme. The oxidized enzyme exhibits absorption maxima at 450 nm (extinction of 37.8 mA/~ 'cirr l ) and 325 nm (46.6 mM^crn" 1 ). The enzyme has an extinction change at 450 nm upon reduction of 26.6 mAf^cm" 1 , 11 of which 14.4 mAf ^cm^ 1 is due to reduction of the two iron-sulfur centers and 12.2 inM^cm" 1 to reduction of the flavin.24 At 550 nm, the entirety of the 6.7 mM^crn" 1 extinction change is due to reduction of the two iron-sulfur centers. These latter values have been determined from a comparison of the spectral properties of the native enzyme with enzyme that has had the flavin removed by incubation with CaCl2.24 The flavin semiquinone that is observed at intermediate levels of enzyme reduction in the course of reductive titrations is the neutral semiquinone. This form has long wavelength

FIGURE 1.

The visible absorption (A) and circular dichroism (B) spectra of milk xanthine oxidase. Solid lines, oxidized enzyme; dashed lines, dithionite-reduced enzyme.

26

Chemistry and Biochemistry of Flavoenzymes

absorbance extending beyond 650 nm, and an extinction coefficient at 600 nm of approximately 4.5 mM^cm" 1 . 19 ' 25 Below pH 7.0, the semiquinone accumulates sufficiently in reductive titrations that the addition of the first few aliquots of reducing equivalents results in an absorbance increase above 550 nm. Transient accumulation of the neutral semiquinone also results in an overshoot of the baseline in the reaction of reduced enzyme with molecular oxygen below pH 7.19 It was initially concluded that there was a considerable disparity in the contributions of the two iron-sulfur centers to the spectral change observed on reduction of xanthine oxidase,23 but more recent work has demonstrated that in fact the two centers exhibit comparable extinction changes upon reduction.26 In addition to the absorption changes attributable to the iron-sulfur centers, these sites are also responsible for the pronounced circular dichroism exhibited by xanthine oxidase (Figure IB). 8 As with the absorption change, the two ironsulfur centers contribute comparably to the circular dichroism change seen observed upon reduction.26 While the resonance Raman spectrum of xanthine oxidase indicates that the iron-sulfur centers are typical 2Fe/2S of the spinach ferredoxin variety,27 recent Mossbauer experiments indicate that one of the centers has an unusually large quadrupole splitting and may be a Rieske-type center containing a nitrogenous distal ligand rather than the expected cysteine thiolate26 There is at present no clearly defined absorption or absorption change attributable to the molybdenum center of the native enzyme in the visible region, although there is evidence from magnetic circular dichroism studies that under certain conditions the Mo(V)) valence state may have weak absorption.28 There are, however, two complexes of pteridine compounds with the enzyme molybdenum center which give long-wavelength absorbance in the 600 to 650 nm region attributable to charge-transfer bands. The first of these is with violapterin (the product of enzymatic action on lumazine) and reduced enzyme,29 and represents the Ered-P complex in the catalytic cycle with this substrate. The second chargetransfer complex is with oxidized enzyme and 2-amino-4-hydroxy-6-formylpteridine.30 This pteridine is a slow substrate for xanthine oxidase, being converted to the corresponding 6carboxylic acid,31 and the charge-transfer complex seen with the enzyme apparently represents a particularly stable EOX*S complex. The two charge-transfer complexes thus represent catalytic intermediates just inward from Eox and Ered in the reductive half-reaction of xanthine oxidase. In both cases the participation of the molybdenum center is unequivocally established by loss of the long-wavelength absorbance upon reaction of the enzyme with cyanide. The observation of a charge-transfer complex would ordinarily be taken to reflect direct coordination of the heterocycles to the molybdenum, but the observation that removal of the catalytically required sulfur results in loss of the charge-transfer band (as opposed to merely shifting its maximum to another wavelength) suggests that the pteridines may be interacting only indirectly with the molybdenum in a manner mediated by the essential sulfur. By far the most generally useful spectroscopic tool in studying xanthine oxidase to date has been electron paramagnetic resonance (EPR) spectroscopy. Each of the redox-active centers has an EPR-detectable oxidation state (FADH-, Mo(V) and the reduced forms of the two iron-sulfur centers), and EPR has been used extensively both as a probe of the level of reduction of the several centers and the chemical nature of the paramagnetic species. The signals of the flavin and molybdenum centers are typically observed in the temperature range 100 to 300 K, while those of the iron-sulfur centers, below 30 K. The signal of reduced Fe/S II cannot be observed above 30 K owing to extremely rapid spin relaxation.8 The most commonly encountered EPR signals seen with xanthine oxidase are shown in Figure 2. The EPR signal of the flavin semiquinone exhibits an isotropic EPR signal with a g value of 2.0006, near that of the free electron, and a 19.4 G linewidth indicative of the blue neutral form (Figure 2A).8 This result is consistent with the absorption properties of the semiquinone.25 The reduced iron-sulfur centers exhibit considerably broader EPR signals than either

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FIGURE 2. The electron paramagnetic signals observed with xanthine oxidase. (A) the signal of the neutral flavin semiquinone obtained at 150 K. (B) the signals of the two iron-sulfur centers (upper), and the simulated spectra of Fe/S I (middle) and Fe/S II (lower), at 22 K. (C) the more commonly encountered signals attributable to the molybdenum center, from top to bottom: the Very Rapid signal seen with the substrate 2-hydroxy-6methylpurine as reductant, the Rapid Type 1 signal seen with xanthine as reductant, the Slow signal of the inactive desulfo form of the enzyme with sodium dithionite as reductant, and the Desulfo Inhibited signal observed on incubation of desulfo enzyme with excess ethylene glycol; all spectra recorded at 150 K. The Very Rapid signal was obtained at pH 10.0, all others at pH 8.5. All signals are calibrated to a microwave frequency of 9.54 GHz. The iron-sulfur data are redrawn from Reference 26. The flavin signal was obtained by computer subtraction of a composite spectrum comprised of both flavin and Rapid molybdenum signals.

the flavin or molybdenum centers, with g values of 2.022, 1.932, and 1.894 for Fe/S I and 2.110, 1.991, and 1.902 for Fe/S II (Figure 2B).26 There are a variety of signals attributable to the molybdenum center of the enzyme, with the particular signal observed depending upon the experimental conditions (Figure 2C). The Very Rapid and Rapid signals are named on the basis of the relative rates at which they are transiently observed in the course of enzyme reduction by xanthine.32 They are readily distinguishable from one another on the basis of their anisotropy and presence of magnetically coupled protons. The Very Rapid signal is nearly axial with g 1 2 3 = 2.025, 1.955, and 1.949, respectively, with no magnetically coupled protons.2'32 The Rapid Type 1 signal has §1,2,3 = 1.989, 1.969, and 1.964, respectively, with one strongly (aave = 13 G) and one weakly (aave - 3 G) coupled proton, both of which are approximately isotropically coupled.

28

Chemistry and Biochemistry of Flavoenzymes

The Rapid Type 2 signal has g l i 2 > 3 = 1.991, 1.969, and 1.963, respectively with two strongly and (again approximately isotropically) coupled protons having aave = 15 and 10 G.32 The Very Rapid signal is thought to be an authentic catalytic intermediate,112 principally on the basis of its kinetics of formation and decay.22-33 The Rapid signals, on the other hand, can also be seen upon reduction of enzyme by reagents such as sodium dithionite in the absence of substrate,2 and in all likelihood simply represent the equilibrium population of Mo(V) in partially reduced enzyme either in the presence of low (Type 1) or high (Type 2) concentrations. While the Very Rapid signal decays at approximately the same rate as the Rapid (Type 1) signal appears in the reaction of xanthine oxidase with xanthine under anaerobic conditions,22 the species giving rise to the Very Rapid signal does not appear to simply break down to that giving rise to the observed Rapid signal. The two signals do not represent the unprotonated and protonated forms, respectively, of a single enzyme form existing in a prototropic equilibrium,34 and on the basis of the difference in anisotropy of their EPR signals in all likelihood differ significantly in molybdenum coordination geometry. The differences in the structures of the species giving rise to the Rapid Type 1 and Type 2 signals are unknown, although it has been suggested that they differ in the orientation of the bound xanthine at the molybdenum center.2 The desulfo form of the molybdenum center exhibits a distinctive Mo(V) EPR signal, termed Slow in keeping with the above kinetic nomenclature. This signal has nearly axial symmetry with g, 2 3 = 1.975, 1.970, and 1.957, respectively, and a single strongly and isotropically coupled proton (aave = 16 G).32 The Slow signal is particularly difficult to saturate with microwave power in the EPR experiment, and is occassionally seen at helium temperatures in monitoring the EPR of the iron-sulfur centers, particularly when dithionite rather than xanthine is used as reductant. A final molybdenum signal of note is that observed with desulfo enzyme after incubation with ethylene glycol, termed Desulfo Inhibited, having §1,2,3 = 1.980, 1.973, and 1.967, respectively, with no detectable proton coupling.35 Extensive use has been made of this signal given its unusual stability toward air-oxidation, which makes it possible to obtain this signal in the absence of other EPR signals in the enzyme. The EPR properties of the molybdenum center of xanthine oxidase have been extensively reviewed.2

B. THERMODYNAMICS AND KINETICS OF INTERNAL ELECTRON TRANSFER Since the reductive and oxidative half-reactions of xanthine oxidase take place at separate sites of the enzyme, electron transfer between them is an integral aspect of catalysis. In this regard, the enzyme may be thought of not as a complex enzyme but rather as a simple electron transport system in which the redox-active sites are held at a (more or less) fixed distance and orientation with respect to one another within a single polypeptide. The distances between redox-active centers in xanthine oxidase are not known, however, with a single exception. On the basis of the magnetic interaction between the molybdenum center and Fe/ S I observed in the EPR signals of the centers, a distance of 10 to 20 A has been estimated.36 The lower limit is arrived at by assuming an entirely dipolar (as opposed to exchangecoupled) interaction, and to the extent that an exchange-coupled pathway exists, the calculation underestimates the true intersite distance. That this is likely to be the case was underscored in the original work by noting the quite isotropic nature of the coupling, suggesting that an exchange mechanism did in fact contribute to the overall magnetic coupling.36 The Mo-Fe/S I distance based on the observed magnetic coupling between the centers has subsequently been refined to 11 ± 3 A.37 An attempt has also been made to extract intersite distance information from the power saturation behavior of each of the EPR signals as a function of the oxidation states of surrounding sites.38 While the distances are in agreement with the above results based on magnetic coupling (intersite distances fall in the

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29

10 to 20 A range and the Mo-Fe/S I distance in particular at 11 ± 3 A), the difficulties of extracting two to three distances to other paramagnetic centers from the power saturation properties of a single EPR signal justify considerable conservatism in the interpretation of the data. The caveats to be taken into consideration in the application of the technique, particularly to biological systems, have been discussed.39 In any case, it appears on the basis of the evidence presently available that the redox-active centers of xanthine oxidase are as nearly far apart as possible given the likely physical dimensions of the enzyme molecule. A major advance in our understanding of the oxidation-reduction behavior of xanthine oxidase took place with the suggestion by Olson et al.23 that a rapid multiple oxidationreduction equilibrium existed among the centers of the enzyme that was governed exclusively by the relative reduction potentials of the sites involved. According to the rapid equilibrium model developed on the basis of this hypothesis, the sequence in which the several centers became reduced in the course of reductive titrations of enzyme with sodium dithionite was determined by the relative affinities of the sites for reducing equivalents. Thus, at any given level of enzyme reduction, those sites having higher reduction potentials would be expected to be predominantly reduced at the expense of those sites having lower potentials. Given a set of relative reduction potentials for the several centers, an equation was derived (Equation 1) that described the level of reduction of each center in the

enzyme as a function of the overall level of enzyme reduction.23 In this equation, K} to K5 are the equilibrium constants for the partition of a reducing equivalent between Fe/S II (chosen arbitrarily) and Fe/S I, FAD/FADH-, FADH-/FADH2, Mo(VI/V) and Mo(V/IV), respectively; Y is defined as the fraction of reduced Fe/S II divided by the fraction oxidized. A single set of relative reduction potentials related to K! to K5 by the Nernst equation was found that gave reduction profiles for each of the centers that closely matched the observed profiles obtained in reductive titrations. The relative potentials of the centers thus obtained have since been confirmed by direct potentiometric determination, following Mo(V) and flavin semiquinone by liquid-nitrogen temperature, EPR, iron-sulfur reduction by liquidhelium EPR, and flavin hydroquinone by the absorbance change at 450 nm.40 It has subsequently been shown, however, that the relative potentials of the centers in xanthine oxidase vary with temperature.41 The fact that EPR was used at different temperatures to monitor iron-sulfur reduction, FADH- and Mo(V) precludes a quantitative interpretation of the data, since the distribution of reducing equivalents could well be different at each temperature. In other work in which a series of flavin analogues of varying oxidation-reduction potential were substituted for the native cofactor in xanthine oxidase, however, it was found that the reduction profiles at room temperature for each of the centers were altered in a manner quantitatively consistent with a rapid equilibrium model.42 While the results of this study were originally treated by comparing data for the several centers at different temperatures, a comparison of the flavin hydroquinone reduction profile obtained at room-temperature for each of the enzyme forms unequivocally demonstrate the trend in reduction profiles predicted by the rapid equilibrium model (Figure 3). Thus, in enzyme containing the high-potential 8-chloro FAD, the flavin center became reduced much earlier in the course of room-temperature reductive titrations compared with the flavin of native enzyme, whereas in enzyme containing the low-potential 6-hydroxy FAD, the flavin site became reduced much later, again relative to the FAD of native enzyme. In each case, the reduction profiles of the other,

30

Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. Reduction profiles of the flavin centers of xanthine oxidase containing 8(C1) (circles), 8(SH) (squares) and 6(OH) (triangles) FAD. The profiles were obtained from room-temperature titrations of enzyme with sodium dithionite at pH 8.5. Redrawn from data in Reference 42.

unperturbed centers of the enzyme gave the reduction profiles predicted by the model, although these were necessarily monitored at different temperatures. The reductive halfreaction kinetics of enzyme containing the same flavin analogues also indicated that the oxidation-reduction equilibrium within the enzyme was rapidly achieved.42"44 A comparison of the rate of flavin vs. iron-sulfur reduction (monitored at different wavelengths) indicated that at times as short as 50 ms a disparity in the distribution of reducing equivalents between the flavin and iron-sulfur centers occurred, and always favored the site having the higher reduction potential.43*44 Thus, with high-potential flavins in the enzyme the flavin site itself was reduced more rapidly than the iron-sulfur centers (i.e., the flavin was preferentially reduced in the partially reduced enzyme intermediates generated in the course of the reaction). The reverse was true with low-potential flavins. These observations could be explained most conveniently if the rates of equilibration of reducing equivalents between the molybdenum center and the flavin on the one hand, and the molybdenum and iron-sulfur centers on the other were at least comparable to the rate at which reducing equivalents were introduced into the enzyme at the molybdenum center by xanthine. In addition to the accurate prediction of the reduction profiles observed in the titrations with xanthine oxidase containing the flavin analogues, the rapid equilibrium model also provides an explanation for the observed dependence of enzyme activity on the flavin midpoint potential in a series of flavin analogues upon incorporation into enzyme.44 Vmax exhibits a sigmoidal dependence on the midpoint potential of the flavin, with activity relative to native enzyme increasing from less than 10% for enzyme containing low-potential flavins to 100% for enzyme containing high-potential flavins. Studies of the oxidative half-reactions of the enzyme forms containing low-potential flavins demonstrate that without exception low activity is not due to an unreactive flavin in the oxidative half-reaction (in fact, the oxidative half-reaction kinetics are consistently faster than with the native cofactor44). The observed dependence can be accounted for in the context of a rapid equilibrium model from a consideration of the difference between the midpoint potentials of the flavin and molybdenum centers. With enzyme containing a high-potential flavin (8-chIoro or 2-thio), the distribution of reducing equivalents within the partially reduced enzyme encountered during turnover is expected to be such that the flavin is largely reduced and the molybdenum oxidized. Under these circumstances both centers are in the appropriate oxidation state to

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participate in their respective half-reaction of the catalytic cycle and turnover is unimpeded. As the midpoint potential of the flavin is lowered, however, reduced molybdenum and oxidized flavin will tend to predominate. For an enzyme containing flavin analogues of extremely low midpoint potential (8-hydroxy, 6-hydroxy, and 1-deaza), the molybdenum center is predominantly reduced at the expense of the flavin in partially reduced enzyme, and neither center is in the appropriate oxidation state for catalysis to take place. To the extent that this distribution of reducing equivalents occurs in the steady state, the enzyme is expected to be inactive. This can be quantified using the Nernst equation and the difference in midpoint potentials for the two centers, and the predicted response of enzyme activity to flavin midpoint potential accurately describes the data. In order to simulate the data well, however, it is necessary to use a molybdenum midpoint potential that is approximately 45 mV higher relative to the flavin than is observed with the free enzyme. This apparently reflects the fact that substrate binding raises the potential of the molybdenum center by this amount, a conclusion that is supported by both potentiometric45 and ligand-binding studies of xanthine oxidase.46 The process by which the oxidation-reduction properties of the centers in complex enzymes such as xanthine oxidase can influence the kinetic properties of these enzymes has been referred to as thermodynamic control.20 The rapid equilibrium model has come to be generally accepted in describing the equilibrium oxidation-reduction properties of xanthine oxidase, and much attention has focused on the determination of the reduction potentials of the redox-active centers and the factor which influence them, particularly pH. The initial evidence that the relative reduction potentials of xanthine oxidase were pH-dependent came from titrations of enzyme with sodium dithionite at several pH values, monitoring enzyme reduction spectrophotometrically.42 The relative reduction of flavin vs. iron-sulfur centers was monitored by comparing the fractional absorbance changes at 450 and 550 nm observed in the course of reductive titrations. Plots of the fractional absorbance change at 550 nm (where the spectral change is due entirely to the reduction of the two iron-sulfur centers) vs. that at 450 nm (where flavin reduction contributes appreciably to the observed spectral change) were simulated using the equations derived on the basis of a rapid equilibrium model, and used to estimate the difference between the flavin midpoint potential and the average of the iron-sulfur potentials (Figure 4). Such simulations indicated that on lowering the pH from 10 to 7 the flavin midpoint potential increased by approximately 70 mV with respect to the average of the iron-sulfur potentials.42 Such plots provide a convenient, if qualitative, method whereby changes in the relative values for the reduction potentials of the redox-active sites of multicenter enzymes can be estimated. The principal shortcoming of the procedure as applied to complex flavoproteins is the need to assume that the flavin semiquinone does not accumulate appreciably in the course of the reductive titration so as not to interfere with the monitoring of the ironsulfur reduction at 550 nm (an assumption that breaks down below pH 7 in the case of xanthine oxidase). Subsequent potentiometric work has established the pH dependence of the reduction potentials of xanthine oxidase on a more quantitative basis.45-47 In the work of Porras and Palmer,47 all centers were monitored at room temperature, the flavin semiquinone and Mo(V) by EPR, and the iron-sulfur centers by circular dichroism. A set of oxidation-reduction mediators were utilized to ensure adequate communication between the electrodes of the potentiometric apparatus and the redox-active centers in the enzyme.47 At pH 8.3 (i.e., at the pH optimum for activity), the reduction potentials for Fe/S I, Fe/S II, FAD/FADH-, FADH-/FADH2, Mo(VI/V), and Mo(V/IV) were -332, -224, -319, -238, -360, and — 320 mV vs. NHE, respectively. In the case of the flavin center, both the maximum amount of semiquinone generated and the poised potential at which it was observed decreased as the pH increased. A plot of the quinone/semiquinone (EJ and semiquinone/hydroquinone (E2) half-potentials indicated that the predominant pH effect was a decrease in Ej of

32

Chemistry and Biochemistry of Flavoenzymes DO

FIGURE 4. Plots of the percentage absorbance change at 550 nm vs. that at 450 nm for xanthine oxidase at pH 10 (solid circles), pH 8.5 (solid triangles), and pH 7.0 (solid squares), and aldehyde oxidase at pH 7.8 (open circles). The xanthine oxidase data are redrawn from Reference 42, the aldehyde oxidase data from Reference 166.

approximately 170 mV over the pH range 7 to 10. The result was consistent with the obligatory uptake of a proton to form the neutral semiquinone on one-electron reduction of the flavin, with the associated pK for the semiquinone being greater than 9. E2, on the other hand decreased by only 60 mV over the same pH range of 7 to 10. The pH dependence for the two flavin half-potentials was consistent with prototropic equilibria associated with both semiquinone and hydroquinone having pKs of 9.5 and 7.4, respectively. The data could be accounted for by Equation 2:

Throughout the pH range examined, E2 was greater than E 15 as manifested in the low integrated spin concentrations (5 to 10% of the enzyme concentration) of the observed flavin semiquinone EPR signal. Each of the iron-sulfur centers exhibited a pH dependence consistent with a shift in the pK of an associated ionizable group in each center from 6.4 in the oxidized form to 8.0 in the reduced. Throughout the pH range 7 to 10 one of the iron-sulfur centers (presumably Fe/S II, although it was impossible to demonstrate this conclusively from the room temperature circular dichroism data of the study) exhibited a reduction potential approximately 100 mV more positive than the other, a difference considerably greater than observed in the low temperature work.40-45 The two half-potentials of the molybdenum center also exhibited a significant pH dependence. The Mo(VI/V) potential decreased in a linear fashion by 190 mV over the pH range 7 to 10, whereas the Mo(V/IV) couple decreased only 80 mV over the same pH range, owing to coupling of electron uptake to a prototropic equilibrium associated with the Mo(V) state having a pK of approximately 8.4. This being the case, it was concluded that there must be an ionization associated with the Mo(VI) state having a comparable pK in order to account for the approximate linearity of the slope in the potential of the Mo(VI/V) couple as a function of pH. The pH profiles for both Ej and E2 were

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adequately described by Equation 3:

Thus, as with the flavin, uptake of the first reducing equivalent by the molybdenum center is tightly coupled to proton uptake, although in the case of the molybdenum center protonation can precede reduction of the fully oxidized form. A comparison of the reduction potentials of xanthine oxidase obtained at room temperature47 with those obtained at low temperature45 indicate that there are substantial changes observed upon freezing the enzyme. Not only are the potentials for Fe/S II, FAD/FADH-, and both molybdenum potentials 30 to 80 mV more positive at room than low temperature (at pH 8.3), but the pH dependence of the observed potentials is also effected, particularly in the cases of the molybdenum center and Fe/S II. Some of the effects of temperature on the reduction potentials appear to be due to the buffer used rather than temperature per se, however, since it has been shown that the low- and room-temperature potentials obtained in Bicine and Ches buffers, for example, agree rather well (although disparities of 20 to 30 mV remained for the Mo(V/IV) and Fe/S II couples).48 Pyrophosphate, which has long been used in studies of xanthine oxidase, appears to be a particularly poor choice of buffer on the basis of the pronounced shifts in enzyme reduction potentials upon freezing and Bicine is recommended as a more suitable substitute. The kinetics as well as thermodynamics of electron transfer has also been the subject of extensive investigation with xanthine oxidase. The initial effort to determine the rates at which electrons moved from one redox-active site of xanthine oxidase to another was made using the flash photolysis technique.49 In the procedure as applied to xanthine oxidase, laser photoexcitation of 5-deazalumiflavin (dFl) gave the strongly oxidizing dFl*, which rapidly oxidized EDTA present in the solution to form the excited dFl*' radical and decayed back to the ground state to give finally the strongly reducing dFl*.50 After initial reduction of the enzyme by the dFl' thus formed, subsequent intramolecular electron transfer was monitored spectrophotometrically. A variety of enzyme forms containing one or another of the enzyme centers rendered redox-inert by chemical modification were examined, and an overall scheme for electron transfer within the enzyme obtained. In general, the rates obtained were surprisingly slow, however, and in particular a rate of 12 s'1 was assigned to the equilibration of a reducing equivalent between the iron-sulfur and flavin centers. Under the conditions of the flash photolysis experiment (20 mAf phosphate, pH 7.0), the apparent limiting rate of Fe/S oxidation in the flash phase of the oxidative half-reaction (as monitored at 550 nm in the reaction of reduced enzyme with molecular oxygen) is approximately 90 s" 1 . 19 It necessarily follows that 90 s ~ ! represents a minimum value for the rate constant associated with the transfer of reducing equivalents from the iron-sulfur centers to the flavin. The flash photolysis studies have recently been reexamined using higher concentrations of the 5deazalumiflavin photosensitizer and improved instrumentation.51 On the basis of this new data, the prominent kinetic process taking place at 12 s" 1 observed in the earlier work is attributed to an artifact, and a new value of 100 s" 1 assigned for the equilibration of a reducing equivalent between the flavin and Fe/S II at pH 7; at pH 8.5 the rate constant increases only marginally to 118 s"1.51 The 12 s"1 process appears to be unrelated to the dithionite-reducible disulfide bond that is known to exist in xanthine oxidase,52 as enzyme containing disulfide-blocked enzyme also exhibits the 12 s^ 1 process. The chemical nature of the 12 s~ l process is at present unknown. Tollin and co-workers168 have recently compared the rates of electron transfer within xanthine oxidase and xanthine dehydrogenase by the flash photolysis technique. In this study slow (kobs ~ 12 s"1) kinetic processes are still observed, complicating the interpretation of the experimental results. It is to be noted that

34

Chemistry and Biochemistry of Flavoenzymes TABLE 1 The Oxidation-Reduction Potentials of Xanthine Oxidase at Low and High pH53 pH6.1 E Fraction (mV vs. NHE) reduced

Fe/S II Fe/S I FAD/FADHFADH-/FADH2 Mo(VI/V) Mo(V/IV)

-150 -261 -203 -168 -239 -301

0.90 0.17 0.12 0.82 0.24 0.00

pH9.2

E (mV vs. NHE)

Fraction reduced

-255 -345 -365 -252 -430 -350

0.96 0.43 0.02 0.83 0.00 0.00

FIGURE 5. The dependence of the spectrum of partially reduced xanthine oxidase on pH. Upper solid curve, oxidized enzyme; dashed curved, enzyme reduced with approximately three electron-equivalents of sodium dithionite, pH 6.1; lower solid line, enzyme at the same level of reduction, pH 9.6; dot-dashed line, enzyme fully reduced with sodium dithionite. Inset, the absorbance change observed at 525 nm upon mixing partially reduced xanthine oxidase in dilute glycine buffer, pH 10, with anaerobic, concentrated MES buffer (final pH of 6.2) at 25°C. The associated rate constant for the kinetic process is 155 s"1. Redrawn from Reference 53.

the rate constants for intramolecular electron transfer obtained by the pulse radiolysis technique were criticized in this paper on the grounds that a stoichiometric excess of reducing equivalents was generated in the pulse experiments, resulting in an excessive reduction of enzyme. The criticism is unfounded. A second approach to determining the rates of internal electron transfer within xanthine oxidase has taken advantage of the known pH-dependence of the reduction potentials of xanthine oxidase.53 Table 1 shows the potentials for the several centers of the enzyme at pH 6.1 and 9.2, along with the expected distributions of reducing equivalents within enzyme containing three of the six equivalents required for complete reduction (calculated using the equations derived on the basis of a rapid equilibrium model23). That the distribution of reducing equivalents in the enzyme at high and low pH is in fact quite different is indicated by the absorption spectra of partially reduced enzyme at pH 6 and 10 (Figure 5). When

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partially reduced enzyme at pH 10 (in dilute buffer) is mixed with concentrated anaerobic buffer at pH 6, a single-exponential process exhibiting a rate constant of 155 s" 1 is observed (Figure 5, Inset).53 When the direction of the pH-jump is reversed the observed rate doubles to 330 s" 1 . The only factors determining the observed rate constant are found to be the final pH of the reaction mixture and the temperature.53 Throughout the pH range 5.5 to 10 the observed rate constants are approximately tenfold greater than kcat under the same conditions, consistent with the assumptions of the rapid equilibrium model. The temperature dependence of the oxidation-reduction equilibration indicates activation energies of 19.8 and 9.2 kcal/ mol at pH 6 and 8.5, respectively. These values are quite high for simple electron transfer reactions, a point which will be taken up below. In the pH-jump experiments a comparison of the extent of the kinetically observed reaction with the static difference spectrum obtained spectrophotometrically indicates that there is an appreciable spectral change taking place in the dead-time of the stopped-flow apparatus (approximately 2 ms). The wavelength dependence of the dead-time spectral change is characteristic of a change in the level of ironsulfur reduction, at least above 400 nm. On the basis of their wavelength dependences and the calculated changes in the distribution of reducing equivalents shown in Table 1, the kinetically observable spectral change has been assigned to equilibration between the flavin (the FAD/FADH* couple) and Fe/S I, and the dead-time spectral change ascribed to equilibration between the molybdenum center (the MoVI/Mov couple) and Fe/S I.53 The pH-jump technique suffers from two potentially serious limitations from the standpoint of general applicability. First, it is limited by the dead-time of the stopped-flow apparatus and is thus unable to observe rate constants larger than approximately 500 s ~ ] . Second, it is clear from Table 1 that many of the rates for individual equilibration processes within xanthine oxidase cannot be determined since some of the centers (Fe/S II, for example, and both FADH2 and MoIV) do not change in their net level of reduction with a change in pH. Finally, the possibility of pH-dependent conformational changes must be taken into account and, as in the case of the above studies,53 appropriate controls carried out. These considerations notwithstanding, the technique has demonstrated that at least some rates of electron transfer within xanthine oxidase are consistent with the assumptions of the rapid equilibrium model and the known oxidative and reductive half-reaction kinetics of the enzyme. Further, the pH-jump technique circumvents the need to use powerful reducing agents and sophisticated instrumentation to rapidly introduce reducing equivalents into the enzyme. Given the amount of energy placed into the system in the course of generating reducing equivalents either photolytically or radiolytically, this is a significant advantage. The effects of directly exciting the protein chromophores by the laser pulse used to generate the reducing equivalents in the flash photolysis experiment, for example, are largely unknown, but likely to be quite large (as is the case with the deazaflavin photosensitizer used in the flash photolysis work). An examination of the conditions under which the initial flash photolysis experiments were performed indicate that as much as half of the light absorbed by the sample was absorbed by the enzyme rather than the deazaflavin photosensitizer.49 Finally, the pH-jump technique has the advantage of directly probing the involvement of protons in oxidation-reduction equilibria. In the case of xanthine oxidase, the first reduced flavin product seen in the photolytic studies is the neutral semiquinone, and protonation thus not only occurs but occurs very rapidly. A recent examination of the solvent kinetic isotope effect of the pH jump experiment169 indicates that electron equilibration within xanthine oxidase exhibits a solvent kinetic isotope effect of 7 for the pH jump in either direction. A proton inventory experiment indicates that the effect is due to a single exchangeable proton, assigned to be the N5-H of the neutral flavin semiquinone. The results were interpreted to indicate that the N5-H bond is partially broken in the course of electron transfer from the semiquinone to the iron-sulfur acceptor in the reaction. In an effort to further examine the rates of internal electron transfer within xanthine

36

Chemistry and Biochemistry of Flavoenzymes

oxidase, a third technique, pulse radiolysis, has recently been employed to investigate the rates of electron transfer within xanthine oxidase.54 Like the flash photolysis method, pulse radiolysis relies upon very rapidly introducing reducing equivalents into the enzyme and monitoring any subsequent internal electron transfer spectrophotometrically. Pulse radiolysis has the advantage over flash photolysis in that the target enzyme is not itself excited by the process of generating the reducing equivalents. When appropriate attention is paid to the experimental conditions, it is possible to steer the radiolytic chemistry in such a way as to generate either strongly reducing or oxidizing species whose chemical reactivity is appropriate for the study of redox-active proteins, and avoid undesirable reactions such as the ionization of aromatic amino acid residues.55 Using CO2* as the proximal reductant of enzyme, it has been demonstrated that (in 20 mM pyrophosphate, pH 8.5) the FAD of xanthine oxidase is reduced at a rate near the diffusion-controlled limit, and a subsequent electron transfer from the FADH- thus formed to one of the iron-sulfur centers (presumably Fe/S II on the basis of its much higher reduction potential relative to Fe/S I at ambient temperature) at a rate of 287 s"1. The kinetics of this latter process are found to be independent of the concentration of xanthine oxidase concentration, as expected for an internal equilibration process. The observed rate is remarkably similar to that assigned to the 330 s"1 process assigned to the equilibration of Fe/S I and flavin obtained from the pH-jump experiments (particularly when allowance is made for the 6°C difference in temperature in the two experiments). It is not possible at present to say, however, whether this agreement in observed rate constant is simply a coincidence or whether it is a manifestation of the specific manner in which the sites interact with one another in the enzyme. If, for example, only Fe/S I interacted directly with the flavin site then Fe/S II would be observed to equilibrate with the flavin at the same rate as Fe/S I, since subsequent transfer of a reducing equivalent from Fe/S I to Fe/S II would not be expected to give any appreciable absorbance change26 regardless of the rate of electron transfer. In other radiolytic experiments using the strongly oxidizing N3-54 rather than the reducing CO^, it is found that reduced enzyme is reoxidized by one electron at the flavin center to form FADH% which then returns to FADH2 by electron transfer from one of the iron-sulfur centers (presumably Fe/S I on the basis of its lower potential). The observed rate constant for this equilibration between Fe/S I and FADHVFADH2 couple is found to be 170 s'1, The application of pulse radiolysis to the study of electron transfer within xanthine oxidase has recently been extended.170 Using the neutral radical of Af-methylnicotinamide to rapidly introduce reducing equivalents into xanthine oxidase at the molybdenum center, evidence was found for electron transfer from the molybdenum to one of the iron-sulfur centers occurring with a rate constant of 8,500 s"1 at pH 6.0, 20°C. Subsequent electron transfer from the iron-sulfur to the flavin took place at 125 s" 1 , in good agreement with previous pH jump experiments when the difference in temperature was taken into account.53 These results indicate that the iron-sulfur centers of xanthine oxidase do indeed mediate electron transfer from the molybdenum to the flavin of the enzyme, as has long been thought. Experiments were also performed using the same 5-deazaflavin radical (generated radiolytically) that was used in the flash photolysis experiments. The results gave a rate constant for electron transfer between the flavin and one of the iron-sulfur centers of 145 s" 1 at pH 6.0 and 218 s"1 at pH 8.5, in agreement with previous pulse radiolysis and pH jump studies. Significantly, no slow kinetic processes such as have been observed with the flash photolysis technique were observed in the pulse radiolysis experiments. In all the pulse radiolysis work it was found necessary to block the reducible disulfide bond to prevent its one-electron reduction.54 A similarly reactive disulfide is also found in azurin.171 Both reductive and oxidative radiolytic experiments support the conclusion of the pHjump studies and the more recent flash photolysis work that reducing equivalents equilibrate among the redox-active centers of xanthine oxidase rapidly with respect to the rate of overall

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SCHEME 2. Proposed structure for the adduct of the FAD of xanthine oxidase with l-methyl-2-(bromomethyI)-4,7-dimethoxybenzimidazole.

catalysis. From the kinetic data53-54 and the known reduction potentials of the centers involved,47 rates for the forward and reverse electron transfer steps between the Fe/S I and the flavin under a variety of conditions can be calculated. The results indicate that the ironsulfur to flavin transfer is approximately 160 s^ 1 , and is essentially independent of the thermodynamic driving force for electron transfer. All of the influence of the thermodynamics of the system appears to be on the rate of electron transfer from the flavin to Fe/S I (which increases from 5 s"1 at pH 7 to 165 s"1 at pH 8.5).53 Since the rate of equilibration slows down as the pH is lowered in the pH-jump experiments, a plausible explanation for this observation is that electron transfer out of the flavin center is at least partially rate-limited by the rate of deprotonation of the reduced flavin participating in the equilibrium (FADHin the pH-jump experiments, FADH2 in the case of the oxidative pulse radiolysis experiment). This hypothesis is consistent with the observed high activation energies for the pH-jump kinetics, and the observation that a Bronsted plot of the logarithm of the flavin-to-Fe/S rate constant vs. pH gives a slope of approximately 0.7.55a It would be quite significant, should it be demonstrated in other systems as well, that protonation-deprotonation steps can be partially rate-limiting. C. THE FLAVIN SITE AND THE OXIDATIVE HALF-REACTION From a strictly chemical point of view, the FADH2 of xanthine oxidase is known to react with iodoacetamide (but not iodoacetate) to form a 4a carboxymethyl covalent adduct.56 The reaction is a manifestation of the ability of the reduced cofactor to serve as an effective nucleophile, and it has recently been shown that reaction of reduced enzyme with 1-methyl2-(bromomethyl)-4,7-dimethoxybenzimidazole also results in a 4a adduct57 (see Scheme 2). These two reactions indicate that the 4a position of the isoalloxazine ring is readily accessible to solvent. This is also true of the 8 position, on the basis of the conversion of 8-chloro FAD to the 8-mercapto derivative by reaction with sulfide.42 As stated above, the reaction of reduced xanthine oxidase with molecular oxygen takes place at the FAD of the enzyme. The reaction is markedly biphasic above pH 7, and the two phases exhibit different dependences on oxygen concentration, the faster phase having hyperbolic [oxygen] dependence whereas the slow phase is linear.l9 The apparent limiting rate of the fast phase increases monotonically with pH, from 13 s"1 at pH 7 to 350 s" 1 at pH 10. The Kd for oxygen in the fast phase increases from 80 \iM to 1.4 mM over the same range in pH, however, with the effect that kfast/Kd remains approximately constant at 2.0 x 105 M"^" 1 as the pH increases.19 This is reflected in the fact that double reciprocal plots of l/k fast vs. 1/[O2] as a function of pH gives a pattern of parallel lines.19'58 The apparent pK associated with this inhibition of oxygen reduction by protons is approximately 8. It has been suggested that the pK is associated with the N,-O2 region of the isoalloxazine ring,58

38

Chemistry and Biochemistry of Flavoenzymes

FIGURE 6. The kinetics of the reaction of reduced xanthine oxidase with molecular oxygen at pH 8.5. Solid line, the absorbance change at 450 nm due to reoxidation of the enzyme. Dashed line, the time course of superoxide generation, obtained by subtracting the transients observed at 550 nm upon mixing reduced enzyme with an oxygenated cytochrome c solution in the absence and presence of superoxide dismutase.

since it is thought that a negative charge in this region of the ring structure facilitates the reaction of the hydroquinone with O2 to give peroxide.59 In contrast to the observed pH dependence of the fast phase of the oxidative half-reaction, the second-order rate constant for the slow phase decreases somewhat from 1.8 x 104 M~ l s~^ at pH 6 to 7.3 X 103 M~ ! s ~ l at pH 10. The differences in kinetic order and pH dependence of the fast and slow phases of the reaction of reduced enzyme with oxygen are clearly indicative of different chemical mechanisms operating in the two phases. On the basis of a variety of pieces of indirect evidence, the kinetics of the oxidative half-reaction were interpreted in the original study19 to reflect the multistep oxidation of fully reduced enzyme to the one-electron reduced level (fast phase) and the subsequent reaction of the one-electron reduced enzyme thus generated to give the fully oxidized enzyme (slow phase). This interpretation has been confirmed in two independent studies in which the kinetics and stoichiometry of the formation of products of the oxidative half-reaction (i.e., superoxide and peroxide) were directly monitored. In the work by Hille and Massey,60 reduction of cytochrome c by superoxide61 was used to follow the formation of superoxide in the course of the reaction with molecular oxygen. Superoxide production was found to be biphasic after a pronounced 100 ms lag period during which most of the fast phase of enzyme reoxidation occurred (Figure 6). On the basis of the amount of cytochrome c reduced in the course of the reoxidation reaction, it was concluded that two equivalents of superoxide were generated per enzyme molecule reoxidized, with one equivalent generated rapidly and the second slowly. The data for both enzyme reoxidation and superoxide generation were quantitatively interpreted in terms of the following scheme, in which the first four reducing equivalents removed from the enzyme by molecular oxygen result in the formation of two equivalents of peroxide, while the last two removed form two equivalents of superoxide:

SCHEME 3. The stoichiometry of the oxidative half-reaction of xanthine oxidase. The kinetics of enzyme reoxidation and superoxide generation in the reaction of reduced

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enzyme with O2 were successfully simulated using a single intrinsic rate constant of 35 s" 1 for steps one through three in Scheme 3,60 attenuated by the fractional amount of FADH2 present in the partially reduced form of the enzyme reacting with oxygen (calculated assuming a rapid equilibrium model). In agreement with theory,23 the observed limiting slope in semilogarithmic plots of the fast phase of the enzyme reduction data underestimated the intrinsic rate constant required in the simulation to accurately fit the data by a factor of two (reflecting the sequential nature of the reoxidation scheme). The reoxidation scheme shown above was confirmed by examining the reoxidation of two-electron reduced enzyme (generated by the reaction of enzyme with substoichiometric xanthine so that no individual enzyme molecule could react with more than a single molecule of xanthine) and alloxanthinecomplexed enzyme (an analogue of four-electron reduced enzyme in which binding of alloxanthine at the reduced molybdenum center effectively locks the molybdenum in the reduced Mo(IV) valence state). For each of these forms of xanthine oxidase, the kinetic parameters from Scheme 3 obtained for fully reduced enzyme yielded good fits to the kinetics of both enzyme reoxidation and superoxide generation. The clearest evidence for Scheme 3 was from the oxidative half-reaction kinetics of two-electron reduced enzyme, where both enzyme reoxidation and superoxide generation were found to be biphasic.60 Identical rates for both fast and slow phases were observed for both enzyme reoxidation and superoxide generation, and the stoichiometry of superoxide generation indicated one superoxide formed per enzyme molecule reoxidized in each kinetic phase. In the study of the oxidative half-reaction undertaken by Porras and Palmer,58 not only was superoxide production followed by its reduction of cytochrome r, but hydrogen peroxide production followed as well by virtue of its formation of a quasi-stable complex with cytochrome c peroxidase (termed Compound I by analogy with the catalytic cycle of horseradish peroxidase) that exhibits distinctive spectral properties.62 The superoxide yield in this study was found to vary with pH, increasing from approximately 0.7 O^ enzyme below pH 7 to 1.8 at pH 9. The results are in reasonable agreement with the first study, which was carried out exclusively at pH 8.5. Given that the superoxide yield in these experiments was independent of the cytochrome c concentration above 125 |xM, it is unlikely that the low superoxide yield at low pH was due to the acceleration of spontaneous superoxide dismutation.63 These results suggest that at low pH two-electron reduced enzyme is able to reduce O2 to peroxide in a two-electron process. The stoichiometry of the Compound I yield in the reaction (carried out in the absence of cytochrome c) was found to be essentially constant as a function of pH at 2.0 to 2.3 H2O2/enzyme. Since superoxide formed in the oxidative half-reaction would be expected to comproportionate to form peroxide and molecular oxygen under the experimental conditions, the theoretical yield for peroxide is expected to be three per enzyme molecule reoxidized. The experimentally determined value of 2.3 was judged to underestimate the true peroxide yield as it was shown in control experiments that cytochrome c peroxidase Compound I reacted directly with reduced xanthine oxidase to reoxidize the latter at a nonnegligible rate. This not only decreased the number of reducing equivalents departing xanthine oxidase to form peroxide but also lowered the amount of Compound I ultimately detected.58 The oxidative sequence shown in Scheme 3 accounts for the long-noted observation that in steady-state turnover experiments high concentrations of oxygen and low concentrations of xanthine favor the production of superoxide.64 It has been pointed out58 that high concentrations of oxidizing substrate and/or low concentrations of reducing substrate are expected to result in predominantly oxidized enzyme in the steady state, with a concomitantly greater likelihood of oxygen encountering the two- and one-electron reduced enzyme whose oxidation results in the formation of superoxide. The scheme also accounts at least qualitatively for the observed decrease in superoxide yield from 80% of the total flux of reducing equivalents passing through the enzyme to 20% on going from pH 9 to 6. Over this pH

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Chemistry and Biochemistry of Flavoenzymes

range the limiting rate for the fast phase of the oxidative half-reaction decreases by a factor of 20, while that for the reductive half-reaction decreases by only a factor of two.19 Thus at saturating levels of oxygen and xanthine, lowering the pH will result in higher levels of enzyme reduction in the steady state, decreasing the likelihood that oxygen will encounter the one- and two-electron reduced enzyme species. This effect is expected to predominate over the relative acceleration of the superoxide-generating slow phase relative to the fast phase as the pH is lowered. The paradoxial result is that the yield of superoxide is expected to decrease, as is observed, under those conditions favoring the formation of flavin semiquinone, the oxidation state of the isoalloxazine ring most commonly associated with oneelectron transfer. The sequence shown in Scheme 3 makes no statement as to the chemical basis for the switch from a peroxide- to a superoxide-generating mechanism in the oxidative half-reaction of xanthine oxidase. The crux of the problem is that the FADH2 of a six- or four-electronreduced enzyme reacts with molecular oxygen to give peroxide, yet the FADH2 of a twoelectron-reduced enzyme reacts quantitatively, or nearly so, to form superoxide at pH 8.5. It has been proposed19 that the generation of peroxide by xanthine oxidase proceeds by two sequential one-electron transfer reactions rather than via a single two-step reaction, with electron transfer from other sites in the enzyme rapidly regenerating FADH2 from the FADHformed after the transfer of the first reducing equivalent to oxygen: FADH2 + O2 ?± FADH2 • O2 FADH2 • O2 -> FADH"-O2- + H+ H + 4- e- + FADH— O2- -> FADH2~O2FADH2-O2- -> FADH--H2O2 FADH"~H2O2 -» FADH- + H2O2 SCHEME 4. Proposed sequence for the generation of hydrogen peroxide in the oxidative half-reaction of xanthine oxidase by successive one-electron steps.

According to Scheme 4, only the FADH-/FADH2 couple is operative in the oxidative halfreaction, even in the formation of H2O2. This scheme emphasizes one possibly significant difference between FADH2 in two-electron reduced xanthine oxidase and other more reduced forms of the enzyme: in the case of two-electron reduced enzyme there is no reservoir of reducing equivalents in other sites of the enzyme to regenerate FADH2 from FADH' (step 3 of Scheme 4). It is possible that in the absence of such a reservoir the transfer of the remaining electron in the flavin semiquinone to the enzyme-bound superoxide is sufficiently slow relative to superoxide dissociation that the latter process predominates. Recent preliminary results with xanthine oxidase containing 5-thia FAD provide indirect support for the proposed scheme.553 This analogue of the native cofactor65 is capable of participating only in oxidation-reduction chemistry that corresponds to that of the FADHV FADH2 couple of the native cofactor (E = +400 mV vs. NHE)66 (See Scheme 5.) Since

SCHEME 5.

The structures of oxidized and reduced 5-thiariboflavin.

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5-thia FAD does not exhibit any appreciable visible absorbance, incorporation into deflavo xanthine oxidase was determined empirically on the basis of the enhanced reactivity of the reduced enzyme toward molecular oxygen upon incorporation of the flavin. The reconstituted enzyme reacted readily with molecular oxygen, but no superoxide was detected when cytochrome c was added to the reaction mix. The implication is that the 5-thia enzyme is able to generate peroxide by sequential one-electron steps, similar to the chemistry shown in Scheme 4. Should future work confirm and extend these preliminary observations, it would constitute substantive evidence that the sequence shown in Scheme 4 operates in the oxidative half-reaction of the native enzyme. In addition to molecular oxygen, other oxidizing substrates for xanthine oxidase include 2,6-dichlorophenolindophenol, methylene blue, phenazine methosulfate, and ferricyanide. Unlike oxygen, however, these oxidants of enzyme react at the molybdenum center and/or the iron-sulfur centers rather than the flavin, since removal of the flavin has little effect on these reactions.24 A reaction that does take place at the flavin center is the reduction of enzyme by NADH. 24 Although quite slow at neutral pH, this reaction permits the reduction of desulfo xanthine oxidase with a reagent other than sodium dithionite. The obligatory twoelectron nature of enzyme reduction by NADH has been found to be convenient in generating two-electron reduced forms of desulfo xanthine oxidase to examine the rate at which this partially reduced enzyme comproportionates with oxidized enzyme to give two equivalents of one-electron reduced enzyme (t1/2 of approximately 1 h at pH 8.5, 25°C, comparable to the rate observed with native enzyme).23 D. THE MOLYBDENUM CENTER AND THE REDUCTIVE HALF-REACTION The hydroxylation of xanthine by xanthine oxidase occurs at the molybdenum center of xanthine oxidase, and after internal electron transfer to the iron-sulfur and flavin chromophores results in the bleaching shown in Figure 1. From stopped-flow experiments monitoring the visible spectral change associated with the anaerobic reduction of enzyme by xanthine at pH 8.5, double reciprocal plots of the observed rate constant vs. xanthine concentration give klim = 17.5 s" 1 and Kd = 13 jxA/.23 Similarly, the reaction has been monitored using the rapid-quench technique in conjunction with EPR to follow iron-sulfur, flavin, and molybdenum reduction.22-23 The rapid quench EPR results for iron-sulfur reduction are in good agreement with the stopped-flow data.22 As mentioned above, two Mo(V) EPR signals are typically observed transiently: a Very Rapid signal (observed maximally at 5 to 10 ms after mixing enzyme with 200 (jiM xanthine), and a Rapid signal (observed maximally at approximately 50 ms after mixing). The appearance and decay of both signals is slowed by a factor of 1.5 to 2.0 upon deuteration of xanthine at the 8 position.22 Because xanthine oxidase has the capacity to take up a total of six equivalents, the reductive half-reaction involves the sequential reaction of three xanthine molecules with each functional enzyme molecule, and this complicates the interpretation of the above experimental results. In the case of the EPR studies, the Rapid signal appears at approximately the same rate as the Very Rapid signal decays, but the Rapid signal apparently contains xanthine rather than uric acid bound at the molybdenum center.2 It thus appears that the species giving rise to the Rapid signal represents a Michaelis complex in which a second (or third) xanthine molecule is bound at the molybdenum center of partially reduced enzyme, and in effect precedes the species giving the Very Rapid signal in a given cycle with xanthine. The necessarily sequential nature of the reaction of xanthine oxidase with excess xanthine also manifests itself in stopped-flow experiments, where it has been shown that the observed rate constant for the reaction at any given xanthine concentration underestimates the intrinsic rate constant for the reductive half-reaction by a factor of approximately two.23 This has been clearly demonstrated by performing the reductive half-reaction experiment under conditions of excess enzyme rather than substrate so that no enzyme molecule reacts more than once with xanthine.

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Chemistry and Biochemistry of Flavoenzymes

Under these conditions, the intrinsic limiting rate constant for the reductive half-reaction has been shown to be approximately 25 s" 1 . 23 This kinetic effect of the sequential nature of the reaction under conditions of excess xanthine has also been shown to account for the observed faster apparent rate of reduction of deflavo xanthine oxidase relative to the native enzyme,24 since in the former case reaction with only two equivalents of xanthine occurs and the reaction is thus expected to go to completion more rapidly.23 Experiments with xanthine oxidase that has been reacted with fluorodinitrobenzene have shown that a lysine residue is present near the molybdenum center.67 The reaction results in the modification of two lysine residues and a reduction in the overall rates of both catalysis and the reductive half-reaction by a factor of six. The oxidative half-reaction is unaffected by the chemical modification, and the results have been taken to indicate that modification of the lysine derivatives partially occluded the xanthine binding site at the molybdenum center.67 Under anaerobic conditions, incubation of the dinitrophenyl-derivatized enzyme with xanthine results in the reduction of one of the nitro groups of each dinitrophenyllysine residue to the amine. The kinetics of this reaction are biphasic, with approximately 50% of the associated absorbance change taking place in each of the two phases. The faster of the two phases (t1/2 of approximately 15 min) is dependent on enzyme concentration, suggesting an intermolecular reduction process, whereas the slower phase (t1/2 of approximately 3 h) is independent, indicative of an intramolecular reduction process.67 In addition to the catalytically essential sulfur discussed above whose loss gives the inactive desulfo form of xanthine oxidase, the molybdenum center also requires an organic cofactor for activity. The initial evidence for the presence of this cofactor was genetic,68 it being observed that a variety of pleiotropic mutants in Aspergitlus nidulans failed to form functional forms of both xanthine oxidase and nitrate reductase (another molybdenumcontaining enzyme that catalyzes the dehydroxylation of nitrate to nitrite, a reaction xanthine oxidase itself is capable of when supplied with a source of external reducing equivalents69). Subsequent reconstitution experiments using extracts of a mutant strain of Neurospora crassa (nit-1) which was capable of making apo nitrate reductase but lacked the ability to synthesize the cofactor, demonstrated that the cofactor was common to all molybdenum enzymes other than nitrogenase.70-71 It has recently been shown that the reconstitution can be quantitatively achieved.72 Elucidation of the structure of the cofactor has been a protracted affair, owing in large part to the fact that the extracted cofactor is extremely air-sensitive and rapidly degrades into multiple inactive products.73 While it has been known for some time that the structure incorporates a pteridine ring, it is only very recently that evidence which may be regarded as chemically definitive has been reported.74-75 On the basis of mass spectral, proton NMR, and energy dispersive X-ray analysis, the structure shown in Scheme 6 has been proposed for the oxidized form of the cofactor obtained after covalent modification with iodoacetamide. Methylation of one thiol of the cofactor was regarded as inconclusive in the report by Kramer et al. ,74 its presence has subsequently been confirmed by Fish and Massey75 using a somewhat different extraction procedure. Further, on the basis of both proton NMR75 and ENDOR76 experiments, it has been concluded that the side chain of the cofactor is saturated rather than existing as an enedithiol as originally reported.74 In the ENDOR work, resonances attributed to H-OS-Mo protons were observed76 in the Mo(V) ENDOR of the desulfo enzyme when chemically modified with ethylene glycol. This form of the enzyme was chosen for the ENDOR study because of the ease with which the enzyme could be prepared with only the molybdenum center paramagnetic. These results support the conclusion that the cofactor side chain is a saturated vicinal dithiol (see Scheme 6). Spectroscopic evidence indicates that it is the tetrahydropterin form of the structure shown, which spontaneously oxidizes to the pterin level in the extraction procedure.75 Ultimately, proof of structure will rely upon chemical synthesis, a step which has already been taken for urothione,77 the

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SCHEME 6. Proposed structure of the iodoacetamide derivative of the pterin cofactor of the molybdenum center of xanthine oxidase and other molybdenum hydroxylases.

metabolic end-product of the pterin ring under conditions of aberrant pterin cofactor metabolism.78 On the basis of the known presence of thiols (and possibly thioether) in the molybdenum coordination sphere of the xanthine oxidase,79-80 the cofactor has been proposed to coordinate directly to the molybdenum in the active site.73 This is consistent with the ENDOR results cited above, and with the observation that the isolated cofactor forms a discrete complex with molybdenum (V) which is observable by EPR.81 Two EPR signals of the molybdenum/ cofactor complex, designated 1 and 2, have been observed under conditions of controlled oxidation in dimethyl sulfoxide solution. Signal 1 is axial with gparallel = 2.0292 and g^. pendicuiar = 1-9776, while Signal 2 is rhombic with g t = 1.9776, g2 - 1.9590, and g3 = 1.9407. Signal 1 is observed initially when the heat-extracted cofactor is poised at approximately - 50 mV (vs. NHE), and decays to Signal 2 over a period of an hour. In the presence of thiophenol, the g values of Signal 1 are shifted somewhat (to gparanei = 2.026 and gperpendicuiar = 1.980), and the signal becomes stable indefinitely.81 The thiophenol spectrum obtained with 95Mo-enriched material is considerably different from the signal of 95MoO(SC6H5)4) ~~, indicating that the thiophenol modification is not due to displacement of the pterin cofactor from the molybdenum coordination sphere.81 As the integrated spin concentration of Signal 1, either in the absence or presence of thiophenol, is comparable to the activity of the extracted cofactor in the apo nitrate reductase assay (approximately 80%), it has been concluded that this signal arises from functional cofactor. Cofactor-complexed molybdenum from sulfite oxidase gives a thiophenol Signal 1 extremely similar to that seen with xanthine oxidase-derived material, and on the basis of this information the structure shown in Scheme 7 has been proposed for the thiophenol complex.81 Since this complex is competent in the

SCHEME 7. Proposed structure for the extracted cofactor of xanthine oxidase complexed to molybdenum.

nitrate reductase reconstitution assays, it has been concluded that the thiophenol ligands are reversibly displaced by two protein thiols upon reconstitution of the apo nitrate reductase. The molybdenum center of nitrate reductase has a second oxo group which must also be incorporated into the reconstituted enzyme, (this must in all likelihood be cis to the existing oxo group82) and barring some extensive ligand rearrangement upon reconstitution, it is thus likely that one of the four equitorial thiolates shown in Scheme 7 is in fact trans to the oxo group. This second oxo group in the reconstituted molybdenum center of nitrate reductase could be derived either from water present in the reconstitution incubation or from the dimethyl sulfoxide used as solvent in the extraction of the cofactor, since this has been

44

Chemistry and Biochemistry of Flavoenzymes

shown to be an effective oxo donor in inorganic systems likely to prove relevant to the molybdenum hydroxylases (see below). The role of the cofactor in catalysis is at present the subject of some conjecture. On the basis of the known oxidation-reduction stoichiometry of the enzyme, it is extremely unlikely that the pterin ring undergoes a change in oxidation state on reduction of enzyme by either xanthine or sodium dithionite. It has recently been suggested that the cofactor acts as the proximal donor to substrate in the hydroxylation reaction (Scheme 8), accepting an oxo

SCHEME 8. droxylase.

Proposed reaction mechanism for the biopterin-utilizing enzyme phenylalanine hy-

oxygen from the molybdenum center to form a 4a hydroxy species.83 While there is precedent for a pterin 4a hydroxide in the catalytic mechanism of phenylalanine hydroxylase,84-85 for example, the route of formation for such a reactive chemical species most likely proceeds via the 4a peroxide (formed by reaction of the dihydropterin with molecular oxygen, possibly with the involvement of the non-heme iron of the enzyme in oxygen activation85). The fate of the oxygen of the pterin 4a-hydroxide is to become water, however,84 and it is the distal oxygen of the putative 4a peroxide that is incorporated into product (Scheme 8). Formation of a pterin 4a hydroxide via oxo transfer from a molybdenum center, even a reduced one, is unlikely to be thermodynamically favorable. The biosynthesis, structure, and possible catalytic role of the pterin cofactor have recently been reviewed by K. V. Rajagopalan.172 A particularly interesting new development with regard to the molybdenum center of xanthine oxidase has been the report of the presence not only of the phosphate group of the molybdenum cofactor but also of a phosphoserine residue which is detectable by 31P-NMR.86 The latter residue exhibits a resonance in the inactive desulfo form of the enzyme approximately 3 ppm downfield from an external phosphoric acid standard. This resonance disappears, presumably due to line broadening as the phosphate group becomes less conformationally constrained, in the active form of the enzyme. The -3 ppm resonance is also lost on treatment of xanthine oxidase with alloxanthine, an enzyme inhibitor known to bind to the molybdenum center. Both the - 3 ppm resonance of the phosphoserine residue and a + 1 ppm resonance attributed to the phosphate group of the pterin cofactor lose a substantial amount of intensity upon formation of a stable Mo(V)-containing species with ethylene glycol. These observations have been taken to indicate that both the cofactor phosphate and the phosphoserine residue are within 10 A of the paramagnetic molybdenum atom.86 The phosphoserine is sufficiently exposed to solvent that it is to be broadened by Mn(II) in the solvent, but not so exposed that it is susceptible to hydrolysis by any of several phosphatases tested. This result is somewhat at odds with the conclusion drawn from an ENDOR experiment using enzyme dissolved in D2O in which minimal effects on the ENDOR signal were interpreted to reflect only limited solvent accessibility to the molybdenum center.76 The phosphoserine has been suggested to be the undesignated nucleophile that has been postulated to abstract the C(8) proton from xanthine in the course of catalysis,86 but a specific catalytic role for the group remains to be established. Recent reports from two different laboratories using 31P NMR173 and both 31P NMR and ENDOR174 have cast considerable doubt on the existence of a phosphoserine residue in xanthine oxidase. In both studies, functional enzyme was found to contain only three phosphate groups: two attributable to FAD, and one to the pterin cofactor of the molybdenum center.

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From a structural standpoint, many of the basic elements of the molybdenum coordination sphere have been elucidated by X-ray absorption spectroscopy.79'80'87"89 This technique involves scanning a sample with X-ray light of sufficient energy to ionize bound electrons (typically in the Is shell of the molybdenum) into the continuum. Upon ionization, the electron emerges from the absorbing atom as a photoelectron wave that is backscattered by atoms in its immediate environment. This backscattering may interfere either constructively or destructively with the photoelectron wave, depending upon the frequency of the photoelectron wave (a function of its energy, i.e., the incident X-ray energy less the energy to excite the bound electron into the continuum) and the distance between the absorber and the scatterer. The interference either enhances or diminishes the probability of X-ray absorption, with the effect that as the energy of the incident X-ray light is scanned above the absorption edge of the absorber a pattern of damped sine waves is observed. This data, termed X-ray absorption fine structure (EXAFS), can be quantitatively analyzed to yield information regarding the distance between absorber and scatterer, the chemical identity of the scatterer and the number of scatterers of a given type at a given distance from the absorber. The fundamentals of X-ray absorption spectroscopy have been effectively treated,90'91 and application of the technique specifically to biological systems has been comprehensively discussed.92 There is now a consensus from the X-ray absorption data obtained to date with molybdenum hydroxylases that the oxidized enzyme has the structure shown in Scheme 9

SCHEME 9. Proposed structures for the oxidized (left) and reduced (right) molybdenum centers of xanthine oxidase. No particular coordination geometry is to be inferred from the structures as shown.

(left), with the coordination sphere being dominated by a sulfido group (Mo=S, 2.15 A) and a single oxo group (Mo=O, 1.68 A). At least two and perhaps three thiols (Mo-SR, 2.45 A) are also present. On the basis of the Mo=O distance, the formal bond order of the bond is estimated to be almost three.93 Consistent with the known chemistry of dioxomolybdenum complexes,82 reduction of the enzyme molybdenum to the (IV) valence state results in the protonation of the Mo=S group to form Mo-SH (Scheme 9, right).79-80 This is manifested in the X-ray absorption data as the loss of the short Mo=S bond and the appearance of an additional Mo-S in the molybdenum coordination sphere. Formation of Mo-SH in the reduced enzyme readily accounts for the known sensitivity of reduced enzyme to classical thiol reagents such as /7-mercuribenzoate94'95 (p-CMB) and arsenite.7 The pCMB96 and arsenite97'98 complexes of xanthine oxidase have also been examined by EPR, and it is found that both mercury and arsenic nuclei interact extremely strongly with the unpaired electron in the partially reduced complexes. The arsenite complex in particular is of more than passing interest from a mechanistic standpoint since the EPR work has shown that while enzyme is completely inhibited, the complexed molybdenum center retains the ability to bind substrate.97-98 The arsenite complex of xanthine oxidase has also been examined by X-ray absorption spectroscopy,80 where it has been possible by analyzing both As and Mo EXAFS to determine the geometry of the Mo-S-As triangle. It is found that the S-Mo-As angle in the inhibitory complex is approximately 50°, and the Mo-As distance approximately 3 A, too far for the arsenic to be directly coordinated to the molybdenum. The arsenic atom

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Chemistry and Biochemistry of Flavoenzymes

is ideally situated, however, to interact with the lone dxy electron of Mo(V) in the partially reduced complex, accounting for the strong magnetic interaction observed by EPR.97 98 Binding of arsenite to the molybdenum center has a profound effect on the reduction potential of the Mo(VI/V) couple, raising it by approximately 170 mV while leaving the Mo(V/IV) couple essentially unperturbed." Also, despite the absence of proton hyperfine in the EPR signal of the arsenite complex, the Mo(VI/V) couple exhibits a 60 mV/pH unit dependence above a pH of 7." This observation emphasizes that those protons involved in strong magnetic interactions with the molybdenum center need not be those responsible for the pH dependence of the molybdenum reduction potentials. Bleaching of the weak chromophore that is formed on binding arsenite to the oxidized enzyme is associated with reduction to the level of Mo(V).99 The affinity of arsenite for oxidized native and desulfo enzyme is quite similar (Kd = 8 (xM and 20 jJiM, respectively), but on reduction of the native enzyme the Kd drops below 0.1 |xM, a decrease which is not observed with the desulfo enzyme. These results are consistent with the arsenic X-ray absorption data suggesting coordination of the arsenic to a thiol (presumably a cysteine residue in the active site) in addition to the Mo-SH of reduced enzyme.80 The former interaction would reasonably occur in both native and desulfo enzyme, but reduction of native enzyme to give the Mo-SH would provide a bidentate mode of arsenite binding that would account for the much higher affinity of arsenite for the reduced, native molybdenum center. That arsenite displaces the strongly coupled proton from the Mo(V) EPR signal and is itself strongly coupled to the unpaired electron spin, strongly suggests that it is the Mo-SH proton that is strongly coupled in the Rapid Type 1 EPR signal of the molybdenum center. This conclusion has also been reached from a consideration of other EPR properties of the center.2 Since the strongly coupled proton has been shown100 to be derived from the C(8) position of xanthine (the position ultimately hydroxylated), a proton or hydride abstraction role for the Mo=S group in the catalytic cycle is plausible, and consistent with the X-ray absorption data indicating that it is this group rather than the Mo=O that becomes protonated upon reduction.79'80 Somewhat surprisingly, binding of arsenite to the molybdenum center of reduced enzyme greatly enhances the reactivity of that center with molecular oxygen.101 The effect is clearest in the enhancement of the rate of reoxidation of deflavo xanthine oxidase in the presence of arsenite. Whereas the reoxidation of reduced deflavo xanthine oxidase with 125 \LM oxygen is slow (kobs = 0.02 s"1) and monophasic, in the presence of arsenite it is fast and multiphasic (kfast = 110 s ~ l , kslow was approximately 4 s"1).101 A double reciprocal plot of the rate constant for the fast phase as a function of oxygen concentration gives a limiting rate constant of 170 s"1 and a KD of 70 fxAf, indicating that the oxygen reactivity of the arsenite-complexed molybdenum center is comparable to that of the flavin center in native enzyme. In no case is more than one Mo=O group found in any form of functional xanthine oxidase by X-ray absorption spectroscopy, and it is well within the resolution of the technique to distinguish between one and two such groups. In desulfo xanthine dehydrogenase, however, the sulfide group has been shown to be replaced by a second oxo group,79 and this is presumed to be the case also with xanthine oxidase. It is thus extremely likely that it is the Mo=S sulfur that is removed from the molybdenum coordination sphere by reaction with cyanide to give the inactive desulfo enzyme. When the cyanolysis reaction is carried out under anaerobic conditions, enzyme is known to become partially reduced,14 and prior reduction of the enzyme renders it unreactive toward cyanide. These observations can be rationalized if the inactivation reaction were to proceed via nucleophilic attack on the Mo=S sulfur (with the molybdenum atom acting as an electron sink). (See Scheme 10.) Although the substitution of a second oxo group for the sulfido group of the molybdenum coordination sphere lowers the molybdenum midpoint potential by approximately 45 mV at pH 8.347 (principally an effect on the Mo(V/IV) couple), the desulfo enzyme still has a potential

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SCHEME 10. A plausible scheme for the inactivation of oxidized xanthine oxidase by cyanide.

some 60 mV higher than the xanthine uric acid couple at the same pH (440 mV94). Thus the loss of activity on formation of the desulfo enzyme cannot be accounted for on the basis of thermodynamics alone. Any mechanism for the reductive half-reaction must adequately account for the loss of activity on formation of the desulfo enzyme. X-ray absorption spectroscopy has several serious limitations as a structural probe, and these are reflected in the work done to date with the molybdenum hydroxylases. In addition to the requirement of millimolar concentrations of sample, the data analysis yields in the end only a radial distribution function, giving the kind of atom and its distance from the molybdenum but saying nothing directly about the coordination geometry.92 Also, because the technique is so sensitive to the presence of sulfur (which is a particularly strong scatterer of the photoelectron wave) it is frequently impossible to unambiguously establish the presence of nonsulfur ligands in a coordination sphere dominated by sulfur. In some of the inorganic complexes that are plausible model compounds for the enzyme active sites, for example, it has not been possible to unambiguously ascribe features in the X-ray absorption fine structure to nitrogen ligands known to be present from the crystallographic data.87 Further, it is generally not possible to distinguish oxygen and nitrogen on the basis of EXAFS data alone. Thus, while X-ray absorption spectroscopy has made substantive contributions in identifying the oxo and sulfido ligands of the molybdenum coordination sphere of xanthine oxidase, several crucial questions remain unanswered. It is not at present possible, for example, to make a definitive statement regarding the molybdenum coordination geometry or whether substrate chelates directly to the molybdenum at the initiation of the catalytic cycle. At least with regard to coordination geometry, recent advances in the application of L-edge X-ray absorption spectroscopy hold out the promise of contributing substantially to our understanding of the molybdenum coordination sphere of xanthine oxidase and other enzymes.102 An initial report of L-edge X-ray absorption spectroscopy on molybdenum enzymes and model compounds has recently appeared.175 From a mechanistic standpoint regarding the reductive half-reaction of xanthine oxidase, there is some consensus that the reaction is likely to pass through an intermediate having the nascent uric acid bound to molybdenum as Mo(IV)~O-R, which in the (V) valence state would give rise to the Very Rapid molybdenum EPR signal.2'103*104 Formation of such an intermediate must proceed from the Michaelis complex of oxidized enzyme and substrate via an undetermined number of chemical steps. Studies with both the substrate analogue 8bromoxanthine46 and the substrate lumazine105 indicate that formation of the Michaelis complex itself is a two-step equilibrium process. Displacement of the -OR group in the putative Mo(IV)-urate complex by HO-, followed by deprotonation of the Mo-OH thus formed to regenerate Mo=O would provide a plausible way to complete the reductive half-reaction. The nascent uric acid has been proposed to form either by hydride2-22 or proton104 abstraction from C(8) of xanthine by either the oxo or sulfide group. A Mo-OR structure is consistent with a considerable amount of data pertinent to the reaction mechanism (below), but leaves unexplained, for example, why no proton coupling is observed in the Very Rapid signal, yet the C(8) proton of substrate is observed in the subsequently appearing Rapid Type 1 signal which appears subsequently to the Very Rapid signal in rapid-quench experiments. Skibo and coworkers106 have undertaken an investigation of the oxidation of a series of substituted quinazolines by xanthine oxidase in an effort to further elucidate the mechanism

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Chemistry and Biochemistry of Flavoenzymes

of the reductive half-reaction. A Hammett plot of Iog(kcat/KM) vs. the pK of the quinazoline N(3) proton shows kcat/KM to be essentially independent of pK in the range 8 to 9.3, but strongly dependent on pK at higher values (the slope of the plot above pK 9.3 is —2.9). The occurrence of a breakpoint in the plot has been taken to reflect a change in rate-limiting step for the enzyme-catalyzed reaction. In the low pK regime, there is no discernible isotope effect observed for kcat/KM upon deuteration of C(2) (the position that is hydroxylated). By contrast, a primary isotope effect of 5 is found in the high pK regime, an effect that is principally manifested as a decrease in k cat . Noting that on the basis of resonance considerations the N(3) pK should correlate with the susceptibility of C(2) to nucleophilic attack, the kinetic data has been interpreted to reflect rate-limiting hydride abstraction and concerted nucleophilic attack on the substrate C(2) position in the high pK regime. The rate-limiting step in the low pK regime is proposed to be product release, accounting for the independence on substrate pK and absence of a kinetic isotope effect. The data for the high pK regime are not consistent with a mechanism involving rate-limiting proton abstraction followed by nucleophilic attack of the resulting carbanion on ligands to the molybdenum atom as the slope of the Hammett plot is negative. The phosphoserine observed by 31P-NMR has been suggested as a candidate for the nucleophile that attacks substrate,86 although it is unclear why formation of the desulfo enzyme should result in loss of activity in this case. The more likely candidate at the present time would appear to be the Mo=S group that becomes protonated upon reduction of the molybdenum center. Arguments deriving from the quinazoline study have been put forward regarding the well-known inhibition of xanthine oxidase at high concentrations of xanthine (above 200 fiM).107 Initially proposed to be the result of nonproductive binding of xanthine in the active site of the enzyme,18 the mechanism of this noncompetitive inhibition has been of more than passing interest for some time. On the basis of the quinazoline inactivation of xanthine oxidase by covalent modification the FAD site,107 it has been proposed that the site of inhibition is at the flavin.106 That this cannot be the case is reflected in the observation that the reductive half-reaction kinetics of xanthine oxidase under anaerobic conditions also exhibit pronounced inhibition at high xanthine concentrations.46 An alternative mechanism for excess substrate inhibition has been proposed based on tighter binding of product uric acid to the reduced enzyme than to the oxidized form of the center,108 and is based on the known perturbation of the reduction potentials of the molybdenum center on binding uric acid.45 This concept has been extended to include substrate as well as product, given the observed increase in molybdenum reduction potential in the presence of the substrate analogue 8-bromoxanthine.46 Inhibition by 8-bromoxanthine is uncompetitive in steady-state experiments rather than competitive, despite the close structural analogy to substrate, and by virtue of its effect on the reduction potentials of the molybdenum centers is proposed to inhibit xanthine oxidase via the formation of a complex of inhibitor with the reduced molybdenum center, rather than the oxidized form that substrate must interact with for catalysis to take place. The attractiveness of the hypothesis that excess substrate inhibition is due to formation of a nonproductive Ered-S complex is that it does not invoke a second substrate binding site (for which there is no evidence, apart from the observation of excess substrate inhibition). It is consistent (indeed thermodynamically obligatory) for inhibitor or substrate to bind more tightly to reduced than to oxidized enzyme if it raises the midpoint potential of the molybdenum center, and also with the observed pattern of inhibition. From a kinetic standpoint, binding to the reduced rather than the oxidized molybdenum center has the same effect on double-reciprocal plots as would binding to a spacially separated site of oxidized enzyme, since the enzyme containing reduced molybdenum is a different kinetic form than the oxidized enzyme with which the substrate interacts productively. The evidence cited in support of xanthine inhibition taking place at the flavin site, namely that excess substrate inhibition is

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observed in both regions of the Hammett plot of kcat/KM vs. pK, and that the substitutent effects in the quinazoline series on kcat/KM and k; are different, 106 is not necessarily inconsistent with a mechanism for excess xanthine inhibition based on substrate binding to reduced enzyme. Along a different line of investigation, experiments utilizing l8O-labeling have been undertaken in an effort to determine the origin of the oxygen atom incorporated into the hydroxyl group of product.104 It has been known for some time that water is the ultimate source of the incorporated oxygen,109 but EPR studies of enzyme in 17O-water indicating that an oxygen site assigned to be Mo=O becomes labeled during turnover raised the question as to whether the Mo=O group is the proximal oxygen donor to substrate.10:M 10 When xanthine oxidase labeled with 18O at the catalytically labile site and dissolved in 16O-water was reacted with substoichiometric xanthine (so that no enzyme molecule had the opportunity to react more than once with substrate), 92% of the uric acid recovered from the reaction mix contained 18O at the 8 position as determined by mass spectral analysis of the product. When the converse experiment was done with 16O-labeled enzyme in 18O-water, the uric acid was found to contain predominantly 16O at the 8 position. Control experiments demonstrated that the label incorporated into uric acid did not exchange with solvent under the experimental conditions, and that the silylation reaction necessary to volatilize the uric acid for the mass spectral analysis did not displace label. The enzyme thus appears to act in such a way that oxygen cycles out of the molybdenum coordination sphere with each turnover to be subsequently replaced by oxygen from water. The conclusion is that the reductive half-reaction of xanthine oxidase proceeds via the general mechanism shown in Scheme 11.

SCHEME 11. thine oxidase.

The stoichiometry of the reductive half-reaction of xan-

The chemical sequence in Scheme 11 has been referred to for obvious reasons as an oxo transfer mechanism and is consistent with an intermediate of the form Mo(IV)-OR in the reductive half-reaction. Such a mechanism has been given considerable credibility with the recent elaboration of the chemistry of a molybdenum complex that is a likely model compound (LMoO2, Scheme 12) for the active sites of the molybdenum hydroxylases.111-112

SCHEME 12. The molybdenum coordination system developed by Holm and coworkers exhibiting oxo transfer catalytic properties.

The transfer of an oxo group out of a molybdenum coordination sphere has been known for some time,113 but the reactions have typically been complicated by the well-known tendency of dioxomolybdenum(VI) complexes to form JJL-OXO dimers upon reduction.82 The new complex is sterically blocked from dimerizing and has been shown to be capable of cleanly catalyzing the transfer of an oxygen atom from a suitable donor to an acceptor according

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Chemistry and Biochemistry of Flavoenzymes

to the chemistry shown in Scheme 12 with no complications due to dimerization. 111 Despite the known strength of the Mo=O bond (with an enthalpy of formation on the order of - 45 kcal/mol114), it is sufficiently labile in solution to permit the removal of the oxygen with its transient replacement by a solvent ligand in the model system. A variety of oxo donors and acceptors have been examined, and the suitability of a given pair for turnover has been adequately accounted for on thermodynamic grounds.114 The specific thermodynamic criterion is that AHdonor > AHLMo0 > AHacceptor where the enthalpy changes are for the generalized oxidation reaction X + V2 02 (g) -> XO

Suitable oxo donors include t-butylperoxide, nitrate, dimethyl sulfoxide, and methanol; acceptors include acetaldehyde, formate, sulfite, and triphenylphosphine.114 Several of these (notably nitrate, acetaldehyde, and sulfite) are known substrates for molybdenum hydroxylases, and the data provide an adequate explanation as to why sulfite is hydroxylated and nitrate dehydroxylated by the relevant molybdenum-containing enzyme. The chemistry shown in Scheme 12 differs from that of the reductive half-reaction of xanthine oxidase (Scheme 11) in that the oxo donor becomes reduced in the model compound chemistry. In the enzymatic reaction, on the other hand, the oxo donor is water and the oxo group is regenerated in such a way as to generate a pair of protons and two reducing equivalents (transferred from the molybdenum center to other sites in the enzyme) rather than the molecule of hydrogen which would be expected by strict analogy to Scheme 12 with H2O as the oxo donor. The result is that regeneration of the oxo group, per se, is not an oxidative event from the standpoint of the enzyme molybdenum center, as is necessarily the case in the chemistry of the inorganic complex. The oxidized LMoO2 complex is capable of being reduced by thiophenol, to give the disulfide, and in the presence of an oxo donor will turn over liberating stoichiometric water.114 This reaction is in essence the reverse of the reaction catalyzed by xanthine oxidase, and is analogous to the dehydroxylation of nitrate to nitrite by nitrate reductase utilizing external reducing equivalents provided by NADPH.115 The evidence indicating that the catalytically labile oxygen site is in fact the Mo=O group of oxidized enzyme comes from EPR studies using 17O-labeled enzyme.103-110 It was found in this study that exchange of 17O into the molybdenum coordination sphere was slow (1 to 2 h) for the enzyme alone, but quite rapid (seconds) during turnover. Enzyme in 17Owater was reduced with either xanthine to generate the Very Rapid EPR signal,103 with formamide to generate the Rapid (Type 1) signal110 or with xanthine followed by complexation with arsenite to give the distinctive EPR signal of that complex.98 In the last two cases, clear evidence was found for a single 17O superhyperfine interaction that was strongly and anisotropically coupled to the electron spin, and was interpreted in both cases as arising from a Mo=O group. In the Very Rapid signal the superhyperfine interaction remained strong but is nearly isotropic, consistent with the formation of a Mo-O-R group, with the (now oxidized) purine moiety covalently bound at the molybdenum center. Given the structure of the arsenite-inhibited molybdenum center proposed on the basis of the X-ray absorption data, the Mo=O group is the obvious candidate for the site labeled with 17O after turnover in 17O-water and inhibition with arsenite.98 Recent EPR work with a homologous series of 33 S- and 17O-labeled inorganic molybdenum(V) complexes indicate that at least in sixcoordinate complexes Mo-17OH is strongly coupled whereas Mo=O is only weakly coupled to the electron spin, consistent with the Mo-OR assignment.116-117 This study leaves unexplained, however, why the presumed Mo=O of the Rapid and arsenite signals of xanthine

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oxidase is strongly, if anisotropically, magnetically coupled, although this could presumably be due to differences in the coordination geometry of the molybdenum center of the enzyme and the inorganic complexes. X-ray absorption data for a variety of forms of xanthine oxidase thought to be relevant to the catalytic cycle indicate that the Mo=O function is the most persistent feature of the molybdenum coordination sphere.118 In addition to being present in both oxidized and reduced enzyme, it is present in the complex of oxidized enzyme with the slow substrate, 2-amino4-hydroxy-6-formylpteridine,30 and of reduced enzyme with violapterin, the product of enzymic action on the pterin lumazine.29'105 Furthermore, the inhibitory complex of reduced enzyme with alioxanthine has been found to contain a Mo=O group.1183 Alioxanthine binds extremely tightly to the reduced molybdenum center and on the basis of its EPR properties in the Mo(V) state is thought to be a close structural analogue of the catalytic intermediate giving rise to the Very Rapid EPR signal.119 It may be concluded from the EXAFS work on the alioxanthine complex that either the species giving rise to the Very Rapid signal also has a Mo=O, or that the alioxanthine complex is not after all the close structural analogue to the Very Rapid species it has been assumed to be.2 Given the extremely similar EPR g values in the Very Rapid and alioxanthine EPR signals, the former interpretation must be regarded as the more likely possibility. The mechanistic implication is that the oxo group of the molybdenum coordination sphere is regenerated in the reductive half-reaction prior to product dissociation from the enzyme active site. The balance of observations at the present time makes a fairly strong case for the catalytically labile oxygen site being the Mo=O group of the molybdenum center. These include: (1) the observation that 17O is rapidly incorporated into the molybdenum coordination sphere only under turnover conditions, and that coupling of 17O in the Rapid (Type 1) and arsenite complex signals, at least, are qualitatively consistent with the presence of Mo=O; (2) the nearly quantitative incorporation of 18O substrate when the catalytically labile (and anisotropically coupled in the EPR) site of the enzyme is labeled with 18O; and (3) the impressive chemical precedent for oxo transfer in at least certain molybdenum complexes. The most important work that remains is to establish the geometry of the molybdenum coordination sphere and role of the pterin cofactor, and to address the issue of whether substrate coordinates to the molybdenum atom in the course of catalysis. This work will involve both a detailed spectroscopic investigation (XAS and resonance Raman in particular) of the inorganic model complexes, and further characterization of the various forms of the enzyme as well. To the latter end, recent resonance Raman work with the complex of reduced xanthine oxidase with violapterin is particularly encouraging.120 This complex exhibits longwavelength absorption that has been attributed to a charge-transfer complex between the molybdenum center and the pterin.29 The molybdenum center is specifically involved (presumably as the charge-donor, given that it is reduced in the complex), since cyanolysis of the enzyme results in the loss of the long-wavelength absorbance. The long-wavelength absorbance of these complexes may prove to be particularly significant vis a vis the model compound work since the reduced LMo(IV)O(DMF) complex of the model system also exhibits long-wavelength absorbance.111 Application of resonance Raman spectroscopy to the molybdenum center of xanthine oxidase should prove extremely useful in addressing the outstanding questions enumerated above regarding the nature of the molybdenum center in xanthine oxidase. The resonance Raman spectrum of dimethylsulfoxide reductase from E. coli (an enzyme containing a molybdenum center as its sole redox-active site) has recently been reported by Spiro and coworkers.176 Evidence is found for dithiolene coordination to molybdenum, consistent with the pterin cofactor being directly coordinated to the metal.

III. XANTHINE DEHYDROGENASE Xanthine dehydrogenase is distinguished from xanthine oxidase on the basis of its

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Chemistry and Biochemistry of Flavoenzymes

specificity for oxidizing substrate, preferring NAD + to molecular oxygen, whereas xanthine oxidase is unable to utilize NAD + as oxidant. The two most common sources of the dehydrogenase are avian (either chicken121-122 or turkey123) and rat liver,124 although the enzyme from Veillonella alcalescens (also known as Micrococcus lactilyticus125) has also been described.126 The bacterial protein is unique among all the enzymes considered here in that it utilizes another protein, namely ferredoxin, as its oxidizing substrate.127 Purification of each of the three xanthine dehydrogenases relies principally upon calcium phosphate and cation exchange chromatography procedures. A. PHYSICOCHEMICAL PROPERTIES Xanthine dehydrogenase from all sources has the same subunit and cofactor constitution as does xanthine oxidase, being a homodimer of Mr 300,000 with each subunit containing one molybdenum center, one flavin adenine dinucleotide, and a pair of 2Fe/2S centers. By contrast with xanthine oxidase, however, the dehydrogenase (at least from chicken liver) possesses no dithionite-reducible disuifide bond, and reductive titrations of the enzyme with sodium dithionite go to completion with the expected six reducing equivalents.128 In one of the few chemical modification studies done with xanthine dehydrogenase, it has been shown that a tyrosine residue is essential for reduction of NAD + at the flavin site of the chicken liver protein.129 Using 5'-[p-(fluorosulfonyl)benzoyl]adenosine (5'-FSBA), a reactive nucleotide analogue that is an effective active site directed reagent, it was demonstrated that a single tyrosine residue of xanthine dehydrogenase was modified which resulted in the loss of NAD + -reducing activity, but not xanthine-oxidizing or oxygen-reducing activity.129 Inactivation was competitively inhibited by NAD + , and the kinetics of inactivation were consistent with complex formation between enzyme and inactivator prior to the inactivation reaction. Inactivation was reversible upon extended incubation with dithiothreitol, consistent with the conclusion that a tyrosine residue was modified. Inactivation of the enzyme with 5'-FSBA resulted in a spectral change similar to that observed on binding NAD + and also increased the flavin reduction potentials, although not to the same extent as did binding of NAD + . 130 Together, the above results strongly indicate the presence of a NAD + binding site at the flavin site of xanthine dehydrogenase which is blocked upon inactivation of enzyme with 5'-FSBA. Inactivation of chicken liver xanthine dehydrogenase with radiolabeled 5'-FSBA followed by proteolytic digestion with subtilisin and S. aureus protease V8 resulted in the isolation by HPLC chromatography of a labeled peptide having the following sequence: Lys-Phe-Phe-Thr-Gly-X-Arg-Lys-Thr-Ile-Val-Lys-Pro-Glu where X designates the covalently modified tyrosine residue.130 Interestingly, recent pulse radiolysis studies of milk xanthine oxidase indicate the presence of a tyrosine residue whose oxidative modification inactivates the enzyme.131 The UV/visible absorption and circular dichroism of the dehydrogenase are very similar to that of the oxidase, the only appreciable differences being in the 430 to 470 nm region of the absorption spectrum, where fine structure of the spectrum attributable to the flavin center is somewhat different in the two enzymes. The EPR properties of the V. alcalescens132 and turkey liver123 xanthine dehydrogenases have been examined and, particularly with regard

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to their molybdenum centers, are also found to be quite similar to xanthine oxidase. Both dehydrogenases exhibit Rapid and Slow Mo(V) signals (the former giving corresponding hyperfine interactions with protons to the comparable oxidase signal). Unlike xanthine oxidase, however, the dehydrogenases from both sources give Mo(V) EPR signals as isolated. These signals closely resemble the Desulfo Inhibited EPR signal of xanthine oxidase, and are associated with nonfunctional enzyme obtained from preparations utilizing exposure to acetone (either in forming dry powders from fresh liver or as fractionation agent). The xanthine dehydrogenase from turkey liver gives a Very Rapid signal in a rapid-quench EPR experiment with xanthine as reductant,123 but attempts to observe a Very Rapid EPR signal in the dehydrogenases from other sources have been unsuccessful. The turkey enzyme also has the more striking homology to xanthine oxidase in its Fe/S EPR signals, with both a relatively narrow, high-temperature signal (g lj2(3 = 2.017, 1.932, 1.906) and a considerably broader, low-temperature signal (g 1 2 3 = 2.08, 2.00, 1.95). In the case of the Veillonella protein, by contrast, only a single Fe/S EPR signal is observed down to 8 K.132 The linewidths of the flavin semiquinone EPR signals of both bacterial and turkey dehydrogenases clearly indicate the formation of the blue neutral semiquinone.132 The first difference in the physicochemical properties of xanthine dehydrogenase and xanthine oxidase that might relate to the difference in function was shown to be in the oxidation-reduction potentials of the flavin center. The principal potentiometric difference between the two proteins is a 130 mV lower potential for the FADH-/FADH2 couple of the turkey dehydrogenase relative to the milk oxidase (the values for the FAD/FADH' and FADHV FADH2 couples were -360 mV and -240 mV in the oxidase, and -360 mV and -370 mV in the dehydrogenase, respectively).133 These results were obtained using a multiple temperature experimental protocol, however, and suffer from the caveat referred to earlier with regard to the potentiometric work with xanthine oxidase, namely that perturbations of the reduction potentials may well occur on varying the temperature. More recent experiments with the dehydrogenase (D) and oxidase (O) forms of rat xanthine dehydrogenase have shown unambiguously, however, that at room temperature the D form of the enzyme has a lower midpoint potential than the O form of the protein, and that the flavin semiquinone is in fact stabilized to a considerably greater extent in the D form.83 Together, these observations indicate that the FADHVFADH2 couple is decreased substantially in the conversion from the O to the D form of the rat liver enzyme, whereas the FAD/FADH* couple is relatively unaffected. These conclusions are consistent with the potentiometric results obtained with the turkey liver enzyme. B. KINETIC PROPERTIES The rapid kinetic properties of xanthine dehydrogenase have been most extensively examined in the case of the chicken liver enzyme, and are found to be considerably more complex than is the case with xanthine oxidase.128 A significant difference between xanthine dehydrogenase and xanthine oxidase from the standpoint of the reductive half-reaction is the inability of xanthine to bring about complete reduction of the dehydrogenase. The anaerobic reaction of xanthine dehydrogenase with xanthine exhibits four kinetic phases, the slowest takes place at 0.006 s"1 and has been taken to reflect the oxidation of reduced functional enzyme by oxidized nonfunctional enzyme (followed by rapid reduction of the now partially reoxidized functional enzyme). The three catalytically relevant phases have been interpreted in terms of Scheme 13.

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Chemistry and Biochemistry of Flavoenzymes

SCHEME 13. Proposed scheme for the reductive half-reaction of chicken liver xanthine dehydrogenase using xanthine as reductant.

On the basis of its wavelength dependence and dependence on xanthine concentration, the fastest phase represents the rapid reduction of enzyme by a first xanthine molecule in a preequilibrium process (KD = 280 jjuM, k,im = 180 s" 1 ). The second phase, which is independent of xanthine concentration, and is attributed to the dissociation of product uric acid from this two-electron reduced enzyme, followed by the rapid redistribution of reducing equivalents to one of the iron-sulfur centers and FADH-. After the formation of the free two-electron reduced enzyme, a second xanthine molecule reacts (again via a preequilibrium) to give free four-electron reduced enzyme. Unlike xanthine oxidase four-electron reduced xanthine dehydrogenase does not react further with xanthine, at least not on a timescale relevant to catalysis. This is reflected not only in the incomplete bleaching of functional enzyme upon reduction with excess xanthine, but also in the ability of uric acid to oxidize dithionite-reduced dehydrogenase to an appreciable extent.128 This reaction is found to take place at sufficiently low uric acid concentrations that the amount of uric acid generated in the faster phases of the reductive half-reaction would prevent the full reduction of the enzyme, thus accounting for the observed incomplete reduction of enzyme in the reaction with xanthine. A situation similar to that described above for the reductive half-reaction of xanthine dehydrogenase also arises in its oxidative half-reaction, i.e., the reaction of reduced enzyme with NAD+.128 Excess NAD+ does not fully oxidize dithionite-reduced xanthine dehydrogenase but brings the enzyme to approximately the two-electron reduced level in an apparently monophasic kinetic process. The observed rate constant exhibits hyperbolic dependence on NAD + concentration, with klim = 27 s" 1 and KD = 80 (xM. Further, just as uric acid partially oxidizes reduced xanthine dehydrogenase, so NADH is able to partially reduce the enzyme, suggesting that the endpoint level of enzyme reduction is determined by a thermodynamic equilibrium. That this is the case is reflected in the significantly greater reoxidation of reduced enzyme by 3-acetylpyridine-NAD (which has a reduction potential approximately 60 mV more positive than NAD + )134-135 than by NAD + itself.128 Interestingly, the reaction of reduced enzyme with NAD+ but not the reaction of oxidized enzyme with

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xanthine exhibits excess substrate inhibition. 128 The overall results from the rapid kinetics work are fully consistent with the conclusion that uric acid release in the reductive halfreaction is rate-limiting during turnover, and that reducing equivalents equilibrate rapidly within the enzyme relative to the rates at which they are introduced into or removed from the enzyme during turnover. On the basis of this argument, a lower limit of 165 s"1 for rates of electron transfer within xanthine dehydrogenase has been estimated.128 The reaction of chicken liver xanthine dehydrogenase with molecular oxygen has also been examined.136 Under steady-state conditions at pH 7.8, kcat = 0.7 s"1 at 4°C, compared with 1.7 s~ [ for NAD + as oxidant under the same conditions. The timecourse of the oxidative half-reaction using O2 as oxidant is multiphasic with the fastest phase occurring at a bimolecular rate of 1.9 x 103 M^s" 1 , which compares with a rate of 1.0 x 104 M^s' 1 for the slower of the two phases in the oxidative half-reaction of xanthine oxidase at the same pH and 25°C. The oxidative half-reaction kinetics are sufficiently fast with molecular oxygen as oxidizing substrate that uric acid release in the reductive half-reaction remains at least partially rate-limiting during turnover.136 Experiments monitoring superoxide generation by means of cyrochrome c reduction61 indicate that on average three of the six reducing equivalents in reduced enzyme depart as superoxide.136 This is consistent with the observation that in steady-state experiments approximately 40% of the throughput of reducing equivalents through the enzyme form superoxide. Enzyme-monitored turnover experiments in the absence and presence of cytochrome c indicate a good correlation between the occurrence of FADH* in the steady state and the formation of superoxide, and it thus appears that superoxide is formed principally via reaction of molecular oxygen with the semiquinone and peroxide by reaction with the flavin hydroquinone (in a presumably two-electron process). This contrasts with the proposed mechanism for the oxidative half-reaction of xanthine oxidase.19 Such a difference would be quite significant in terms of how the flavin reacts with molecular oxygen, superoxide formation being dependent primarily on the level of oxidation of other centers in the enzyme in the case of the oxidase, and simply on the steady-state level of semiquinone in the case of the dehydrogenase. Such a difference in oxygen reactivity could possibly reflect differences in the flavin half-potentials, the midpoint potential being higher in the case of the oxidase, but the thermodynamic stability of the semiquinone greater in the case of the dehydrogenase. Interestingly, chemical modification of the tyrosine at the NAD + binding site has no effect on the reactivity of the flavin of the dehydrogenase toward molecular oxygen.129 Perhaps the principal difference between the (milk) oxidase and (chicken liver) dehydrogenase is that the latter but not the former possesses a binding site for NAD + . This site has been investigated by examining the effect of 3-aminopyridine adenine dinucleotide (A AD), a nonreducible NAD + analogue.128 The presence of A AD in reductive titrations of the chicken liver dehydrogenase with dithionite results in a substantial perturbation of the absorbance changes observed in the course of the titration. The results are consistent with a substantial increase in the FAD/FADH* half-potential to a level comparable to that of the Fe/S II couple in the presence of A AD, resulting in a doubling of the maximum amount of semiquinone formed in the course of the reductive titration.128 In light of the effect of AAD on the flavin reduction potentials, it is quite surprising that AAD reduces the superoxide yield in the reaction of reduced dehydrogenase with molecular oxygen sixfold while reducing the rate of the overall oxidation by only a factor of two. The result has been rationalized in terms of slow superoxide dissociation from the AAD-complexed enzyme to permit further reduction to hydrogen peroxide, but the observation is nonetheless counterintuitive given the proposed mechanism of superoxide generation by direct reaction with FADH- on the dehydrogenase. C. THE DEHYDROGENASE/OXIDASE INTERC ONVERSION Avian and rat xanthine dehydrogenases differ from one another in that the rat protein

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Chemistry and Biochemistry of Flavoenzymes

can be reversibly interconverted from a dehydrogenase form to an oxidase form by the oxidation or chemical modification of protein thiols. 137 - 138 By contrast, conversion of the avian dehydrogenase to the oxidase form is irreversible, although the dehydrogenase form is considerably more stable than in the case of the rat protein.121'122 With the rat enzyme, the conversion from the dehydrogenase to the oxidase form is associated with a complete loss of reactivity toward NAD + and a fourfold increase in reactivity toward molecular oxygen.139 A substantial decrease in the ability of the enzyme to be reduced by NADH is also observed. Oxidation of thiols in the dehydrogenase form of the rat liver enzyme apparently results either directly or indirectly in the loss of a NAD + binding site in the vicinity of the flavin. Other observations also suggest a substantial difference in the environment of the flavin center of the dehydrogenase and oxidase. The flavin thiol present in milk xanthine oxidase reconstituted with 6-SH FAD is reactive with the thiol reagent methyimethanethiosulfonate, whereas reconstituted 6-SH xanthine dehydrogenase from chicken liver is relatively unreactive toward the reagent, reflecting a more buried 6 position in the dehydrogenase than in the oxidase.140 The 8-SH derivatives of both enzymes react readily with this same reagent, reflecting accessibility to solvent.140 Chicken liver xanthine dehydrogenase containing flavin derivatives with ionizable functional groups at the 6 or 8 positions of the flavin ring have also provided some insight into the nature of this structural constraint on the part of the polypeptide. It has been shown that the flavin analogue exist in protonated and deprotonated forms having markedly different spectral properties, and that protein interactions with the bound analogue can markedly perturb the pK associated with the ionization.141 Such perturbations can be conveniently monitored spectrophotometrically, and reflect a tighter binding of one form of the cofactor over another to the protein under investigation. In the case of milk xanthine oxidase, there is little discrimination between the binding of the protonated and deprotonated forms of 8-hydroxy and 8-mercapto FAD, and only a modest preference for the deprotonated forms of 6-hydroxy and 6-mercapto FAD.140 In contrast, chicken liver xanthine dehydrogenase has an overwhelming preference for the protonated forms of all four flavin analogues.140 The effect reflects an increase in the pKs for the bound cof actors of at least four pH units*. On the basis of these observations, it has been proposed that a negatively charged amino acid residue is present in the flavin binding pocket of the dehydrogenase (but not the oxidase) that stabilizes the protonated forms of the flavin analogues, and since the 8 position of the flavin was shown to be relatively solvent accessible the charged group was proposed to be localized at the N(l) end of the isoalloxazine ring.140 It is at present unclear whether the presence of this negative charge is directly responsible for the increase in the thermodynamic stability of the semiquinorse in xanthine dehydrogenase relative to the oxidase, although it is conceivable that it could. In this context, M. elsdenii flavodoxin, which also stabilizes the flavin semiquinone, does not have nearly so pronounced an effect upon the pK of the ionizable flavin analogues upon binding them.141 Thus it appears that there is more than one mode of polypeptide interaction with the flavin that can give rise to stabilization of the flavin semiquinone. It is tempting to speculate that these might reflect the two alternate thermodynamic approaches which can be envisaged, i.e., increasing the reduction potential of the FAD/FADH* couple or decreasing the potential of the FADHV FADH2 couple. Other work has focused on the enzymatic reactivity of xanthine dehydrogenase containing *

Nishino and co-workers have recently investigated differences in the environment of the FAD site of rat liver xanthine dehydrogenase in the dehydrogenase and oxidase forms.177 The study shows that the dehydrogenase and oxidase forms of the enzyme are selectively regenerated from deflavoenzyme when the flavin reconstitution was carried out in the presence or absence of dithiothreitol, respectively. Further, as with the chicken liver enzyme, the dehydrogenase form of the rat liver enzyme is found to bind the protonated forms of 8-SH and 6-OH FAD, while the oxidase form preferred the anionic forms of these chemically modified flavins.

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chemically modified flavin analogues. It is found that all forms of the chicken liver enzyme examined behave in reductive titrations in a manner very similar to xanthine oxidase and consistent with a rapid equilibrium hypothesis.142 Thus with high-potential flavins present in the enzyme (2-thio, 4-thio, 8-C1) the flavin center is reduced earlier in the course of titrations of enzyme with sodium dithionite than the iron-sulfur centers, while with lowpotential flavins (5-deaza, 8-OH, 6-SH) iron-sulfur reduction precede that of the flavin. Interestingly, all forms of the dehydrogenase give high yields of superoxide in steady-state experiments. As with native enzyme, the yield was typically 40 to 50% at pH 7.8, but in the case of 4-thio xanthine dehydrogenase it approached 100%.142 With both high-and lowpotential flavins in the chicken liver enzyme, the xanthine/O2 activity increased relative to the xanthine/NAD + activity, from approximately 1:300 for the native enzyme to 1:3 for 2thio xanthine dehydrogenase and 1:90 for the 6-SH and 6-OH derivatives. On the basis of these observations, it is clear that the difference between oxidase and dehydrogenase involves more than just the oxidation-reduction thermodynamics (lower midpoint potential and stabilized semiquinone relative to the oxidase), since the flavin potentials of 2-thio xanthine dehydrogenase are comparable to those of native xanthine oxidase, yet the former is in fact a relatively poor oxidase. This conclusion is supported by the studies described above with 3-aminopyridine adenine dinucleotide indicating the presence of a nucleotide binding site in the dehydrogenases but not the oxidase.128 Chemically modified flavin analogues have also been used with xanthine dehydrogenase to address the dependence of the catalytic activity on the reduction potential of the flavin. It is found with both the chicken liver enzyme142 and rat liver enzyme in the D form143 that a plot of the relative xanthine/NAD + activity as a function of the flavin midpoint potential gives a bell-shaped curve centered on the potential of the native cofactor. This dependence has been attributed to the thermodynamic stringency of reduction of NAD + in the oxidative half-reaction: when the flavin potential exceeds that of NAD + , the oxidative half-reaction is expected to be adversely effected.135 This thermodynamic constraint is obviously less serious on the xanthine/O2 reaction; and both D and O forms of the rat dehydrogenase give approximately sigmoidal plots of xanthine/O2 activity as a function of flavin midpoint potential reminiscent of the results with milk xanthine oxidase (although it is to be noted that in the case of the O form of the rat dehydrogenase, the 2-thio derivative falls well off the curve, having approximately half the activity of the 8-C1 enzyme, another high-potential form).143 In the case of the chicken liver dehydrogenase, on the other hand, no systematic dependence of the xanthine/O2 on the flavin midpoint potential is observed142 and this raises the question as to whether the redox-active centers in this enzyme interact in a different manner with regard to electron transfer (perhaps subtly so) than is the case with the other dehydrogenases and xanthine oxidase. D. MOLECULAR GENETICS Xanthine dehydrogenase is the protein encoded by the rosy locus of Drosophila melanogaster, and the gene has proven extremely useful in studying both genetic fine structure and population genetics owing to the ease in determining phenotype.l44 The gene has recently been cloned and sequenced, and found to be a 3766 base pair sequence consisting of four exons and three introns.145 It encodes a protein of 1335 amino acid residues having a calculated molecular weight of 146,898 Da, consistent with direct determination of the protein molecular weight from Drosophila.146 Somewhat disappointingly, two protein sequence database searches turned up no significant hornologies with known proteins. There is a six residue identity, however, from residues 194 to 199 of the predicted amino acid sequence of the Drosophila protein, with the 5'-FSBA-labeled peptide described above from the chicken liver protein. The conserved sequence is Phe-Phe-Thr-Gly-Tyr-Arg, and presumably delineates a portion of the nucleotide-binding site of the Drosophila protein. 13° This

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homology notwithstanding, it is extremely surprising that no other significant homologies to the Drosophila xanthine dehydrogenase sequence are found, given the number of centers present in the dehydrogenase and the relatively large number of classes of proteins which might be related to one or another of them. In addition to the work with Drosophila, the genetic sequencing of rat liver xanthine dehydrogenase has also been initiated.147 The work to date has involved the construction of a bacteriophage lambda expression library, immunological screening of the library using rabbit antibody to the rat liver dehydrogenase, and determination of the size of xanthine dehydrogenase mRNA by Northern blot analysis using the cDNA insert from the bacteriophage clone.147 The mRNA is found to be quite large relative to the cDNA insert, 5.9 vs. 1 kbp, indicating that the original insert did not contain the entirety of the structural gene for xanthine dehydrogenase. In addition to a comparison between the rat and fly sequences when this work is completed, it will be interesting to establish whether the mammalian gene also exhibits the unusually high polymorphism found in Drosophila. Xanthine dehydrogenase from rat liver has recently been cloned and its sequence analyzed.178 The protein is found to have considerable sequence homology to the dehydrogenase from Drosophila119^ The 5'-FSBA-labeled peptide obtained from the NAD binding site of the chicken liver protein corresponds to residues 387 to 400 in the rat liver sequence and 390 and 403 in the Drosophila sequence. Hydropathy analysis of the rat liver protein suggests that the iron sulfur centers are most likely in the N-terminal 184 amino acid residues of the rat liver protein. Work related to the molecular genetics and domain structure of xanthine dehydrogenase and related enzymes has recently been reviewed.181

IV. ALDEHYDE OXIDASE Aldehyde oxidase has been known since 1954 to be a separate enzyme having overlapping substrate specificity with xanthine oxidase.148 The conventional sources of the enzyme are pig149 or rabbit150 liver, rabbit appearing to be the more convenient source owing to difficulties with contaminating catalase in the pig liver preparation.151 Like xanthine oxidase, aldehyde oxidase is a homodimer of Mr 300,000 and has the identical complement of redox-active centers: molybdenum, FAD, and a pair of 2Fe/2S centers. Also like xanthine oxidase, the physiological oxidant for aldehyde oxidase is molecular oxygen. The two enzymes are distinguishable in that aldehyde oxidase has negligible xanthine-oxidizing activity, but will convert Af-methylnicotinamide to the 6-pyridone152 and 2-amino-4-hydroxy-6-formylpteridine to the corresponding carboxylic acid,153 reactions which xanthine oxidase catalyzes only extremely slowly. Also, menadione is a potent inhibitor of aldehyde oxidase but not of xanthine oxidase.150 This inhibition apparently takes place at the flavin site of aldehyde oxidase, and suggests that there are subtle differences in the flavin binding site as well as the molybdenum centers of the two enzymes. Aldehyde oxidase also has the ability to reduce several different sulfoxides (notably the sulfa drug Sulindac, ds-5-fluoro-2-methyl-l-[p(methylsulfinyl) benzylidenyl]indene-3-acetic acid) to the corresponding sulfide.154 Assuming this reaction takes place at the molybdenum center of the enzyme, it is analogous to the reaction catalyzed by the molybdenum-containing biotin-S-oxide reductase from E. coli.155-156 The similarity of the two reactions in fact raises the possibility that the two proteins may in fact be one and the same, albeit from different sources. Among the straight-chain aliphatic aldehydes, kcat is optimal with butyraldehyde at 0.9 s^ 1 (based on a molecular weight for the enzyme of 300,000) and KM optimal with valeraldehyde at 1.25 mA/. 157 The combined effects give an optimal kcat/KM for valeraldehyde of 8.2 x 103 M-'s' 1 . The highest value for kcat/KM found among the aliphatic aldehydes investigated was 2 X 104 M ~ l s ~ l with 2-ethylbutyraldehyde. The relative substrate specificities of aldehyde oxidase and xanthine oxidase for a wide variety of aromatic heterocycles

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has also been examined.158 Among the compounds investigated, xanthine oxidase had the higher relative activity toward carbon-substituted compounds, whereas aldehyde oxidase had the higher for nitrogen-substituted compounds. From this work, the general trend in Vmax among the aromatic heterocycles was: pteridines > pyrimidines > purines, and in KM: pteridines < purines < pyrimidines, although there were rather wide ranges in both values based on the nature of the ring substitutions examined. Among the best substrates for aldehyde oxidase was pteridine itself, which was hydroxylated at a comparable rate to valeraldehyde and had a KM of only 40 jjiM.158 Other substrates exhibiting high turnovers with aldehyde oxidase were 2-hydroxypyrimidine, 3-methylhypoxanthine, and 6-cyanopurine. Of these, 3methylhypoxanthine exhibited both the highest turnover with aldehyde oxidase and the greatest discrimination between xanthine oxidase and aldehyde oxidase.158 It was noted on examination of the data compiled in this study that it is unlikely that the physiological role of aldehyde oxidase is the hydroxylation of aliphatic aldehydes, a NADH-dependent aldehyde dehydrogenase having a much higher affinity for acetaldehyde.159'160 It was concluded that the enzyme functioned instead as a substituted purine oxidase in vivo. A. PHYSICOCHEMICAL PROPERTIES The molybdenum center of aldehyde oxidase possesses the same pterin cofactor common to all other molybdenum hydroxylases, as evidenced by the ability of acid extracts of the enzyme to reconstitute nitrate reductase activity in extracts of the Neurospora crassa mutant incapable of making the cofactor.70 Like xanthine oxidase, aldehyde oxidase possesses a cyanolyzable sulfur atom at the molybdenum center that is required for activity, and some 50% of the enzyme as routinely isolated is to be found as the inactive desulfo form of the enzyme.161 By analogy to xanthine oxidase and xanthine dehydrogenase the cyanolyzable sulfur is expected to be a Mo=S group, although no X-ray absorption spectroscopy has been done with aldehyde oxidase to date. In contrast to the cyanolysis of xanthine oxidase, which requires an hour or longer to go to completion, the inactivation of aldehyde oxidase by CNis essentially instantaneous.157 Similarly, inhibition by both arsenite and p-chloromercuribenzoate exhibit much more rapid kinetics with aldehyde oxidase compared with xanthine oxidase. In the case of both cyanide and arsenite inhibition, the rate of inhibitor dissociation from the complexed enzyme is sufficiently great that inhibition is reversible and competitive in nature, 162 in contrast to the slow and uncompetitive inhibition observed with xanthine oxidase. It is a general characteristic of slow-binding enzyme inhibitors that they exhibit uncompetitive inhibition, regardless of the manner in which they interact with their target enzymes,163 however, and the difference in inhibition pattern observed with aldehyde oxidase and xanthine oxidase does not necessarily reflect fundamentally different modes of interaction of arsenite and cyanide with the two enzymes. In general, while it appears that aldehyde oxidase and xanthine oxidase do exhibit differences in substrate specificity and reactivity toward inhibitors, these are rather subtle and the molybdenum centers of the two enzyme are likely to be fundamentally very similar. The spectral properties of aldehyde oxidase are by and large quite similar to those of xanthine oxidase. The UV/visible absorption spectrum of aldehyde oxidase is indistinguishable from xanthine oxidase, in both the native and deflavo forms of the protein.164 By contrast, the EPR properties of aldehyde oxidase are quite distinct from xanthine oxidase. Perhaps the most singular difference is the presence of an air-stable Mo(V) EPR signal in aldehyde oxidase as isolated that is extremely similar to that observed with xanthine dehydrogenase.165 This signal exhibits no proton hyperfine, has very little g-anisotropy, and bears a striking resemblance to the air-stable EPR signal seen with turkey xanthine dehydrogenase and the signal observed upon reaction of desulfo xanthine oxidase with ethylene glycol (Figure 2C).164 As stated above in the case of xanthine dehydrogenase, this signal appears to be associated with nonfunctional enzyme from preparations that incorporate

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Chemistry and Biochemistry of Flavoenzymes

protein extraction from an acetone powder. The molybdenum center of aldehyde oxidase also exhibits a Rapid Mo(V) signal which can be observed either by partial reduction of enzyme with substoichiometric amounts of A^-methylnicotinamide165 or by rapid-quench of the reaction of enzyme with excess substrate.166 In the latter experiments, no signal corresponding to the Very Rapid signal seen with xanthine oxidase is observed with aldehyde oxidase.166 In both equilibrium and kinetic experiments the enzyme exhibits the characteristic EPR signal of the neutral flavin semiquinone.165 Like xanthine oxidase, the two iron-sulfur centers of aldehyde oxidase can be distinguished on the basis of their linewidths and temperature dependence.165 The broader signal corresponds rather well with that of Fe/S II in xanthine oxidase, with g-values for the former signal of 2.106, 2.003, and 1.915. The signal from aldehyde oxidase differs from that seen with xanthine oxidase in that extremely high microwave powers are required to observe the signal even at 20 K. The narrower Fe/S I signal (i.e., the one observable at liquid-nitrogen temperatures)164 is remarkable in exhibiting almost axial symmetry with g-values of 2.018, 1.930, and 1.918. As with xanthine oxidase and xanthine dehydrogenase, there is EPR evidence for magnetic coupling between the molybdenum center and Fe/S I (i.e., the center giving the narrower of the two iron-sulfur EPR signals) of aldehyde oxidase, indicative of a site-site distance of approximately 11 A. 167 Although other interactions are suggested on the basis of the power saturation behavior of the various signals, in general these interactions are much weaker in aldehyde oxidase than in xanthine oxidase.167 The oxidation-reduction potentials of aldehyde oxidase have been determined using a combination of room-temperature absorption and cryogenic EPR spectroscopy to follow the reduction of the several centers as a function of the poised potential of the system.16? While the study suffers from the criticism discussed above regarding the inadvisability of comparing data at ambient and cryogenic temperatures, some generalizations with regard to xanthine oxidase can be made. First, on the basis of plots of the fractional absorbance change at 450 nm vs. that at 550 nm it is clear that the flavin midpoint potential is of comparable magnitude to the average of the iron-sulfur potentials, as is the case in xanthine oxidase. The data for aldehyde oxidase fall well within the range observed for xanthine oxidase at several pH values (Figure 4).167 Given the dependence of the 450/550 plots on the relative magnitudes of these values, the extent to which the data for aldehyde oxidase fits in with the xanthine oxidase data reflects the underlying similarities between the two proteins. B. KINETIC PROPERTIES The steady-state kinetics for aldehyde oxidase with A^-methylnicotinamide (NMN) as substrate are 5.4 s"1 for kcat and 400 pjVf for the NMN K M , giving a value for kcat/KM of 1.4 x 104 M~ ! s~'. l64 Surprisingly, catalysis is independent of the concentration of oxidizing substrate using either molecular oxygen or potassium ferricyanide. In both cases a set of horizontal (within experimental error) lines are observed in double reciprocal plots of 1/V vs. l/[oxygen] or l/[ferricyanide].164 The conclusion from the data taken at face value is that the Km for both substrates is much smaller than the experimental range (125 fiM for oxygen and 40 \^M for ferricyanide). DCIP, on the other hand, gives a well-behaved pattern of lines in a double reciprocal plot which intersect on the x-axis and yield a KM for DCIP of 1 juiM.164 The reductive half-reaction kinetics of aldehyde oxidase using NMN as reductant are strikingly similar to what is observed with xanthine oxidase using xanthine as reductant. The reaction is markedly biphasic with the slow phase being attributed to the slow reduction of nonfunctional enzyme by reduced functional enzyme after reaction with substrate. The fast phase exhibits hyperbolic dependence on NMN concentration and gives a limiting apparent rate of reduction of 12.3 s" 1 and a KD of 200 (xM for NMN (compared with 17 s" 1 and 13 (xM for the xanthine/xanthine oxidase reaction23). Somewhat surprisingly, aldehyde oxidase does not exhibit excess substrate inhibition in the reductive half-reaction

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when yV-methylnicotinamide is used as reducing substrate. By contrast with xanthine oxidase, aldehyde oxidase reacts with molecular oxygen to give a single kinetic phase. The reaction is second-order with an associated rate constant of 5.3 x 105 M^s" 1 , fivefold faster than the slow phase of the oxidative half-reaction of xanthine oxidase. A substantial amount of superoxide is generated in the oxidative half-reaction, as reflected in the absolute requirement for oxygen in order to obtain reduction of cytochrome c, and the inhibition of this aerobic cytochrome reduction by superoxide dismutase.164-166 The stoichiometry of superoxide has not been quantitated to date, however. As in the case of xanthine oxidase, that the oxidative half-reaction of aldehyde oxidase takes place at the flavin site is indicated by the loss of reactivity toward molecular oxygen on removal of the enzyme FAD,164 but no extensive flavin substitution study has yet been undertaken.

V. CONCLUDING REMARKS Several key issues with regard to the structure and function of the molybdenum hydroxylases remain to be addressed in the future. With the structure of the pterin cofactor fairly well established, the mode of its interaction with the molybdenum atom in the enzyme active site must be elucidated, and any active role of the cofactor itself in catalysis established, particularly in regard to the phosphate group. Similarly, the catalytic role, if any, of the phosphoserine residue shown to be present at (or near) the active site must be addressed in future studies. These issues bring up the larger concern of establishing the chemistry that leads up to the putative Mo-OR intermediate of the catalytic cycle. To this end further work with homologous series of substrates along the lines of the quinazoline studies described here (and extended to include kinetic isotope experiments of the reductive half-reaction) hold out considerable promise. It would be of particular interest to determine whether any of the quinazoline species, for example, give Very Rapid-type EPR signals in the anaerobic reaction with enzyme, and the nature of differences in kinetic isotope effect on the reductive half-reaction with low-pK (limiting product release) and high-pK (limiting hydride abstraction). Such studies would undoubtedly address the issue of how the Mo=O group of oxidized enzyme might come to be covalently modified to form a Mo-OR intermediate, and the manner in which the Mo=O group might be regenerated from such an intermediate. The application of resonance Raman to studies of the molybdenum hydroxylases promises also to make a major contribution with regard to the reductive half-reaction of the molybdenum hydroxylases, particularly with regard to the manner in which substrate interacts with the molybdenum and the coordination geometry of the charge-transfer species amenable to the technique, A second major area for future work is a further examination of the rates of intramolecular electron transfer within these enzymes and the factors that influence it. Agreement between results obtained from the flash photolysis technique on the one hand and the pH-jump and pulse radiolysis techniques on the other are in fair agreement at neutral pH, but diverge at high pH where the enzyme is more active. Future experiments must address in particular the pH-dependence of the rates of internal equilibration of reducing equivalents. In this regard, studies with enzyme substituted with chemically modified flavin derivatives of varying oxidation-reduction potential should prove extremely useful not only with regard to the issue of the pH dependence of electron transfer but also the dependence of electron transfer on the thermodynamic driving force of the reaction. Given the differences in the oxidation-reduction potentials of the flavin in the dehydrogenase relative to the oxidase, studies with the former would also be extremely useful. Finally, with the amino acid sequences of molybdenum hydroxylases coming to light, there is hope that concrete structural information will be forthcoming in the near future. The identification of a homologous region in the amino acid sequences of chicken liver and

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Chemistry and Biochemistry of Flavoenzymes

Drosophila xanthine dehydrogenases points the way to future studies aimed at extending the homology between these two proteins. It would obviously be of interest to establish the molybdenum-binding domain that these proteins presumably have in common in such studies.

ACKNOWLEDGMENTS The author wishes to thank Drs. V. Massey and L. M. Schopfer for providing material prior to publication. Work in the author's laboratory is supported by grants from the National Institutes of Health (AR 38917 and ES 04882) and the National Science Foundation (DMB04421).

REFERENCES 1. Hille, R. and Massey, V., Molybdenum-containing hydroxylases: xanthine oxidase, aldehyde oxidase, and sulfite oxidase, in Molybdenum Enzymes, Spiro, T. G-, Ed., John Wiley & Sons, New York, 1985, chap. 9. 2. Bray, R. C., The inorganic biochemistry of molybdoenzymes, Quart Rev. Biophys., 21, 3, 1988. 3. Cramer, S. P. and Stiefel, E. I., Chemistry and biology of the molybdenum cofactor, in Molybdenum Enzymes, Spiro, T. G., Ed., Wiley-Interscience, New York, 1985, Chap, 8. 4. Dixon, M. and Thurlow, S., Studies on xanthine oxidase: the dynamics of the oxidase system, Biochem. /., 18, 976, 1924. 5. Bray, R. C., Molybdenum iron-sulfur flavin hydroxylases and related enzymes, Enzymes, 12, 1975, chap. 6. 6. Massey, V., Iron-sulfur flavoprotein hydroxylases, in Iron-Sulfur Proteins, Vol. 1, Ehrenberg, A., Ed., Academic Press, New York, 1973, chap. 10. 7. Mackler, B., Mahler, H. R., and Green, D. E., Studies on metalloflavoproteins. I. Xanthine oxidase, a molybdoflavoprotein, J. Biol. Chem., 210, 149, 1954. 8. Palmer, G. and Massey, V., Electron paramagnetic resonance and circular dichroism studies on milk xanthine oxidase, J. Biol Chem., 244, 2614, 1969. 9. Corran, H. S., Dewann, J. G., Gordon, A. H., and Green, D. E., Xanthine oxidase and milk flavoprotein, Biochem. J., 33, 1694, 1939. 10. Hart, L. I., McGartoll, M. A., Chapman, H. R., and Bray, R. C., The composition of milk xanthine oxidase, Biochem. J., 116, 851, 1970. 11. Massey, V., Brumby, P. E., Komai, H., and Palmer, G., Studies on milk xanthine oxidase: some spectral and kinetic properties, J. Biol. Chem., 244, 1682, 1969. 12. Morrison, M. and Hultquist, D. E., Lactoperoxidase: isolation, J. Biol. Chem., 238, 2847, 1963. 13. Nathans, G. R. and Kirby Hade, E. P., Bovine milk xanthine oxidase: purification by ultrafiltration and conventional methods which omit addition of proteases; some criteria for homogeneity of native xanthine oxidase, Biochim. Biophys. Acta, 526, 328, 1978. 14. Massey, V. and Edmondson, D., On the mechanism of inactivation of xanthine oxidase by cyanide, J. Biol. Chem., 245, 6595, 1970. 15. Edmondson, D., Massey, V., Palmer, G., Beacham, L. M., HI, and Elion, G, B., The resolution of active and inactive xanthine oxidase by affinity chromatography, J. Biol. Chem., 247, 1597, 1972. 16. Massey, V., Komai, H., Palmer, G., and Elion, G. B., On the mechanism of inactivation of xanthine oxidase by allopurinol and other pyrazolo[3,4-d] pyrimidines, J. Biol. Chem,, 245, 2837, 1970. 17. Nishino, T., Nishino, T., and Tsushima, K., Purification of highly active milk xanthine oxidase by affinity chromatography on Sepharose 4B/folate gel, FEBS Lett., 131, 369, 198L 18. Hofstee, B. H. J., On the mechanism of inhibition of xanthine oxidase by the substrate xanthine, /. Biol. Chem., 230,235, 1955. 19. Olson, J. S., Ballou, D. P., Palmer, G., and Massey, V., The reaction of xanthine oxidase with molecular oxygen, /. Biol Chem., 249, 4350, 1974. 20. Palmer, G. and Olson, J. S., Concepts and approaches to the understanding of electron transfer processes in enzymes containing multiple redox centers, in Molybdenum and Molybdenum-Containing Enzymes, Coughlan, M., Ed., Pergamon Press, Elmsford, New York, 1980, Chap. 5. 21. Greenlee, L. and Handler, P., Xanthine oxidase: influence of pH on substrate specificity, J. Biol. Chem., 239, 1090, 1964.

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22. Edmondson, D., Ballou, D., Van Heuvelen, A., Palmer, G., and Massey, V., Kinetic studies on the substrate reduction of xanthine oxidase, J. Biol. Chem., 248, 6135, 1973. 23. Olson, J. S., Ballou, D. P., Palmer, G., and Massey, V., The mechanism of action of xanthine oxidase, J. Biol. Chem., 249,4363, 1974. 23a. D'Ardenne, S. C. and Edmondson, D. E., Kenetic isotope effect studies on milk xanthine oxidase and chicken liver xanthine dehydrogenase, Biochemistry, 29, 9046, 1990. 24. Komai, H., Massey, V., and Palmer, G., The preparation and properties of deflavo xanthine oxidase, J. Biol. Chem., 244, 1692, 1969. 25. Miiller, F., Hemmerich, P., Ehrenberg, A., Palmer, G., and Massey, V., The chemical and electronic structure of the neutral flavin radical as revealed by electron spin resonance spectroscopy of chemically and isotopically substituted derivatives, Eur. J, Biochem., 14, 185, 1970. 26. Hille, R., Hagen, W. R., and Dunham, W. R., Spectroscopic studies on the iron-sulfur centers of mik xanthine oxidase, J. Biol. Chem., 260, 10569, 1985. 27. Willis, L. J. and Loehr, T. M., Resonance Raman studies of the flavin and iron-sulfur centers of milk xanthine oxidase, Biochemistry, 24, 2678, 1985. 28. Peterson, J., Godfrey, C., Thomson, A. J., George, G. N., and Bray, R. C., Detection by lowtemperature circular-dichroism spectroscopy of optical absorption bands due to molybdenum(V) in the form of xanthine oxidase giving the desulpho inhibited e.p.r. signal, Biochem. J., 233, 107, 1986. 29. Davis, M. D., Olson, J. S., and Palmer, G., Charge transfer complexes between pteridine substrates and the active center molybdenum of xanthine oxidase, J. Biol. Chem,, 257, 14730, 1982. 30. Hille, R. and Massey, V., Tight binding inhibitors of xanthine oxidase, Pharmacol. Ther., 14, 249, 1981. 31. Kalckar, H., Kieldgaard, N. O., and Klenow, H., Inhibition of xanthine oxidase by 6-pteridylaldehyde, J. Biol. Chem., 174, 771, 1948. 32. Bray, R. C. and Vanngard, T., 'Rapidly appearing' molybdenum electron paramagnetic resonance signals from reduced xanthine oxidase, Biochem. J.t 114, 725, 1969. 33. Bray, R. C., Palmer, G., and Beinert, H., Direct studies on the electron transfer sequence in xanthine oxidase by electron paramagnetic resonance spectroscopy. II. Kinetic studies employing rapid freezing, J. Biol. Chem., 239,2667, 1964. 34. Tsopanakis, A. D., Tanner, S. J., and Bray, R. C., pH-jump studies at subzero temperatures on an intermediate in the reaction of xanthine oxidase with xanthine, Biochem. J., 175, 879, 1978. 35. Lowe, D. J., Barber, M. J., Pawlik, R. T., and Bray, R. C., A new nonfunctional form of milk xanthine oxidase containing stable quinquivalent molybdenum, Biochem. J., 155, 81, 1976. 36. Lowe, D. J. and Bray, R. C., Magnetic coupling of the molybdenum and iron-sulphur centers in xanthine oxidase and xanthine dehydrogenase, Biochem. J., 169, 471, 1978. 37. Coffman, R. E. and Buettner, G. R., General magnetic dipolar interaction of spin-spin coupled molecular dimers. Application to an EPR spectrum of xanthine oxidase, /. Phys, Chem., 83, 2392, 1979. 38. Barber, M. J., Salerno, J. C., and Siegel, L. M., Magnetic interactions in milk xanthine oxidase, Biochemistry, 21, 1648, 1982. 39. Beinert, H. and Orme-Johnson, W. H., Electron spin relaxation as a probe for active centers of paramagnetic species, in Magnetic Resonance in Biological Systems, Ehrenberg, A., Ed., Pergamon, Oxford, 1967, 221. 40. Cam mack, R., Barber, M. J., and Bray, R. C., Oxidation-reduction potentials of molybdenum, flavin, and iron-sulphur centers in milk xanthine oxidase, Biochem. J., 157, 469, 1976. 41. Williams-Smith, D. L., Bray, R. C., Barber, M. J., Tsopanakis, A. D., and Vincent, S. P., Changes in apparent pH on freezing aqueous buffer solutions and their relevance to biochemical electron-paramagnetic resonance spectroscopy, Biochem. J., 167, 593, 1977. 42. Hille, R., Fee, J. A., and Massey, V., Equilibrium properties of xanthine oxidase containing FAD analogs of varying oxidation-reduction potential, J. Biol. Chem., 256, 8933, 1981. 43. Hille, R., Redox interactions of the FAD in xanthine oxidase with other centers of the protein, in Flavins andFlavoproteins, Edmondson, D. E., and McCormick, D. B., Eds., Walter de Gruyter, New York, 1987, 391. 44. Hille, R. and Massey, V., The kinetic behavior of xanthine oxidase containing chemically modified flavius, J. Biol. Chem., 26, in press. 45. Barber, M. J. and Siegel, L. M., Oxidation-reduction potentials of molybdenum, flavin, and iron-sulfur centers in milk xanthine oxidase; variation with pH, Biochemistry, 21, 1638, 1982. 46. Hille, R. and Stewart, R. C., The inhibition of xanthine oxidase by 8-bromoxanthine, J. Biol. Chem.t 259, 1570, 1984. 47. Porras, A. G. and Palmer, G., The room temperature potentiometry of xanthine oxidase: pH dependent redox behavior of the flavin, molybdenum, and iron-sulfur centers, J. Biol. Chem., 257, 11617, 1982. 48. Spence, J. T., Barber, M. J., and Siegel, L. M., Determination of the stoichiometry of electron uptake and the midpoint reduction potentials of milk xanthine oxidase at 25°C by microcoulometry, Biochemistry, 21, 1656, 1982.

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49. Bhattacharyya, A., To 11 in, G. G., Davis, M., and Edmondson, D. E.; Laser flash photolysis studies of intramolecular electron transfer in milk xanthine oxidase, Biochemistry, 22, 5270, 1983. 50. Massey, V. and Hemmerich, P., Photoreduction of flavoproteins and other biological compounds catalyzed by deazaflavins, Biochemistry, 17, 9, 1978. 51. Edmondson, D. E., Hazzard, J. T., and Tollin, G., Laser flash photolysis studies of intramolecular electron transfer between the FAD and Fe/S II centers in xanthine oxidase, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. M., Eds., Walter de Gruyter, New York, 1987, 403. 52. Hille, R. and Massey, V., The presence of a reducible disulfide bond in milk xanthine oxidase, J. Biol. Chem., 257, 8898, 1982. 53. Hille, R. and Massey, V., The equilibration of reducing equivalents within milk xanthine oxidase, J. Biol. Chem., 261, 1241, 1986. 54. Anderson, R. F., Hille, R., and Massey, V., The radical chemistry of milk xanthine oxidase as studied by radiation chemistry techiques, J. Biol. Chem., 261, 15870, 1986. 55. Buxton, G. V., Basic radiation chemistry of liquid water, in The Study of Fast Processes and Transient Species by Electron Pulse Radiolysis, Baxendale, J. H. and Busi, F., Eds., D. Reidel Publishing Co., Dordrecht, Holland, 1982, 241. 55a. Hille, R., Unpublished results. 56. Komai, H. and Massey, V., Alkylation of milk xanthine oxidase, in Flavins and Flavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 399. 57. Skibo, E. B., Noncompetitive and irreversible inhibition of xanthine oxidase by benzimidazole analogues acting at the functional flavin adenine dinucleotide cofactor, Biochemistry, 25, 4189, 1986. 58. Porras, A. G., Olson, J. S., and Palmer, G., The reaction of reduced xanthine oxidase with oxygen: kinetics of peroxide and superoxide formation, J. Biol. Chem., 256, 9096, 1981. 59. Massey, V. and Hemmerich, P., Active site probes of flavoproteins, Biochem. Soc. Trans., 8, 246, 1980. 60. Hille, R. and Massey, V., Studies on the oxidative half-reaction of xanthine oxidase, J. Biol. Chem., 256, 9090, 1981. 61. McCord, J. M. and Fridovich, I., The reduction of cytochrome c by milk xanthine oxidase, J. Biol. Chem., 243, 5753, 1968. 62. Yonetani, T., Studies on cytochrome c peroxidase. II. Stoichiometry between enzyme, H2O2 and ferrocytochrome c and enzymic determination of extinction coefficients of cytochrome c, J. Biol. Chem., 240, 4509, 1965. 63. Rabani, J. and Nielsen, S. O., Absorption spectrum and decay of Oi and HO2 in aqueous solutions by pulse radiolysis, /. Phys. Chem., 73, 3736, 1969. 64. Fridovich, I., Quantitative aspects of the production of superoxide anion radical by milk xanthine oxidase, J. Biol. Chem., 245, 4053, 1970. 65. Janda, M. and Hemmerich, P., 5-Deaza- and 5-thiaflavins: a simple pathway to antimetabolites of vitamin B2, Angew. Chem. Intl. Ed. (EngL), 15, 443, 1976. 66. Fenner, H., Grauert, R., Hemmerich, P., Michel, H., and Massey, V., 5-Thia-5-deazaflavin, a 1 e"transferring flavin analog, Eur. J. Biochem., 95, 183, 1979. 67. Nishino, T., Tsushima, K., Hille, R., and Massey, V., Inhibition of xanthine oxidase by fluorodinitrobenzene, J. Biol. Chem., 257, 7348, 1982. 68. Pateman, J. A., Cove, D. J., Rever, B. M., and Roberts, D. B., A common cofactor for nitrate reductase and xanthine dehydrogenase which also regulates the synthesis of nitrate reductase, Nature, 201, 58, 1964. 69. Dixon, M. and Thurlow, S., Studies of xanthine oxidase. III. The reduction of nitrates, Biochem. J., 33, 989, 1924. 70. Ketchum, P. A., Cambier, H. Y., Frazier, W. A., HI, Madansky, C. H., and Nason, A., In vitro assembly of Neurospora assimilatory nitrate reductase from protein subunits of a Neurospora mutant and the xanthine oxidase or aldehyde oxidase systems of higher animals, Proc. Natl. Acad. Sci. U.S.A., 66, 1016, 1970. 71. Nason, A., Lee, K.-Y., Pan, S,-S., Ketchum, P. A., Lamberti, A., and DeVries, J., In vitro formation of assimilatory reduced nicotinamide adenine dinucleotide phosphate: nitrate reductase from a Neurospora mutant and a component of molybdenum-enzymes, Proc. Natl. Acad. Sci. U.S.A., 68, 3242, 1971. 72. Hawkes, T, R. and Bray, R. C., Quantitative transfer of the molybdenum cofactor from xanthine oxidase and from sulfite oxidase to the deficient enzyme of the nit-1 mutant of Neurospora crassa to yield active nitrate reductase, Biochem. J., 219, 481, 1984. 73. Johnson, J. L., Hainline, B. E., Rajagopalan, K. V., and Arison, B. H., The pterin component of the molybdenum cofactor: structural characterization of two fluorescent derivatives, J. Biol. Chem., 259, 5414, 1984. 74. Kramer, S. P., Johnson, J. L., Ribeiro, A. A., Millington, D. S., and Rajagopalan, K. V., The structure of the molybdenum cofactor: characterization of di-(carboxamidomethyl)molybdopterin from sulfite oxidase and xanthine oxidase, J. Biol. Chem., 262, 16357, 1987. 75. Fish, K. and Massey, V., The molybdenum cofactor of milk xanthine oxidase, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. M., Eds., Walter de Gruyter, New York, 1987, 421.

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76. Edmondson, D. E. and D'Ardenne, S. C., Electron nuclear double resonance spectroscopy of the desulfoinhibited molybdenum (V) center in bovine milk xanthine oxidase, Biochemistry, 28, 5924, 1989. 77. Taylor, E. C. and Reiter, L. A., Studies on the molybdenum cofactor. An unequivocal total synthesis of (±)-urothione, J. Am. Chem. Soc., I l l , 285, 1989. 78. Johnson, J. L. and Rajagopalan, K. V., Structural and metabolic relationship between the molybdenum cofactor and urothione, Proc. Natl. Acad. Sci. U.S.A., 79, 6856, 1982. 79. Cramer, S. P., Wahl, R., and Rajagopalan, K. V., Molybdenum sites of sulfite oxidase and xanthine dehydrogenase. A comparison by EXAFS, /. Am. Chem. Soc., 103, 7721, 198L 80. Cramer, S. P. and Hille, R., Arsenite-inhibited xanthine oxidase-determination of the Mo-S-As geometry by EXAFS, /. Am. Chem. Soc., 107, 8164, 1985. 81. Hawkes, T. R. and Bray, R. C., Studies by electron-paramagnetic-resonance spectroscopy of the environment of the metal in the molybdenum cofactor of molybdenum-containing enzymes, Biochem. J., 222, 587, 1984. 82. Stiefel, E. L, The coordination and bioinorganic chemistry of molybdenum Prog. Inorg. Chem., 22, 1, 1977. 83. Rajagopalan, K. V., Pterin cofactors, J. CellBiol., 107, 441a, 1988. 84. Lazarus, R. A., Dietrich, R. F., Wallick, D. E., and Benkovic, S. J., On the mechanism of action of phenylalanine hydroxylase, Biochemistry, 20, 6834, 1981. 85. Dix, T. A., Bollag, G. E., Domanico, P. L., and Benkovic, S. J., Phenylalanine hydroxylase: absolute configuration and source of oxygen of the 4a-hydroxytetrahydropterin species, Biochemistry, 24, 2955, 1985. 86. Davis, M. D., Edmondson, D. E., and Miiller, F., 31P nuclear magnetic resonance and chemical studies of the phosphorus residues in bovine milk xanthine oxidase, Eur. J. Biochem., 145, 243, 1984. 87. Berg, J. M., Hodgson, K. O., Cramer, S. R, Corbin, J. L., Elsberry, A., Pariyadath, N., and Stiefel, E. I., Structural results relevant to the molybdenum sites in xanthine oxidase and sulfite oxidase. Crystal structures of MoO2L, L=(SCH2CH2)2NCH2CH2X with X-SCH3, Af(CH3)2, J. Am. Chem. Soc., 101, 2774, 1979. 89. Bordas, J., Bray, R. C., Garner, C. D., Gutteridge, S., and Hasnain, S. S., X-ray absorption spectroscopy of xanthine oxidase, Biochem. J., 191, 499, 1980. 90. Teo, B. K., Ed., EXAFS: Basic Principles and Data Analysis, Springer-Verlag, New York, 1986. 91. Konigsberger, D. C. and Prins, R., X-Ray Absorption Spectroscopy: Principles, Applications, Techniques, of EXAFS, SEXAFS, andXANES, Wiley-Interscience, New York, 1988. 92. Scott, R. A., X-ray absorption spectroscopy, in Structural and Resonance Techniques in Biological Research, Chance, B., Ed., Bell Telephone Laboratories, Murray Hill, 1984, chap. 4. 93. Cotton, F. A. and Wing, R. M., Properties of metal-to-oxygen multiple bonds, especially molybdenumoxygen bonds, Inorg. Chem., 4, 867, 1965. 94. Green, D. E., Studies on reversible dehydrogenase systems. II. The reversibility of the xanthine oxidase system, Biochem. J., 28, 1550, 1934. 95. Harris, J. and Heller man, L., Sulfhydryl groups apparently essential to the activity of milk xanthine oxidase, in Inorganic Nitrogen Metabolism McElroy, W. D. and Glass, B., Eds., Johns Hopkins Press, Baltimore, 1956, 565. 96. George, G. N. and Bray, R. C., Formation of the inhibitory complex of p-mercuribenzoate with xanthine oxidase, evaluation of hyperfine and quadrupole couplings of mercury to molybdenum (V) from the electron paramagnetic resonance spectrum, and structure of the complex, Biochemistry, 22, 5443, 1983. 97. Hille, R., Stewart, R. C., Fee, J. A., and Massey, V., The interaction of arsenite with xanthine oxidase, J. Biol. Chem,, 258, 4849, 1983. 98. George, G. N. and Bray, R. C., Reaction of arsenite ions with the molybdenum center of milk xanthine oxidase, Biochemistry, 22, 1013, 1983. 99. Stewart, R. C., Hille, R., and Massey, V., Characterization of arsenite-complexed xanthine oxidase at room temperature, J. Biol. Chem., 259, 14426, 1984. 100. Bray, R. C. and Knowles, P. F., Electron spin resonance in enzyme chemistry: the mechanism of action of xanthine oxidase, Proc. Roy. Soc. A., 302, 351, 1968. 101. Stewart, R. C., Hille, R., and Massey, V., The reaction of arsenite-complexed oxidase with oxygen, J. Biol. Chem., 260, 8892, 1985. 102. Cramer, S. P., Biochemical application of x-ray absorption spectroscopy, in X-Ray Absorption Spectroscopy: Principles, Applications, Techniques of EXAFS, SEXAFS, and XANES, Konigsberger, D. C. and Prins, R., Eds., Wiley-Interscience, New York, 1988, chap. 7. 103. Gutteridge, S. and Bray, R. C., Oxygen-17 splitting of the very rapid molybdenum (V) e.p.r. signal from xanthine oxidase, Biochem. J.t 189, 615, 1980. 104. Hille, R. and Sprecher, H., On the mechanism of action of xanthine oxidase, /. Biol. Chem., 262, 10914, 1987. 105. Davis, M. D., Olson, J. S., and Palmer, G., The reaction of xanthine oxidase with lumazine, J. Biol. Chem., 259, 3526, 1984.

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106. Skibo, E. B., Gilchrist, J. H., and Lee, C.-H., Electronic probes of the mechanism of substrate oxidation by buttermilk xanthine oxidase: role of the active-site nucleophile in oxidation, Biochemistry, 26, 3032, 1987. 107. Skibo, E. B., Noncompetitive and irreversible inhibition of xanthine oxidase by benzimidazole analogues acting at the functional flavin adenine dinucleotide cofactor, Biochemistry, 25, 4189, 1986. 108. Morpeth, F. F. and Bray, R. C., Inhibition of xanthine oxidase by various aldehydes, Biochemistry, 23, 1332, 1984. 109. Murray, K. N., Watson, J. G., and Chaykin, S., Catalysis of the direct transfer of oxygen from nicotinamide-TV-oxide to xanthine by xanthine oxidase, J. BioL Chem., 241, 4798, 1966. 110. Bray, R. C. and Gutteridge, S., Numbers and exchangeability with water of oxygen-17 atoms coupled to molybdenum (V) in different reduced forms of xanthine oxidase, Biochemistry, 21, 5992, 1982. 111. Berg, J. M., and Holm, R. H., A model for the active sites of oxo-transfer molybdoenzymes: synthesis, structure, and properties, /. Am. Chem. Soc., 107, 917, 1985. 112. Berg, J. M. and Holm, R. H., A model for the active sites of oxo-transfer molybdoenzymes: reactivity, kinetics, and catalysis, J. Am. Chem. Soc., 107, 925, 1985. 113. Chen, G. J.-J., McDonald, J. W., and Newton, W. E., Synthesis of Mo(IV) and Mo(V) complexes using oxo abstraction by phosphines. Mechanistic implications, Inorg. Chem., 15, 2612, 1976. 114. Harlan, E. W.,Berg, J. M., and Holm, R. H., Thermodynamic fitness of molybdenum (IV, VI) complexes for oxygen atom transfer reactions, including those with enzymatic substrates, J. Am. Chem. Soc., 108, 6992, 1986. 115. Adams, M. W. W. and Mortenson, L. E., Mo reductases: nitrate reductase and formate dehydrogenase, in Molybdenum Enzymes, Spiro, T. G. Ed., John Wiley & Sons, New York, 1985, 519. 116. Hanson, G. R., Wilson, G. L., Bailey, T. D., Pilbrow, J. R., and Wedd, A. G., Multifrequency electron spin resonance of molybdenum (V) and tungsten (V) compounds, J. Am. Chem. Soc., 109, 2609, 1987. 117. Dowerah, D., Spence, J. T., Singh, Raghuvir, Wedd, A. G., Wilson, G. L., Farchione, F., Enemark, J. H., Kristofzski, J., and Bruck, M., Molybdenum (VI) and molybdenum (V) complexes with N,N'Dimethyl-A^,N'-bis(2-mercaptophenyl)ethylenediamine. Electrochemical and electron paramagnetic resonance models for the molybdenum (VI/V) centers of the molybdenum hydroxylases and related enzyme, J. Am. Chem. Soc., 109, 5655, 1987. 118. Hille, R., George, G. N., Eidsness, M. K., and Cramer, S. P., EXAFS analysis of xanthine oxidase complexes with alloxanthine, violapterin, and 6-pteridylaldehyde, Inorg. Chem., 28, 4018, 1989. 118a. Turner, N. A., Bray, R. C., and Diakun, G. P., Information from e.x.a.f.s. spectroscopy on the structures of different forms of molybdenum in xanthine oxidase and the catalytic mechanism of the enzyme, Biochem, /., 260,563, 1989. 119. Hawkes, T. R., George, G. N., and Bray, R. C., The structure of the inhibitory complex of alloxanthine (l//-pyrazolo[3m4- Phe, Met-110 -> Leu, Lys-204 -> Glu, His-217 -> Leu, and Tyr-224 -* Phe, did not significantly affect catalytic activity. Furthermore, the same authors suggest that Met-110 is not an essential amino acid, because in human DAAO this residue is substituted by Thr.114 On the contrary the replacements Tyr-228 -^ Phe and His-307 -^ Leu caused a complete loss of enzyme activity. 113 All these results are very much at variance with those obtained by chemical modification studies, as reported in Table 3. Both experimental approaches could be criticized. We shall briefly discuss these points, noting however that final settling of the controversy concerning the topography of the active site must await resolution of the three-dimensional structure. The chemical modification studies on DAAO were performed using highly specific labeling agents, such as N-chloro substituted amino acids, which very often modify only a single residue producing concomitant inactivation of the enzyme. In this case, it may be supposed that the overall conformation of the molecule is not greatly affected by such procedures, though the introduction of a small group could cause significant perturbations to the active site environment and magnify the primary inhibitory effect via steric hindrance or differences in chemical properties of the modified amino acid. The same criticism, however, can be applied to site specific mutagenesis approach, since it results in the presence of *'foreign" acylic side chains which could as well modify the protein architecture at the active site. Furthermore, in the case of DAAO, the mutagenized enzymes were produced in very minute quantities and enzymatic

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activities were measured using the reticulocyte lysate directly. This could give rise to problems with regard to measurement of mutated enzyme kinetic parameters, as indicated by the somewhat scattered values published. 112 ' 113 Moreover, for some chemical modification studies, detailed kinetic analyses of the enzyme mechanism indicated a particular catalytic step specifically involving the modified amino acid residue; the small amounts of mutagenized enzyme available, however, did not permit similarly detailed studies to be performed. Activity assays under certain conditions might fail to detect the specific involvement of an amino acid residue in steps which are not rate limiting. Recent evidence indicating the involvement of chemically modified residues in the active site of DAAO also comes from comparison with glycolate oxidase. The active site of the latter enzyme, which has been recently resolved by crystallographic studies at 2.2 A resolution, 115 shows an array of active site amino acid residues with a similar catalytic role: Arg257, Tyr-24 and Tyr-129 are involved in the substrate binding site, His-254 in proton abstraction from the substrate, while Lys-230 is located near the N(1)-C(2)=O locus of the flavin and Arg-309 electrostatically binds the phosphate group of FMN. The major difference concerns the basic residues which in glycolate oxidase play a different role from that proposed in Scheme 2 for DAAO. Although overall comparison of the primary structures of the two enzymes shows a low degree of homology, it is reasonable to expect a similar topology for the active sites of these two oxidases. We feel in conclusion that chemical modification studies on DAAO still offer a valid method of probing the active site of this enzyme; in the meantime, more refined kinetic analyses of mutagenized enzymes are awaited. D. CLONING AND GENE EXPRESSION Three different laboratories116-118 have attempted molecular cloning and DNA sequencing of DAAO but only Miyake's group117 has so far been able to clone and sequence the cDNA of the complete pig kidney enzyme and, more recently, also of the human kidney D-amino acid oxidase.1M The cDNA for the pig kidney enzyme consists of 3211 nucleotides, presenting a 5'-terminus untranslated region of 198 nucleotides followed by an open reading frame of 1041 nucleotides which encodes the 347 amino acids of the enzyme and a 3'-terminus untranslated region of 1972 nucleotides. Comparison of the cDNA sequence with the amino acid sequence indicates that no signal peptide is present at the N-terminal region of the protein. The in vitro translation of capped RNA transcript yielded a protein with significant catalytic activity, which was strongly inhibited by benzoate.119 Southern blot analysis revealed the presence of a single gene for DAAO in the pig genome. The existence of distinct mRNA species in pig kidney, liver, and brain119 is indicative of some tissue-specific regulatory process for enzyme expression. A human cDNA with a length of 2 Kbase pairs has been isolated which codes for the complete DAAO sequence. The molecular weight of the human kidney enzyme has been calculated as 39,410 from the derived sequence of 347 amino acids (see Figure 5). 84.4% of residues are identical to those of the pig enzyme, but Met-110 is replaced by Thr. All other amino acid residues implicated in the active site (see Section II.C) are conserved.114 The construction and the expression in E. coli of semisynthetic DNA encoding complete DAAO has also been reported.120 As already noted,14'121 pig D-amino acid oxidase is a peroxisomal enzyme. A similar localization has been found for the Rhodotorula gracilis enzyme.69 No specific studies have been carried out on the transport of this enzyme into peroxisomes. The enzyme is synthesized on free ribosomes122 and because of the lack of a signal peptide, it is expected that the mature protein would contain all the information necessary for its translocation into peroxisomes. Recently it has been reported that various peroxisomal proteins have at or near their C-termini a tripeptide, Ser-Lys/His-Leu which could function as a targeting signal;123 interestingly, D-amino acid oxidase has the sequence Ser-His-Leu at its C-terminus.

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E. THE PHYSIOLOGICAL ROLE The physiological role of D-amino acid oxidase is still the subject of considerable debate. In animals, D-amino acid oxidase is localized mainly in kidney, cerebellum, and liver and this must be related in some way to its metabolic function. Localization in peroxisomes is linked to the production of H2O2, but could indicate the existence of a specialized function in metabolism. In an attempt to identify the physiological role of the enzyme, two main lines of research have been pursued. The first retains that D-amino acid oxidase is mainly involved in D-amino acid metabolism. Thus, free D-amino acids have been detected in human and mice plasma, and in increasing quantities in senescence and renal diseases.]24-125 Furthermore, in DAAO" mutant mice the level of neutral D-amino acid, and in particular of D-alanine, is increased in urine126-127 and D-amino acid levels in various organs are significantly higher than those found in DAAO + mice.128 These findings are all consistent with a role for the enzyme in D-amino acid metabolism, even though the comparatively low level of D-amino acids in tissues128 contrast with the high KM of the enzyme for these substrates. The second line of research is based on the assumption that metabolites other than Damino acids may represent the physiological substrates of the enzyme. This suggestion was first formulated by Hamilton52'129 who argued that potential DAAO substrates may originate from the reaction of amines with glyoxylate; it followed, among others, the observation that 7V-alkyl D-amino acids and D-proline are good substrates for the enzyme. In fact, pig kidney DAAO catalyzes oxygen uptake in the presence of relatively high concentrations of glyoxylate plus cysteamine, the activity being higher at the physiological pH of 7.4 than at 8.3. The true substrate would be the adduct formed from the two reagents, thiazolidine-2-carboxylate52 (see Section II.A). This reaction has been confirmed by other laboratories,53-22 but the physiological significance of the reaction in vivo is still a matter of controversy.

III. L-AMINO ACID OXIDASES L-amino acid oxidase activities have been detected in mammals,2-130 birds,13 reptiles,131 invertebrates,132 molds,2-133 and bacteria.2'9-134 In mammals, the L-amino acid oxidase first isolated by Blanchard et al.130 from rat liver and further investigated by Nagano et al.,135 utilizes FMN as coenzyme and is more active on L-hydroxy acids than amino acids. The enzyme properties were substantially re-examined by Cromartie and Walsh136 who consider this flavoprotein as an L-hydroxy acid oxidase rather than an L-amino acid oxidase. In birds a partially purified LAAO obtained from turkey liver13 also appears to have FMN as the prosthetic group. In bacteria, LAAO activity was first detected in Proteus vulgaris9 where it is located in the membrane.137 The Proteus enzyme, like the bacterial D-amino acid oxidase10"12 is a dehydrogenase which uses oxygen as an electron acceptor if coupled to the respiratory system.137 Recently, LAAO activity was found associated with photosystem II of Cyanobacteria and the enzyme partially purified.138 Extensive studies on structural and kinetic properties of LAAO have only been carried out, however, on ophidian enzymes, in particular on the LAAO from Crotalus adamanteus to which we shall mainly refer in what follows. A. CHEMICAL AND PHYSICAL PROPERTIES L-amino acid oxidases from snake venoms have been purified by various procedures involving heat treatment, salt fractionation, gel filtration, ion exchange chromatography, and crystallization.3-139'140 The purified LAAO is substantially stable if maintained in the dark at 0°C, temperature below -5°C causing denaturation of the enzyme.141 Substrates protect LAAO from pH or heat inactivation.140"142 The enzymes isolated from Crotalus adamanteus and Agkistrodon piscivorus are dimers of similar subunits having molecular weights ranging from 130,000 to 140,000 and containing 1 mol of noncovalently bound

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FIGURE 6. Absorbance spectra ofCrotalus adamanteus L-amino acid oxidase. (1) oxidized enzyme; (2) as in (1), after 4 h of irradiation in the presence of 0.09 M EDTA; (3) as in (1), after reduction of the enzyme with an excess of either substrate L-leucine and L-phenylalanine under anaerobic conditions. (From Massey, V. and Curti, B., /. Biol. Chem., 242, 1259, 1967. With permission.)

FAD/monomer.139-140 The scattering of molecular weight values is partially due to the relatively different sugar content. A complete analysis of carbohydrates has been performed only by de Kok and Rawitch.143 Sedimentation coefficients of 6.5 and 6.9 S have been determined for the C. adamanteus and A. piscivorus enzymes, respectively.3'139-140 Three electrophoretically distinct isoenzymes, A, B, and C, have been detected in varying amounts and isolated from C. adamanteus L-amino acid oxidase.140'143 The isoenzyme pattern has been interpreted as representative of subunit composition of the type aa, bb, ab.143 The amino acid composition of the A isoenzyme and of a mixture of the three isoenzymes has been determined by various laboratories with slightly different results.143-144 In particular, the half-cystine content is variable;143-144 according to de Kok and Rawitch143 only four are present as free sulfhydryl groups. Electrofocusing analysis of the C. adamanteus enzyme revealed that the original three isoenzyme pattern could be further resolved into at least eighteen enzymatically active components with some differences in their amino acid composition.145 This microheterogeneity, which partially overlaps the original isoenzyme distribution, could be accounted for by differences in carbohydrate content, in addition to differences in amino acid composition. In spite of this microheterogeneity, the two subunits present a high degree of homology as shown by finger-print analysis following tryptic digestion of the alkylated enzyme:143 in fact, the number of the tryptic peptides is approximately half the value that one would expect from the total arginines plus lysines content of the dimer. Failure to obtain a reconstitutable apoprotein has not allowed studies on the FADapoprotein complex or the use of flavin analogues as active site probes of the enzyme. The spectra of oxidized, fully reduced and one-electron reduced flavin are reported in Figure 6. The enzyme behaves like a flavoprotein oxidase: it reacts rapidly with oxygen, stabilizes the red anionic form of the flavin radical obtained through the EDTA-light procedure and reacts with sulfite.39'41 A Kd of 1.5 x 10~ 4 M has been reported for the sulfiteLAAO complex.41 As in the case of DAAO, sodium borohydride reacts with LAAO leading to the formation of 3,4-dihydroflavin.44 No trace of semiquinone was detected in the reaction of the reduced enzyme with oxygen.

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Chemistry and Biochemistry of Flavoenzymes

Potentiometric titrations on the LAAO from C. adamanteus allowed determination of the redox potentials of both the red semiquinone and anionic dihydroquinone couples. The first electron potential (EF1OX/EF1') appeared pH independent, with an EO of — 0.056 V (pH = 7.8); the second electron transfer potential EF17EFlredH~ was pH-dependent, with an EQ of —0.175 V (pH — 7.8) and a 0.060 V/pH unit slope. The semiquinone species is thermodynamically stable.146 Following the early work of Zeller et al., 147 several studies on the substrate specificity of ophidian L-amino acid oxidases were performed.2-3 At pH 7.0 the preferred substrates appeared to be aromatic or more generally hydrophobic amino acids; polar and basic amino acids being deaminated at much lower rates. Further studies revealed that substrate specificity varied according to the source of the enzyme2 and the pH used in the assay mixture,148 Shape and optimum pH curves depend markedly on the amino acid used as substrate;148 polar, basic amino acids, and glycine are oxidatively deaminated at comparable rates when assayed at their pH optima.148-149 Conversely, L-glutamic acid, Lraspartic acid, and L-proline were not attacked by the enzyme.23 It must be pointed out that all these pH studies were carried out at a single amino acid and O2 concentration, whereas a proper study would require determination of Vmax at saturated concentrations of both substrates. LAAO is also active towards a series of ring substituted amino acids, the rate of deamination being strictly correlated with the nature and position of the substituent.150 Seleno cysteinyl derivatives are also substrates for the enzyme. 151 Like all flavooxidases, LAAO is quite specific for oxygen; very low activities were observed with ferricyanide, methylene blue or iodonitrotetrazolium as final acceptors.41J4J - 152 Aromatic carboxylates are competitive inhibitors of LAAO. Unlike DAAO, L-amino acid oxidase weakly binds benzoate.153 The inhibitors give distinct resolved spectra with LAAO. The Ki and Kd determined through kinetic or spectral titration experiments were in good agreement and of the order of 10~3 M.153 LAAO undergoes reversible inactivation of two types. The first type of inactivation is obtained by raising the pH of the solution from 5.5 to 7.5 and the temperature from 25 to 38°C.154 Temperature influences the rate of the process whereas the pH affects the extent of inactivation. Monovalent anions such as chloride, substrates, and competitive inhibitors of LAAO, prevent inactivation.155 Reactivation is observed by lowering the pH to 5 and incubating the enzyme solution at 38°C for 60 min. The second type of inactivation is caused by storing the enzyme at temperatures between — 5° and — 60°C.141 The rate of inactivation is dependent on the storage pH and on the ionic composition of the medium, but unlike the first type, this inactivation is favored by low pH and is not prevented by chloride ions. Heating the enzyme at 38°C for 60 min at pH 5.0 causes reactivation. Neither type of inactivation involves major changes in the physical properties of the enzyme except for changes in absorption and ORD spectra.141-156 Spectral perturbations are clearly different for the two types of inactivation; in both cases reactivation regenerates the spectrum of native enzyme. These differences were confirmed and extended by Coles et al.157 through circular dichroism studies on complex formation of inactivated enzyme with sulfite or with anthranilic acid, an inhibitor which binds to both inactivated forms. These studies141'156"158 suggest that the two types of inactivation are due to a limited conformational change at the catalytic site of the enzyme which affects both the flavin and the substrate binding region. Differences between the two inactivated enzyme forms were also found by immunochemical techniques:159 in both, the differences disappeared on reactivation of the enzyme. It is interesting to note, in this contest, that antibodies raised against the native enzyme only partially inhibit catalytic activity, as is the case with pig kidney D-amino acid oxidase.160 A third form of inactivated LAAO, the so called y form has been described144 and derives from pH-heat inactivated LAAO following treatment with /?-chloromercuribenzoate.

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TABLE 4 Substrate Specificity and Turnover Numbers of Crotalus adamanteus L-Amino Acid Oxidase162 Substrate

L-arginine L-phenylalanine L-leucine L-valine

Turnover number (moles substrate oxidized/min/mol EFAD) 1,100 280 600 40

B. KINETIC MECHANISM AND ACTIVE SITE STUDIES The kinetic mechanism of LAAO from C. adamanteus has been thoroughly investigated using both steady-state and rapid-reaction techniques. It is noteworthy that no kinetic heterogeneity was observed despite the presence of many isozymes in the purified preparation. Initial attempts to elucidate the kinetic mechanism of LAAO were by Wellner and Meister:161 they explained substrate inhibition of enzymatic activity by an ingeneous scheme involving an enzyme with 2 mol of FAD/active site and a biradical intermediate, E(FADH')2, which reacted more rapidly with the substrate than the fully reduced enzyme, E(FADH2)2. Evidence against this mechanism was presented by Massey and Curti162-163 who put forward a kinetic scheme substantially similar to that proposed for DAAO (Scheme 1 and Table 4). Bright and Porter6 confirmed the latter scheme and extended their kinetic studies by examining the effect of pH on the enzyme reaction.164 Using phenylglycine as substrate and a wide range of O2 concentrations, they observed parallel lines in steady state kinetic experiments when amino acid concentration was varied, but biphasic parallel lines when O2 concentration was varied.164 They also found that the rate constant for the dissociation of Er-P to Er was pH dependent, being smaller at low pH. These results,162"164 are consistent with a kinetic mechanism involving either loop A or loop B or both of Scheme 1, depending on the substrate and pH used. At high pH and O2 concentration lower than 10~4 M, loop B is operative, whereas at low pH and O2 concentration greater or equal to 5 x 10~ 3 M, loop A becomes effective. The curved region of the biphasic lines observed at different O2 concentrations6-164 was interpreted as due to both mechanisms being operative. The inhibition of enzyme activity at high substrate concentration was better explained6*162-163 by assuming the formation of a spurious complex between reduced enzyme and substrate, Er*"S; if this reacts more slowly with oxygen than Er or Er—P, it will tend to accumulate during turnover in presence of high concentrations of substrate, thus explaining the inhibition phenomenon. The catalytic mechanism of flavin reduction in the LAAO reaction (i.e., E0'S —> Er*P, see Scheme 1) is similar to that observed for DAAO, and studies using a(2H)phenylalanine165 have shown C-a proton abstraction from the substrate occurs during the rate limiting step of flavin reduction. A pH dependent deuterium kinetic isotope effect was also observed in the reductive reaction with D,L-ot-2H-leucine as substrate. This was more marked at low pH and was interpreted as due to a change in the rate of C-H bond cleavage as the pH changed from 6 to 9.166-167 In the case of LAAO too, the a-imino acid appears to be the first product released by the enzyme, as shown by experiments in which cyanide was used as a trapping agent.168 Little data are available concerning the active site of LAAO. Studies have been hampered, as already noted, by the fact that the apoprotein cannot be reconstituted by addition of exogeneous FAD to give an active holoenzyme. Chemical modification studies on LAAO are also very limited. Apparently, sulfhydryl reagents are without effect on catalytic activity. 144 Conversely, butanedione causes irreversible loss of activity, arguing for the presence

88

Chemistry and Biochemistry of Flavoenzymes

of arginine(s) at the active site.169 This conclusion is supported by the observation that substrates such as L-phenylalanine and L-methionine protect the enzyme from butandione inactivation.169 A decrease of enzyme activity has been observed also by treating LAAO with diethyl pyrocarbonate;170 hydroxylamine partially reverses this inhibition. Spectroscopic titrations of the treated enzyme revealed the modification of several histidyl residues, but no significant correlation was observed between loss of activity and number of modified residues. The presence of a histidine at the active site was postulated by Singer and Kearney.139 Indirect evidence for the role of an imidazole side chain in the catalytic mechanism has been postulated by Page and Van Etten,167 but this conclusion has been criticized by Porter and Bright.164 The latter authors found that the pK attributed to a histidyl residue acting in the a-proton transfer mechanism could be assigned to the substrate instead of a side chain residue of the enzyme. It must be pointed out that the structural identification of a specific amino acid residue in the enzyme molecule was not accomplished in the above studies. Finally, studies of the reaction of LAAO with vinylglycine171 have shown that the reagent acts as a suicide inhibitor of the enzyme causing a progressive, irreversible inactivation. The loss of catalytic activity is probably related to covalent modification of a nucleophilic group at the enzyme active site, but no specific amino acid residue has been identified. C. THE PHYSIOLOGICAL ROLE The physiological role of LAAO is rather obscure. In mammals, as already pointed out, the role of the enzyme has been questioned, what was once an L-amino acid oxidase of rat kidney is now an L-hydroxy acid oxidase, apparently located both in mitochondria and in peroxisomes.136 With the exception of ophidian L-amino acid oxidases, the stoichiometry of the catalyzed reaction has not been investigated and it is likely that true L-amino acid dehydrogenases have been mistaken for L-amino acid oxidases. On the other hand, the high rate of L-amino acid oxidation found by Krebs1'2 in mammalian kidney homogenates is not consistent with the low turnover number of the former LAAO described by Nagano et al. ,135 thus suggesting that the metabolism of L-amino acids follows other pathways, such as the transaminase-glutamate dehydrogenase system. In species other than higher organisms or mammals, LAAO could be associated with a specific function. In cyanobacterium Anacystis nidulans, LAAO has been isolated as a component of photosystem II.138 In the early stages of evolution, the enzyme could have functioned as a scavenger for oxygen production by photosystem II.172 Oddly enough, the metabolic function of LAAO appears to be more elusive than that of DAAO, in spite of the fact that the former enzyme deals with the physiological L-isomers of amino acids. The hypothesis that LAAO is an ancestral enzyme which in the early times extracted energy by L-amino acid oxidation and was later converted to some secondary function deserves, perhaps, further consideration.

ACKNOWLEDGMENTS The authors would like to thank Drs. A. Negri and L. Pollegioni for helpful discussion and critical reading of the manuscript. The studies in the authors' laboratories were in part supported by grants from Ministero della Pubblica Istruzione and C.N.R., of Italy.

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63. Antonini, E., Brunori, M., Bruzzesi, M. R., Chiancone, E., and Massey, V., Association-dissociation phenomena of D-amino acid oxidase, J. BioL Chem., 241, 2358, 1966. 64. Tojo, H., Horiike, K., Shiga, K., Nishina, Y., Watari, H., and Yamano, T., Self-association mode of a flavoenzyme D-amino acid oxidase from hog kidney. I. Analysis of apparent weight-average molecular weight data for the apoenzyme in terms of model, J. BioL Chem., 260, 12607, 1985. 65. Tojo, H., Horiike, K., Shiga, K., Nishina, Y., Watari, H., and Yamano, T., Self-association mode of a flavoenzyme D-amino acid oxidase from hog kidney. II. Stoichiometry of holoenzyme association and energetics of subunits association, J. BioL Chem,, 260, 12615, 1985. 66. Horiike, K., Shiga, K., Isomoto, A., and Yamano, T., Effect of quasi-substrate on the monomer-dimer equilibrium of D-amino acid oxidase, J. Biochem. (Tokyo), 75, 925, 1974, 67. 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J., Chlorination of an active site tyrosyl residue of Damino acid oxidase by N-chloro-D-leucine, /. BioL Chem., 255, 498, 1980. 104. D'Silva, C., Williams, C. H., Jr., and Massey, V., Electrophilic amination of a single methionine residue located at the active site of D-amino acid oxidase by 0-(2,4-dinitrophenyl)-hydroxylamine, Biochemistry, 25, 5602, 1986. 105. D'Silva, C., Williams, C. H., Jr., and Massey, V., Identification of methionine-110 as the residue covalently modified in the electrophilic inactivation of D-amino acid oxidase by O-(2,4-dinitrophenyl)hydroxylamine, Biochemistry, 26, 1717, 1987. 106. Ronchi, S., Minchiotti, L., Galliano, M., Curti, B., Swenson, R. P., Williams, C. H., Jr., and Massey, V., The complete amino acid sequence of D-amino acid oxidase from pig kidney, in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Jr., Eds., Elsevier/North-Holland, Amsterdam, 1982, 66. 107. Miyake, Y., Fukui, K., Momoi, K., Watanabe, F., and Shibata, T., Biological and medical aspects of D-amino acid oxidase — Biogenesis and in vivo reaction with D-propargylglycine, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., de Gruyter & Co., Berlin, 1987, 501. 108. Marcotte, P. and Walsh, C., Properties of D-amino acid oxidase covalently modified upon its oxidation of D-propargylglycine, Biochemistry, 17,2864, 1978. 109. Singer, T. P. and Guzman Barron, E. S., Studies on biological oxidations. XX. Sulfhydryl enzymes in fat and protein metabolism, J. BioL Chem., 157, 241, 1945. 110. Fonda, M. L. and Anderson, B. M., D-amino acid oxidase. IV, Inactivation by maleimides, J. BioL Chem., 244, 666, 1969. 111. Karplus, A. P. and Schulz, G. E., Refined structure of glutathione reductase at 1.54 A resolution, J. Mol. BioL, 195, 701, 1987. 112. Watanabe, F., Fukui, K., Momoi, K., and Miyake, Y., Effect of site-specific mutagenesis of tyrosine55, methionine-110 and histidine-217 in porcine kidney D-amino acid oxidase on its catalytic function, FEBSLett., 238, 269, 1988. 113. Watanabe, F., Fukui, K., Momoi, K., and Miyake, Y., Site-specific mutagenesis of lysine-204, tyrosine224, tyrosine-228 and histidine-307 of porcine kidney D-amino acid oxidase and the implication as to its catalytic function, J. Biochem. (Tokyo), 105, 1024, 1989. 114. Momoi, K., Fukui, K., Watanabe, F., and Miyake, Y., Molecular cloning and sequence analysis of cDNA encoding human kidney D-amino acid oxidase, FEBS Lett., 238, 180, 1988. 115. Lindqvist, Y. and Branden, C. I., The active site of spinach glycolate oxidase, /. BioL Chem., 264, 3624, 1989.

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116. Inagaki, T., Ohishi, N., Tsukagoshi, N., Ichihara, C., Udaka, S., and Yagi, K., Translation of messenger RNA of pig kidney D-amino acid oxidase in a cell-free system, Biochem. Intern., 13, 1045, 1986. 117. Fukui, K., Watanabe, F., Shibata, T., and Miyake, Y., Molecular cloning and sequence analysis of cDNAs encoding porcine kidney D-amino acid oxidase, Biochemistry, 26, 3612, 1987. 118. Jacobs, P., Brockly, F., Massaer, M., Loriau, R., Guillaume, J. P., Ciccarelli, E., Heinderyckx, M., Cravador, A., Biemans, R., van Elsen, A., Herzog, A., and Bollen, A., Porcine D-amino acid oxidase: determination of mRNA nucleotide sequence by the characterization of genomic and cDNA clones, Gene, 59, 55, 1987. 119. Fukui, K., Momoi, K., Watanabe, F., and Miyake, Y., In vivo and in vitro expression of porcine Damino acid oxidase: in vitro system for the synthesis of a functional enzyme, Biochemistry, 27, 6693, 1988. 120. Ciccarelli, E., Massaer, M., Guillame, J.-P., Herzog, A., Loriau, R., Cravador, A., Jacobs, P., and Bolien, A., Porcine D-amino acid oxidase: production of the biologically active enzyme in Escherichia coli, Biochem. Biophys. Res. Comm., 161, 865, 1989. 121. Perotti, M. E., Gavazzi, E., Trussardo, L., Malgaretti, N., and Curti, B., Immunoelectron microscopic localization of D-amino acid oxidase in rat kidney and liver, Histochem. J., 19, 157, 1987. 122. Fukui, K., Momoi, K., Watanabe, F., and Miyake, Y., Biosynthesis of porcine kidney D-amino acid oxidase, Biochem, Biophys. Res. Comm., 141, 1222, 1986. 123. Gould, S. G., Keller, G. A., and Subramani, S., Identification of peroxisomal targeting signals located at the carboxy terminus of four peroxisomal proteins, J. Cell. Biol., 107, 897, 1988. 124. Nagata, Y., Akino, T., Ohno, K., Katakoa, Y., Ueda, T., Sakurai, T., Shiroshita, K., and Yasuda, T., Free D-amino acids in human plasma in relation to senescence and renal diseases, Clin. Sci., 73, 105, 1987. 125. Nagata, Y., Akino, T., and Ohno, K M The presence of free D-amino acids in mouse tissues, Experientia, 45, 330, 1989. 126. Konno, R., Isobe, K., Niwa, A., and Yasamura, Y., Lack of D-amino acid oxidase activity causes a specific renal aminoaciduria in the mouse, Biochim. Biophys. Acta, 967, 382, 1988. 127. Konno, R., Nagata, Y., Niwa, A., and Yasamura, Y., Spontaneous excretion of D-alanine in urine in mutant mice lacking D-amino acid oxidase, Biochem. J., 261, 285, 1989. 128. Nagata, Y., Konno, R., Yasamura, Y., and Akino, T., Involvement of D-amino acid oxidase in elimination of free D-amino acids in mice, Biochem. J . , 257, 291, 1989. 129. Hamilton, G. A., Peroxisomal oxidases and suggestions for the mechanism of action of insuline and other hormones, Adv. EmymoL, 57, 85, 1985. 130. Blanchard, M., Green, D. E., Nocito, V., and Ratner, S., L-amino acid oxidase of animal tissue, /. BioL Chem., 155, 421, 1944. 131. Zeller, E. A. and Maritz, A., Ubereine neue L-Aminosaure-oxydase, Helv. Chim. Acta, 27, 1888, 1944. 132. Roche, J., Glahn, P. E., Manchon, P., and Thoai, N. V., Sur une nouvelle L-amino acide oxydase, activable par le magnesium, Biochim. Biophys. Acta, 35, 111, 1959. 133. Thayer, P. S. and Horowitz, N. H., The L-amino acid oxidase of Neurospora, J, Biol Chem., 192, 755, 1951. 134. Szwajcer, E., Brodelius, P., and Mosbach, K., Production of oc-keto acids. II. Immobilized whole cells of Providencia sp. PCM 1298 containing L-amino acid oxidase, Enzyme Microb. TechnoL, 4, 409, 1982. 135. Nagano, M. and Danowski, M. N., Crystalline mammalian L-amino acid oxidase from rat kidney mitochondria, J. Biol. Chem., 241, 2075, 1966. 136. Cromartie, T. H. and Walsh, C. T., Rat kidney L-a-hydroxy acid oxidase: isolation of enzyme with one flavin coenzyme per two subunits, Biochemistry, 14, 2588, 1975. 137. Pelmont, J., Arlaud, G., and Rossat, A., L-amino acide oxidases des enveloppes de Proteus mirabilis: properties generales, Biochemie, 54, 1359, 1972. 138. Pistorius, E. K. and Gau, A. E., Presence of a flavoprotein in O2-evolving Photosystem II preparations from the cyanobacterium Anacystis nidulans, Biochim. Biophys. Acta, 849, 203, 1986. 139. Singer, T. P. and Kearney, E. B., The L-amino acid oxidases of snake venom. II. Isolation and characterization of homogeneous L-amino acid oxidase, Arch. Biochem. Biophys., 29, 190, 1950. 140. Wellner, D. and Meister, A., L-amino acid oxidase of Crotalus adamanteus, J. Biol. Chem., 235, 2013, 1960. 141. Curti, B., Massey, V., and Zmudka, M., Inactivation of snake venom L-amino acid oxidase by freezing, J. BioL Chem., 243, 2306, 1968. 142. Paik, W, K. and Kim, S., Studies of the stability of L-amino acid oxidase of snake venom, Biochim. ftiophys. Acta, 139, 49, 1967. 143. de Kok, A. and Rawitch, A. B., Studies on L-amino acid oxidase. II. Dissociation and characterization of its subunits, Biochemistry, 8, 1405, 1969. 144. Wellner, D. and Hayes, M. B., Multiple molecular forms of L-amino acid oxidase, Ann. N. Y. Acad. Sci., 151, 118, 1968.

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145. Hayes, M. B. and Wellner, D., Microheterogeneity of L-amino acid oxidase, /. BioL Chem., 244, 6636, 1969. 146. Stankovich, N. T. and Fox, B. G., Redox potential-pH properties of the flavoprotein L-amino acid oxidase, Biochim. Biophys, Acta, 786, 49, 1984. 147. Zeller E. A., Maritz, A., and Iselin, B., Uber eine neue L-Aminosaure-oxydase (Ophio-L-aminosaureoxydase), Helv. Chim. Acta, 30, 1615, 1945. 148. Paik, W. K. and Kim, S., pH-substrate relationship of L-amino acid oxidase from snake venom and rat kidney, Biochim. Biophys. Acta, 96, 66, 1965. 149. Zeller, A. E., Ramachander, G., Fleisher, G. A., Ishimaru, T., and Zeller, V., Ophidian L-amino acid oxidase, Biochem. J., 95, 262, 1965. 150. Zeller, A. E., Clauss, L. M., and Ohlsson, J. T., Interaction of ophidian L-amino acid oxidase with its substrates and inhibitors: role of molecular geometry and electron distribution, Helv. Chim. Acta, 57, 2406, 1974. 151. Coccia, R., Blarzino, C., Foppoli, C., and Cini, C., Oxidative deamination of Se-(l-carboxyethyl)-, Se(1-carboxypropyl)- and Se-(2-carboxyethyl)-seleno cysteine by snake venom L-amino acid oxidase, Physiol. Chem. Phys. Med. NMR, 20, 115, 1988. 152. Dixon, M. and Webb, E. C., Enzymes, 3rd ed., Longman, London, 1979, 132. 153. de Kok, A. and Veeger, C., Studies on L-amino acid oxidase. I. Effects of pH and competitive inhibitors, Biochim. Biophys. Acta, 167, 35, 1968. 154. Kearney, E. B. and Singer, T. P., The L-amino acid oxidases of snake venom. III. Reversible inactivation of L-amino acid oxidases, Arch. Biochem. Biophys., 33, 377, 1951. 155. Kearney, E. B. and Singer, T. P., The L-amino acid oxidases of snake venom. IV. The effect of ion on the reversible inactivation, Arch. Biochem. Biophys., 33, 397, 1951. 156. Wellner, D., Evidence for conformational changes in L-amino acid oxidase associated with reversible inactivation, Biochemistry, 5, 1585, 1966. 157. Coles, C. J., Edmondson, D. E., and Singer, T. P., Reversible inactivation of L-amino acid oxidase. Properties of the three conformational forms, J. BioL Chem.., 252, 8035, 1977. 158. Kearney, E. B. and Singer, T. P., The L-amino acid oxidases of snake venom. V. Mechanism of the reversible inactivation, Arch. Biochem. Biophys., 33, 414, 1951. 159. Zimmerman, S. E., Brown, R. K., Curti, B., and Massey, V., Immunochemical studies of L-amino acid oxidase, Biochim. Biophys. Acta, 229, 260, 1971. 160. Gavazzi, E., Malgaretti, N., and Curti, B., Immunochemical properties of D-amino acid oxidase, Biochim. Biophys. Acta, 915, 188, 1987. 161. Wellner, B. and Meister, A., Studies on the mechanisms of action of L-amino acid oxidase, J. BioL Chem., 236, 2357, 1961. 162. Massey, V. and Curti, B., On the reaction mechanism of Crotalus adamanteus L-amino acid oxidase, J. BioL Chem., 242, 1259, 1967. 163. Massey, V. and Curti, B., Comparative studies of D- and L-amino acid oxidases, in Flavin and Flavoproteins, Yagi, K., Ed., University of Tokyo Press, Tokyo, 1968, 226. 164. Porter, D. J. T. and Bright, H. J., Interpretation of the pH dependence of flavin reduction in L-amino acid oxidase reaction, J. BioL Chem., 255, 2969, 1980. 165. Porter, D. J. T. and Bright, H. J., Location of hydrogen transfer steps in the mechanism of reduction of L-amino acid oxidase, Biochem. Biophys. Res. Comm., 36, 209, 1969. 166. Page, D. S. and Van Etten, R. L., L-amino acid oxidase. I. Effect of pH, Biochim. Biophys. Acta, 191, 38, 1969. 167. Page, D. S. and Van Etten, R. L., L-amino acid oxidase. II. Deuterium isotope effects and the action mechanism for the reduction of L-amino acid oxidase by L-leucine, Biochim. Biophys. Acta, 227, 16, 1971. 168. Porter, D. J. T. and Bright, H. J., Effect of cyanide on the L-amino acid oxidase reaction, Biochem. Biophys. Res. Comm., 46, 564, 1972. 169. Cristman, M. F. and Cardenas, J. M., Essential arginine residues occur in or near the catalytic site of L-amino acid oxidase, Experientia, 38, 537, 1982. 170. Thome-Beau, F., Le-Thi-Lan, Olomucki, A., and Van Thoai, N., Essential histidyl residues in arginine oxygenase (decarboxylating). Comparison with amino acid oxidases, Eur. J. Biochem., 19, 270, 1971. 171. Marcotte, P. and Walsh, C., Vinylglycine and propargylglycine: complementary suicide substrates for Lamino acid oxidase and D-amino acid oxidase, Biochemistry, 15, 3070, 1976. 172. Pistorius, E. K., Kertsch, R., and Faby, S., Investigation about various possible functions of the L-amino acid oxidase in the cyanobacterium Anacystis nidulans, Z. Naturforsch., 44c, 370, 1989.

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Chapter 4

METHANOL OXIDASE F. Miiller, T. R. Hopkins, J. Lee, and P. I. H. Bastiaens

TABLE OF CONTENTS I.

Introduction

96

II.

General Properties and Sources of Methanol Oxidase A. General Background Information B. Sources and Purification of Methanol Oxidase C. Crystals and Crystalloids of Methanol Oxidase D. Spectral Properties of Methanol Oxidase and Its Azide Complex E. The Interaction Between Enzyme and Formaldehyde and H2O2 F. The Flavosemiquinone of Methanol Oxidase G. Apoprotein of Methanol Oxidase H. Chemical Modification of Methanol Oxidase 1. Modified FAD in Methanol Oxidase 2. Chemical Modification of the Protein

96 96 98 99 100 102 103 104 104 104 107

III.

Catalytic Properties of Methanol Oxidase

107

IV.

Molecular Biology of Methanol Oxidase

109

V.

Fluoresence Properties of Methanol Oxidase

109

VI.

Outlook

115

Acknowledgments

115

References

117

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Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION Enzymes capable of metabolizing one-carbon compounds are widely spread in nature.1 The cofactors of the enzymes involved in the conversion of methanol into formaldehyde differ. In prokaryotic organisms the main cofactor is pyrrolo quinoline quinone, flavin is seldom observed and if it is involved it is associated with metal ions.2 In eukaryotic organisms, on the other hand, the "simple" flavoprotein methanol oxidase plays an important role in the metabolism of methanol.3 Methanol oxidase (EC. 1.1.3.13) is called a "simple" flavoprotein because it appears to contain FAD as sole coenzyme like e.g., D- and L-amino acide oxidase (see Curti et al., Chapter 3, this volume). However, as will be shown below, the biochemical and biophysical properties of the enzyme seem to be more complex than anticipated. Methanol oxidase, often also referred to as alcohol oxidase, is localized in peroxisomes of methylotrophic yeasts (Scheme 1). Up to date four genera of methylotrophic yeasts have been identified: Hansenula, Pichia, Candida, and Torulopsis.4 The oxidase, generally a flavoprotein, in all peroxisomes in eukaryotic cells is always associated with catalase to protect the cell from the noxious action of H2O2*5 Catalase is involved in both the catalatic (dismutation of H2O2) and the peroxidatic (oxidation of methanol by H2O2) reactions. Formaldehyde generated in the peroxisome is dissimilated by cytosolic enzymes to give CO2 and energy as shown in Scheme 1. The assimilative pathway, not shown in Scheme 1, yields cellular biomass. The latter pathway requires the transformation of formaldehyde into dihydroxyacetone. This reaction is catalyzed by dihydroxyacetone synthase, also localized in the peroxisome.4 As a matter of fact the compartmentation of the three key enzymes in methanol metabolism in peroxisomes is a common feature of all yeasts grown on methanol. The function of peroxisomes easily adapts to the changing physiological environment in eukaryotic cells. The degree of response varies widely and present knowledge indicates that the susceptibility of eukaryotic cells for peroxisome induction decreases with the evolutionary development of the organism.3'5 The process of peroxisome proliferation seems on the other hand more easily reversible in lower than in higher organisms.3'5 Yeast cells grown on glucose contain a low number of small peroxisomes. When these cells are transferred to methanol-containing media the size and number of peroxisomes increases dramatically. Only such cells contain a large amount of methanol oxidase, the yield of methanol oxidase can be as high as —40% of the total soluble cellular protein.3 The peroxisomes present in these cells quickly disappear when the cells are transferred to glucose-containing media, due to active degradation.6'7 The "history" of methanol oxidase is unusual from a biochemical point of view and in the context of the development of knowledge in the field of flavoproteins. In 1969 Ogata et al.8 reported the first isolation of a methanol-utilizing yeast. Thereafter the isolation and purification procedures for methanol oxidase were worked out. While the regulation of methanol oxidase in yeasts is fairly well understood and the enzyme has even been cloned and its nucleotide sequence elucidated,9 the biochemical mechanism and physical and chemical properties of the enzyme are still badly understood. Therefore, besides summarizing published data, the emphasis of this chapter will be on some biophysical results obtained recently. It is hoped that these data will stimulate further research on methanol oxidase and contribute to a better understanding of the biochemical properties of the enzyme.

IL GENERAL PROPERTIES AND SOURCES OF METHANOL OXIDASE A. GENERAL BACKGROUND INFORMATION Methanol oxidase is synthesized on free ribosomes and transported into peroxisomes post-translationally. The protein is made in its mature form. As with many other microbody

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SCHEME 1. Schematic representation of the peroxisome and the metabolic and assimilative pathways of methanol. Abbreviations: MOx, methanol oxidase; CAT, cataJase; FDh, formaldehyde dehydrogenase; SFGDh, S-formylglutathion hydrolase; FODh, formate dehydrogenase.

proteins the sequence of methanol oxidase from Hansenula does not contain a microbody uptake signal, at least no signal was detected up to date. Therefore the mechanism of import of methanol oxidase by peroxisome remains obscure and subject to further investigations. Nevertheless it has been speculated that if there is no leading sequence the membrane of the peroxisome might be permeable for which some evidence seemed to exist. However in vivo 31P NMR studies on Hansenula, Candida, and Trichosporon have shown that peroxisomal membrane is impermeable.10 In addition these studies also demonstrated that the pH of the peroxisome is acidic, i.e., 5.8 to 6.O.10 The 31P NMR signal of inorganic phosphate correlate very well with the peroxisomal changes providing thus solid support for the interpretation of the data.10 The control of peroxisome proliferation in Hansenula is complex and cannot be induced by merely overproducing a single peroxisomal protein. The overproduction leads to swelling of preexisting peroxisomes but not to peroxisome proliferation. The number and shape of peroxisomes, which can differ widely, depend on the metabolic state of the cell. Veenhuis et al.11 and Van der Klei et al.12 found that mature peroxisomes do not respond to changing physiological conditions and that methanol oxidase induced by transfer of glucose-grown cells into methanol-containing media enters only proliferating peroxisomes. Hansen et al.13 have shown that methanol oxidase from Hansenula polymorpha does not contain a sequence, as found in other eukaryotic cells, which would serve as a target signal for the protein import into the peroxisomes. However, they showed that by fusion of a carboxy terminal sequence of dihydroxyacetone to bacterial (3-lactamase transport of the heterologous protein into peroxisomes is mediated. This observation was interpreted that the carboxy-terminal of dihydroxyacetone contains a target signal which needs to be further characterized. It remains to be seen if this observation is also a possibility for methanol oxidase. Structural methanol oxidase mutants of Hansenula polymorpha capable of overproduction of the fully active enzyme upon transformation were described by Rogenkamp et al.14

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Chemistry and Biochemistry of Flavoenzymes

The quantities of methanol oxidase produced were as high as about two-thirds of the total cellular protein. The overproduced protein was imported into the peroxisomes. The morphology and stability in cell lysates and the size of peroxisomes were strongly altered as compared with nontransformed yeast cells. The organelles showed a tendency to form rectangular bodies and their lumina were completely filled with the crystalloid structure (see below). The regulation of methanol oxidase in Hansenula polymorpha was studied in continuous cultures using a mixture of glucose and methanol.15 The levels of methanol oxidase mRNA, methanol oxidase in monomeric and octameric form, the FAD/methanol oxidase ratio, and the activity of the enzyme were quantified as a function of the dilution rate D. The study showed that (1) an induction of methanol oxidase mRNA formation occurs up to D = 0.29 h^ 1 , (2) the induction of methanol oxidase synthesis is determined at low D values on the transcriptional level, (3) the methanol oxidase activity at high D values is controlled by FAD incorporation and posttranslation, (4) the yield of protein synthesis was up to 37% of the cellular protein, (5) despite of high levels of methanol oxidase mRNA, decreasing levels, of methanol oxidase activity and protein were found at D values between 0.14 and 0.29 h" 1 , (6) the maximal FAD/methanol oxidase ratio was found to be 6 at D = 0.1 h" 1 correlating with the maximal specific activity of methanol oxidase. The induction of methanol oxidase in yeasts leads to elevation of riboflavin synthase and riboflavin kinase activities.16-17 The induced formation of FAD synthase corresponds well with the synthesis of methanol oxidase in Candida boidinii™ and Hansenula polymorpha.19 Furthermore externally added riboflavin or FMN to cell cultures are transformed efficiently into FAD.20 As already mentioned above the compartmentation of methanol oxidase, catalase and dihydroxyacetone synthase in peroxisomes is a prerequisite for the assimilation and dissimilation of methanol. This has been proven by Hansen and Roggenkamp21 who showed that a mutant of Hansenula polymorpha defective in catalase was not able to grow on methanol as the sole carbon source; growth on glucose or glycerol was not impaired. Transformed mutant cells containing the gene coding for catalase A from Saccharomyces cerevisiae were able to grow on methanol. Furthermore, when the gene was placed under control of the regulatory methanol oxidase promotor from Hansenula polymorpha, high levels of activity subject to glucose repression were obtained. These results seem to be at variance with data from Guiseppin et al.22 who showed that a catalase-negative mutant of Hansenula polymorpha was able to grow on a mixture of glucose and methanol and that methanol oxidase was induced up to a level of 40% of that obtained in the wild-type strain. B. SOURCES AND PURIFICATION OF METHANOL OXIDASE Methanol oxidase has been isolated from various fungi (Table 1). The molecular mass of the protein seems to vary considerably, depending on the strain. However, present knowledge indicates that the subunit molecular mass of all proteins is 70 to 80 kDa and that the majority consists of eight subunits, each subunit containing 1 mol FAD. Methanol oxidase can be easily purified in a few steps.24'26'37-38 The purification can be achieved in 3 to 4 steps: preparation of cell-free extract, ammonium sulfate precipitation, and gel filtration. The conditions of cell growth and method used to prepare the cell-free extract can be crucial factors for the yield of methanol oxidase.3 This is related to the stability of the cell wall depending on the growth conditions of the cells.3 The strength of the cell wall of Hansenula polymorpha has been investigated in dependence of the composition of the culture media and a medium containing methanol as the sole carbon source.39 The cell wall strength was influenced by growth media and dilution rate D. The slowest lysis kinetics and a large fraction of nondegradable cell wall material was observed when the culture medium contained

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TABLE 1 Methanol Oxidase from Various Fungi and Its Molecular Mass and Subunit Composition Source Molds Polyporus sp. Poria contigua Yeasts Kloeckera sp. 2201 Hansenula polymorpha Candida boidinii Candida N16 Candida 25A Pichia pastoris Pichia sp. Phanerochaete chrysosporium

Molecular mass (kDa)

Number of subunits

300 610

4 8

23, 24 25

8 8 8 4; 8 8 8 4 4

26, 27, 28 3,28 29, 30 31, 32 33 34 35 36

570—673 616—669 600 210, 600 520 675 300 310

Ref.

a mixture of glyceroi and methanol. When a mixture of glucose and methanol was used in continuous cultures both the resistance to zymolyase and the cell wall thickness increased at D < 0.1 h" 1 . However, the yield of methanol oxidase by zymolyase lysis is up to 100% higher than that of ultrasonic treatment. At D values > 0.2 h ~ ' the rate of methanol oxidase release increases owing to the increased fraction of thin daughter cells. Therefore the kinetic analysis of zymolyase lysis of the cultured cells combined with physical and enzymatic methods provides a method for the achievement of optimal recovery of methanol oxidase. For the preparation of large amounts of methanol oxidase a few simple procedures have been developed.40 In these procedures the cells were disrupted by bead mill, high pressure shear and nonmechanical means such as solvent lysis using either diethyl ether, chloroform or methylene chloride. The cells were lysed at pH 7 to obtain the homogenate. To increase the stability of the free enzyme the lysis is preferentially done in the presence of azide forming a complex with the enzyme (see below). The cell free extract was obtained by centrifugation, microfiltration or just sedimentation (natural settling). The pH of the cellfree supernatant was then adjusted to 6.3 and the ionic strength brought to a conductivity of 2400, yielding methanol oxidase in crystalline state. Washing the crystals thus obtained by buffer and water yielded a highly purified preparation (95%) in high yields (80%). C. CRYSTALS AND CRYSTALLOIDS OF METHANOL OXIDASE Methanol oxidase appears in the form of crystalloids in the peroxisomes.3-14 The size and number of the crystalloids depend on the growth conditions of the cells. Cells from the exponential growth phase of batch cultures contain a partly crystalline matrix whereas an increase in size of the crystalloids is observed during vegetative reproduction.3 A largely, or completely, crystalline matrix was observed in cells grown in both cultures entering the stationary phase of growth. The peroxisomes present in cells of methanol-limited chemostat cultures invariably showed a completely crystalline structure. The crystallization of methanol oxidase was first achieved as part of the purification procedure described by Janssen and Ruelius24 using polyethylene glycol 6000. Later Tani et al.26 crystallized the enzyme by ammonium sulfate fractionation and also obtained fine thin needles exhibiting an yellow color (Kloeckera, sp.). Methanol oxidase is highly soluble in water (at least 100 mg/ml) the solubility is markedly dependent on the ionic strength of the solution.27 The solubility of methanol oxidase is minimal at an ionic strength of 0.04 to 0.05 M and is about 5 mg/ml at 40°C. Reduced methanol oxidase exhibits an even lower solubility. Hopkins279 made use of this solubility property to crystallize methanol oxidase

100

Chemistry and Biochemistry of Flavoenzymes

FIGURE 1. Light absorption spectra of methanol oxidase from Pichia pastoris in phosphate buffer, pH 7.0 (enzyme concentration with respect to FAD was 0.7 mM). The dashed line represents the enzyme in the "yellow" form and the full line is the spectrum of the azide-complexed enzyme. An estimated difference spectrum between the two spectra is also shown (dotted curves).

in large quantities, as already mentioned above, i.e., the cell free extract was adjusted to pH 6.3 and an ionic strength of 0.04 to 0.05 M. More recently the enzyme from Pichia pastoris was also crystallized and investigated by X-ray techniques.41 The results show that the asymmetric unit of the crystals contains two octamers with a molecular mass of over l,200kDa. Obviously it will be very difficult to solve the structure by conventional techniques. In order to advance and to aid in the resolution of the structure of methanol oxidase Vonk and van Bruggen41 approached the problem by the preparation of two-dimensional crystals from Hansenula polymorpha which were studied by electron microscopy. The crystals exhibit p4 symmetry. The dimension of a unit cell is 12.5 x 12.5 nm2 and contains one enzyme molecule. The data demonstrate that the subunits of methanol oxidase possess an elongated shape and form two layers of four stacked face to face.42 D. SPECTRAL PROPERTIES OF METHANOL OXIDASE AND ITS AZIDE COMPLEX Purified free enzyme exhibits a yellow color as expected for a flavoprotein not containing metal ions or any other chromophoric groups.34 Despite this the visible light absorption spectrum of free methanol oxidase is composed of different species. The absorption spectrum of free methanol oxidase is shown in Figure 1. Addition of azide leads to considerable spectral changes. The appearance of absorbance at longer wavelengths is typical for flavoproteins forming molecular complexes with organic molecules.43 The spectrum of free methanol oxidase resembles but is not identical with spectra of other flavoprotein oxidases, e.g., D- and L- amino acid oxidase (see Curti et al., Chapter 3, this volume). Moreover the spectral shape of free methanol oxidase can vary considerably among

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preparations and source of isolation. In Figure 1 the difference of absorption between the spectra of free and azide-complexed enzyme is also indicated as obtained by calculation. The absorption difference indicates possible maxima at about 500 and 340 nm for the azide complex. The strength of interaction between methanol oxidase and azide differs greatly.37 This observation seems to be associated with the strain of yeast used as a source of the enzyme. For instance the formation of the azide complex with methanol oxidase from Hansenula polymorpha and Candida boidinii is easily reversed by dialysis whereas that of methanol oxidase from Pichia pastoris is much more difficult to reverse.37 The complex can also be dissociated into its constituents by either lowering the pH of the solution between 5.5 to 6.0, raising the temperature of the solution to 50°C or reduction of the enzyme to the FADH2 form in the presence of substrate and in the absence of oxygen.44 Hopkins44 developed an elegant method for the preparation of azide-free, yellow methanol oxidase. About 5 ml of crystalline, red methanol oxidase slurry in 0.05 M phosphate buffer containing 0.02% sodium azide was centrifuged for 10 min at 3000 rpm (20°C). The pellet was washed two times with 10 ml 0.05 M phosphate buffer (pH 7.5) at room temperature. The slurry was then resuspended in 10 mml of the same buffer in a 15 ml screw-cap test tube and made anaerobic by flushing with nitrogen. To the anaerobic solution 2 jml of pure ethanol was added, mixed and the anaerobic solution kept 20 min at room temperature. During this time the color of the slurry changes from red to light pink-yellow typical of the reduced enzyme. The reduced, crystalline slurry was then harvested by centrifugation and washed one time with 10 ml deaerated buffer containing 1 JJL! ethanol. The pellet was drained carefully in the air and redissolved in water to give a straw-yellow, fully active solution of concentrated methanol oxidase. This work also revealed that reduced methanol oxidase crystals are less dense than oxidized crystals of the azide complex suggesting that methanol oxidase undergoes a conformational change upon binding of azide. In addition, it was also noticed that reduced methanol oxidase per se is not unstable since it can be stored for months under anaerobic conditions without loss of activity. Therefore the instability of methanol oxidase observed under certain conditions must probably be ascribed to H2O2 formed during turn-over of the enzyme. The association constant for the azide complex formation for the oxidized enzyme was determined to be 6 x 106M"], with 0.97 binding sites per enzyme subunit (7.7 sites per octameric enzyme).37 The inhibition of the enzyme from Pichia pastoris by azide was found to be of competitive nature, but of linear mixed type inhibition (methanol as substrate).37 The Ki value was determined to be 6 x 10~6M, in fair agreement with the association constant. On the other hand the inhibition of the enzyme from Candida boidinii S2 was reported to be competitive with respect to methanol, Kj = 1.5 x 10~5M.38 In both cases the enzymic activity was not altered by the presence of azide. This observation is in agreement with the type of inhibition observed with the Pichia pastor is methanol oxidase, but is difficult to reconcile with the competitive inhibition of the Candida boidinii enzyme. Assuming that the KM value of methanol (2.6 mM at saturating O2 concentrations37) represents also an approximate dissociation constant of the enzyme-methanol complex, then the much higher affinity of the enzyme for azide than for methanol would indeed strongly inhibit the enzyme in the presence of up to 0.16 mM azide, as reported by Sakai and Tani,38 but is in contradiction with their statement regarding the enzymic activity in the presence and absence of azide. The results published by Sakai and Tani38 show that incubation of methanol oxidase by 0.25 or 2.5 mM azide for 1 to 2 d at 4°C did not affect at all the enzymic activity. Incubation at 30°C only affected the enzymic activity in the presence of 0.25 mM azide, but not in the presence of 2.5 mM azide. Since the experimental procedure was poorly described it is guessed that a stock solution of the enzyme-azide complex was prepared which was diluted many fold for activity measurement diminishing the inhibition by azide. The results obtained

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at 30°C, however, demonstrate clearly the beneficial effect of azide on the stability of the enzyme. E. THE INTERACTION BETWEEN ENZYME AND FORMALDEHYDE, AND H202 There are some indications that formaldehyde37-38 and probably also H2O2, are strongly interacting with the enzyme leading to inactivation of the protein. The degree and ease of inactivation seems to depend on the source of isolation of the enzyme. 27 Nevertheless, while such an interaction could easily inactivate the enzyme by covalent adduct formation between e.g., lysine residues of the protein and formaldehyde, the inactivation by H2O2 could occur by oxidation of certain amino acid residues (e.g., sulfhydryl groups). Recently it has been shown by Sakai and Tani38 that methanol oxidase from Candida boidinii 2S can be protected by catalase from inactivation by H2O2. Interestingly the enzyme-azide complex was much less inactivated by H2O2 than the free enzyme suggesting that azide protects a site in the enzyme from reaction with H2O2. This site could be a sulfhydryl or lysine group forming a noncovalent adduct with H2O2. It must however be mentioned that the nature of the interaction between the free enzyme and H2O2 has not yet been clarified (covalent, noncovalent interaction, degree of reversibility). Formaldehyde, the product of the enzymic oxidation reaction, also interacts with the enzyme in an unusual manner for an enzymic reaction. Highly purified enzyme contains varying amounts of formaldehyde adduct, the amount of adduct depends on the history of the preparation used.37 The adduct is stable enough to survive the purification procedure, but the adduct formation is still reversible. The likely form of the adduct is a Schiff base formation with lysine residues of the protein. Purified methanol oxidase from Pichia pastor is can contain as much as 25% of all available e-amino groups of lysine residues in the form of Schiff base.37 It has been proposed that the Schiff base formation could serve as a formaldehyde buffering system which could explain the apparent overproduction of methanol oxidase in cells grown on methanol. The reduced glutathione-formaldehyde complex is the substrate of formaldehyde dehydrogenase (Scheme 1). Under physiological conditions where glutathione concentrations are not high enough to handle a transient surge in formaldehyde concentration, formaldehyde could be temporarily stored as adduct in methanol oxidase in the peroxisome. The Schiff base formation could also influence the solubility of the protein (facilitation of crystalloid formation) or may mask sites of proteolytic attack from trypsinlike degradative proteases and could thus constitute a possible regulatory mechanism of peroxisome proliferation.37 In Candida boidinii methanol oxidase the Schiff base formation by formaldehyde has also been observed recently.38 The Schiff base content is higher in Candida boidinii than in Pichia pastoris methanol oxidase. The formation of Schiff base in methanol oxidase could also explain an observation by Geissler and Hemmerich.45 These authors found that the flavin spectrum of a methanol oxidase solution becomes bleached upon removal of oxygen, indicating reduction of enzymebound FAD. Since no substrate was present the authors postulated that the enzyme contains bound H2O2 which reacts according to the following equation: Fl-ox + H2O2±q»HFl-red + O2 + H + i.e., removal of O2 would shift the equilibrium to the right and that the reaction is reversible. While the reaction to the left hand side of the equation is trivial for all flavoprotein oxidases it would be very surprising, even despite the fact that the reaction is in principle microscopically reversible, to drive the reaction fully to the right by removal of oxygen. A more reasonable explanation would be that free formaldehyde (i.e., hydrated formaldehyde), a substrate of methanol oxidase and in equilibrium with the Schiff base, is reducing the enzyme.

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This reduction can only be observed under anaerobic conditions: Flox + CH2O • H2O «± HFlred + HCOOH + H + A similar mechanism, i.e., storage of aldehyde in some form in the enzyme, could also hold true for luciferase (see Lee et al., Chapter 6, Vol. II of this series) for which it was proposed by Matheson et al.46 that the bioluminescence reaction could also occur in the absence of the substrate (aldehyde). F. THE FLAVOSEMIQUINONE OF METHANOL OXIDASE Another unusual property of methanol oxidase as a flavoprotein oxidase is the formation of an air-stable anionic flavosemiquinone.47"49 Methanol oxidase isolated from various yeast strains contains varying amounts of flavosemiquinone. The flavosemiquinone form of the enzyme is catalytically inactive, as expected for a flavoprotein oxidase, and can amount between 40 and 70% of the flavin present in the octameric enzyme molecule.45'50 The radical can be easily observed by visible absorption spectrophotometry. The small shoulder at about 400 nm, the intense band at about 370 nm and an increased absorption at about 470 to 500 nm (see Figure 1) are diagnostic for the anionic flavosemiquinone.47 The presence of flavosemiquinone in methanol oxidase is one of the factors mentioned above to contribute to the complexity of the visible absorption spectrum. Therefore it is very difficult to calculate the content of oxidized enzyme in a given preparation from visible spectra. The content of oxidized flavin in methanol oxidase can roughly be estimated by methanol-reduction of the enzyme under anaerobic conditions and substrating the spectrum thus obtained from the original spectrum. Another possibility is to convert the oxidized form of the protein into the semiquinone form by hydroxylamine.45 In contrast to electron paramagnetic resonance spectroscopy (ESR) electron nuclear double resonance (ENDOR) spectroscopy yields more detailed information on a proteinbound radical owing to a much better resolution of the spectra. The ENDOR spectrum of methanol oxidase from Candida boidinii in the semiquinone form contains two distinguishable flavin radical species, both being of the anionic type of radical.51 Interestingly one set of lines belongs to the semiquinone already present in the freshly prepared enzyme. Reduction of such a sample by hydroxylamine yields a second set of lines indicating strongly that the two flavin semiquinones differ in some chemical property or are placed in different environments in the enzyme. The two radicals also differ with respect to their reactivity towards oxygen. The radical produced by hydroxylamine reduction is easily oxidized by O2 whereas the original radical is air-stable. The latter resembles the characteristics of the complex between the anionic flavosemiquinone of lactate oxidase and the product pyruvate.52 (see also Ghisla and Massey, Chapter 8, Vol. II of this series). This complex is rather stable towards dioxygen, but can still be slowly oxidized by O2. The similarity between the two cases, differences are that formaldehyde is much more tightly or even covalently bound to methanol oxidase than pyruvate to lactate oxidase, indicates that similar factors are operating. Mincey et al.50 have studied the accessibility of the flavosemiquinone in methanol oxidase to water by the spin echo technique. It was found that the flavosemiquinone is accessible to water and that the accessibility is significantly decreased in the presence of the inhibitor cyclopropanol. These data then suggest that the inhibitor was able to bind to the active site of the protein. Consequently one could conclude that formaldehyde interacts with the protein at other sites than the active site. In order to influence the oxygen-stability of the flavosemiquinone, binding of formaldehyde must influence either the three-dimensional structure of the active site of methanol oxidase or block access of O2 by for instance steric hindrance. The flavosemiquinone in methanol oxidase is clearly a "physiological" artifact. Therefore the elucidation of the mechanism of formation should be studied leading eventually to

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its suppression and subsequently to the preparation of a more uniform methanol oxidase preparation. The unusual high air-stability of the anionic flavosemiquinone might be related to the interaction of formaldehyde with the two-electron reduced methanol oxidase altering the environment of the active center and/or the structure of methanol oxidase such that an one-electron oxidation is favored. At any rate the finding of Blankenhorn et al.53 that reduced flavin forms an adduct with formaldehyde might be an useful guide in studies on methanol oxidase. The formaldehyde adduct formation of reduced methanol oxidase could also account for part of the inactivation of the enzyme observed under prolonged turn-over conditions. G. APOPROTEIN OF METHANOL OXIDASE A few attempts have been undertaken to resolve methanol oxidase into its constituents apoflavoprotein and FAD, and to reconstitute the native protein. Resolution of the enzyme in a mixture of phosphate buffer (pH = 7.5) containing 6 M urea and 3.5 M KBr and subsequent reconstitution of the protein by column chromatography was achieved.50 The reconstituted protein exhibits a two-banded visible spectrum but the spectrum still deviates from an usual flavoprotein spectrum by an increased intensity of the second band at about 350 nm. The enzymatic activity of such preparations varied between 37 and 58% that of the original enzyme. Considering the already low enzymatic activity of the original enzyme preparation it is obvious that only qualitative data can be obtained from such reconstituted enzyme samples. The protein was also reconstituted with 5-deazaflavin.54 This protein did neither catalyze the oxidation of methanol nor the reduction of formaldehyde by the reduced form of the protein. In view of the great complexity of the chemical properties of methanol oxidase it is most desirable to prepare highly active reconstituted enzyme. If this goal could be achieved, which may involved a major effort, then the biochemical properties of the enzyme could be studied in much more detail than was possible up to date. H. CHEMICAL MODIFICATION OF METHANOL OXIDASE 1. Modified FAD in Methanol Oxidase Owing to the complex mixture of methanol oxidase molecules in isolated preparations there has been some speculations about the chemical nature of the mixture. While it is well recognized that the flavosemiquinone form of methanol oxidase contributes considerably to the mixture in enzyme preparations it was not clear why the residual oxidized flavin in methanol oxidase could not be accounted for in light absorption spectra by calculation using the molar extinction coefficient of FAD.45 Geissler et al.55 have observed that only part of the oxidized methanol oxidase from Candida boidinii was catalytically competent. The catalytically noncompetent flavin accounted for 30 to 35% of the total flavin in the particular preparation used. Geissler et al.55 also found that the oxidized flavin in catalytically noncompetent methanol oxidase exhibits a considerable absorbance in the 370 nm and 460 nm region. They proposed that FAD is bound to a modified active site being not reducible by the substrate. Previous to the studies by Geissler et al.,55 Sherry and Abeles54 isolated two chemically different FADs from methanol oxidase by denaturation and HPLC analyses. From the two FAD fractions thus obtained the second fraction showed identical physical and chemical properties as authentic FAD (retention time of 16.2 min). The other (first) fraction (retention time of 13.8 min) differed from the properties of authentic FAD. Methanol oxidase from Hansenula polymorpha contained 49% FAD and 51% modified FAD and that from Pichia pestoris contained 13% FAD and 87% modified FAD. This variation is probably not related to strain specificity but has its origin in some other factors (see below). Sherry and Aheles54 reported that the light absorption spectra of the two flavins are identical, except that the extinction coefficient of modified FAD at 370 nm is slightly larger relative to that at 445 nm. These data suggest that the unidentified prosthetic group observed

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SCHEME 2. The chemical structures of riboflavin, its periodic acid product, the cyclic acetal, hemiacetal, and xylase.

by Geissler et al.55 is identical with the modified FAD isolated by Sherry and Abeles.54 Consequently the catalytically noncompetent methanol oxidase should contain modified FAD as prosthetic group. It was further shown by Sherry and Abeles54 by 1H NMR that the isoalloxazine ring structure in both FADs was identical. Moreover the structure of the AMP moieties of both flavins, obtained by phospodiesterase treatment of both flavins, were also identical as judged by thin layer chromatography and HPLC. Hence it was concluded that the chemical difference between the two flavins resides in the N(10)-ribidyl side chain. This follows from the facts that both ribityl side chains contain the same number of nonexchangeable protons, determined by 'H NMR, but their chemical shifts are distinctly different. Further proof for the modification of the side chain came from the periodic acid treatment of the two compounds yielding identical products.54 The treatment of modified riboflavin or FMN with periodic acid yields 7,8-dimethyl-10-formylmethylisoalloxazine (Scheme 2).56'57 This suggests that the chemical modification in the structure of the modified FAD in methanol oxidase must be associated with either C(2'), C(3') or C(4') of the ribityl side chain. Based on their data Sherry and Abeles54 proposed that the modified FAD in methanol oxidase is an optical isomer of FAD. If this suggestion is correct then a chemical reaction should occur at one or more of the above mentioned three carbon atoms in order to yield the optical isomer. Bystrykh et al.58 investigated recently the relation between the growth condition, Vmax and K M , on the one hand, and the degree of modification of FAD in methanol oxidase from Hansenula polymorpha, on the other hand. The results confirm the findings of Sherry and Abeles54 and show in addition that the content of modified FAD in methanol oxidase depends apparently on the growth conditions of the yeast. Cells grown under batch conditions yield less modified FAD than under chemostat conditions, but the amount of modified FAD formed was dependent on the dilution rate D, i.e., at low dilution rates more modified FAD is formed than at high Ds. The situation seems to be even more complex. When Bystrykh et al.58 isolated the enzyme at pH 5.8 rather than as usually at pH 7.5 they obtained an almost homogeneous enzyme preparation with respect to the content of modified FAD. The authors also conclude that there is a relation between the enzymatic activity and the content of modified FAD in the enzyme preparations and consequently suggest that modified FAD

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and not FAD is the natural coenzyme of methanol oxidase. In our opinion this is an overstatement and is not supported by the data. For instance according to Table 1 in the paper by Bystrykh et al.58 very little modified FAD is present in cells grown at D = 0.18 h" 1 , at D = 0.05 h ~ f the majority of FAD is present in the modified form. Yet the V max and KM values of the two purified enzyme preparations do not vary drastically (Table V in the paper of Bystrykh et al.58). The light absorption spectrum of purified modified FAD is, as already reported by Sherry and Abeles,54 very similar but not identical with that of FAD, the spectrum seems to possess larger extinction coefficients at 376 nm and 448 nm than FAD at 374 nm and 445 nm.58 The circular dichroism spectra of FAD and modified FAD show opposite cotton effects in the visible region.58 These data are in support of the proposal by Sherry and Abeles54 and confirm modification of FAD in the side chain. Bystrykh et al.58 did the following experiment to shine some light on the mechanism of protein and FAD modification of methanol oxidase by formaldehyde. To an anaerobic solution of methanol oxidase containing only modified FAD 14CH3OH was added followed by NaBH4. It should be noted that under these conditions only the methanol-oxidizing (flavinreducing) half-reaction of the catalytic cycle can occur. Analysis of the sample indicated that 0.8 mol label per mol subunit methanol oxidase was incorporated into the enzyme, also suggesting that the enzyme had an exceptional activity of 80% of that theoretically possible. Further analysis showed that 74% of the label was incorporated into the apoprotein and 26% into the modified FAD.58 These are remarkable results considering the quantitative incorporation of the labeled product into the protein. Also surprising is the fact that modified FAD is also labeled and its HPLC pattern still identical with that of unlabeled modified FAD. The latter result can only be explained if the original modification is reversible, the formation of a cyclic acetal between formaldehyde and the hydroxyl groups for instance at C(3') and C(4') of the ribityl side chain moiety of FAD (Scheme 2). This reaction would require acid catalysis. Furthermore the acetal shown in Scheme 2 could not react with periodic acid but the C(2')-C(3') cyclic acetal would do so. The *H NMR spectra of these acetals should contain two protons more than e.g., FMN. This is in disagreement with the data of Scherry and Abeles54 except if the methylene protons of the cyclic acetals are easily exchangeable which is feasible. On the other hand the 13C NMR spectrum of the acetal would contain one carbon atom more. From the above discussion it can be concluded that the cyclic acetal is possibly not a candidate for the structure of modified FAD in methanol oxidase. In contrast the formation of a hemiacetal seems more likely and has also been observed in model reactions using riboflavin and formaldehyde (Scheme 2).59 The hemiacetal reacts with periodic acid, the hemiacetal formation is reversible and the product is quite stable towards hydrolysis exhibiting a minimum stability at about pH 7.59 The light absorption properties of the hemiacetal resemble those of riboflavin. Interestingly the hemiacetal is stronglydextrorotatory whereas the starting material riboflavin is levorotatory.59 Consequently all presently available data (light absorption, 1H NMR and CD) can be consolidated by the formation of xylose (Scheme 2) or arabinose if C(2') is involved in the hemiacetal formation. Therefore it is proposed that modified FAD in methanol oxidase is either a derivative of xylose or arabinose. The proposed mechanisms require the reversible dissociation of the prosthetic group from the protein which can occur. At any rate it seems obvious that the enzymatic oxidation product formaldehyde is involved in the modification reaction of FAD bound to methanol oxidase. Consequently FAD is the natural cofactor of methanol oxidase rather than modified FAD, as suggested by Bystrykh et al.58 Since modification of FAD occurs only under certain physiological conditions of the cell and the modification represents a merely secondary chemical reaction it is difficult to imagine the biological relevance of such a modification. This interpretation is in line with the present knowledge that the redox potential of flavins is only drastically altered by chemical modification of the isoalloxazine

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nucleus (see Stankovich, Chapter 18, Vol. I of this series). In this context it is tempting to speculate a little further on the mechanism of formation of modified FAD-containing methanol oxidase. It is proposed that the chemical modification of FAD in the cell occurs in free FAD which, depending on the dissociation constant of the enzyme-FAD complex, is present in a finite concentration according to the equilibrium: E-FAD 450 nm. The spectrum is independent of wavelengths which was checked at the additional (emission) wavelengths at 540 nm and 560 nm. These data seem to indicate that methanol oxidase does not only contain conventional flavin but also other chromophore(s). Remarkably, the two maxima at 322 nm and 418 nm in the excitation spectrum correspond with the minima in the absorption spectrum of flavin. The methanol oxidase-azide complex (for the visible absorption spectrum see Figure 1) yields the emission spectrum presented in Figure 3A. The spectrum (excitation at 480 nm) is almost identical with that of the yellow form of the enzyme. The quantum yield of the yellow form is however higher than that of the azide complex. This can be expected on the basis of the close association of azide with the protein in the neighborhood of the flavin as

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FIGURE 3. Corrected luminescence spectra of the "red" form (azide-complexed) of methanol oxidase in phosphate buffer, pH 7.0. (A) Emission spectrum, excitation wavelength was 480 nm. (B) Excitation spectrum, emission wavelength was 580 nm.

evidenced by the visible light absorption spectrum. The excitation spectrum (emission = 540 nm) exhibits two bands with maxima at 335 nm and 475 nm (Figure 3B). Three marked shoulders are also seen at 510 nm, 455 nm, and about 420 nm. The spectrum differs dramatically from that of the yellow form (see Figure 2C). The excitation spectrum shows some resemblance with flavin at least in the long wavelength region.

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The enzyme has also been studied in the presence of methanoi under anaerobic conditions. Any enzyme molecules catalytically competent should be reduced by methanoi and the fluorescence spectrum should therefore reveal the catalytically noncompetent, fluorescent molecules. Figure 4A shows the excitation spectrum (emission = 580 nm) of an oxidized methanoi oxidase solution. The spectrum is very similar with that shown in Figure 2C, although different batches were used in the two experiments. The solution used in Figure 4A was then made anaerobic and methanoi added. The solution thus obtained gave the excitation spectrum (emission = 580 mm) also shown in Figure 4A (lower curve). This experiment was repeated using another batch of enzyme. The resulting excitation spectrum (Figure 4B) is very similar, if not identical with that shown in Figure 4A (lower curve). The excitation spectrum of the catalytically noncompetent fraction of the enzyme shows some similarity with oxidized flavin but the first absorption maximum is shifted to the red by about 30 nm as compared with that of conventional flavin. The emission spectrum of this solution (excitation at 340 nm, results not shown) showed emission bands with decreased intensities as compared to those of oxidized enzyme. It is not easy to explain the complex luminescence spectra but it seems reasonable to conclude the following. The free enzyme used in our studies contains three fluorescent species: the azide complex which exhibits an emission maximum at about 580 nm, catalytically competent enzyme and catalytically noncompetent enzyme. This conclusion is based on the comparison of the above spectra showing that catalytically reduced enzyme exhibits luminescence spectra very similar with those of the azide-complexed enzyme.

VI. OUTLOOK Methanoi oxidase is an interesting flavoprotein both with respect to the quarternary structure and the catalytic mechanism. The fact that high amounts of the enzyme can be easily obtained either by cloning or induction makes this protein a good candidate for a multidisciplinary research project. However, before such an approach can be effectuated several basic biochemical properties of the enzyme must be better understood and therefore further investigated. Properties to be studied with high priority are: (1) to develop growth conditions so that the cells produce a homogeneous enzyme; (2) to clarify the biological relevance of the modified FAD in methanoi oxidase and if not of relevance to find experimental conditions to prevent its formation; (3) to elucidate the mechanism of formation of the protein-bound flavosemiquinone; (4) to investigate the enzymatic mechanism. The above suggestions are also important in view of the fact that methanoi oxidase has been applied as a biosensor and for removal of aspartame from food.73-74 More recently it has been proposed to use the enzyme as an antimalarial agent in combination with ethanol75 and as a means to lower ethanol concentrations in blood (U.S. Patent filed by Phillips Petroleum Co.) These few examples illustrate that potentially the enzyme could lend itself for further applications if a well defined enzyme preparation can be achieved.

ACKNOWLEDGMENTS We thank Miss. S. Affolter for typing the manuscript, Mr. W. Gehrig for the preparation of the drawing, Mr. F. van Mieghem and Mr. I. Roscher for the assistance in some of the experiments. The work from the authors' laboratories was supported in part by The Netherlands Organization for Scientific Research (NWO/SON) (to FM) and in part by an NIH grant GM-28139 (to JL).

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FIGURE 4. Corrected excitation spectra of the "yellow" form of methanol oxidase in the oxidized and substrate-reduced state in phosphate buffer, pH 7.0. (A) Oxidized enzyme (full line) and methanol-reduced enzyme under anaerobic conditions (dotted line) (emission = 580 nm). (B) Substrate-reduced methanol oxidase in phosphate buffer, pH 7.0, under anaerobic conditions. The enzyme used in this experiment was from a different batch than that used in A. The emission wavelength was 580 nm.

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REFERENCES 1. Quayle, J. R., The metabolism of one-carbon compounds by micro-organism, Adv. Microbiol Physiol., 1, 119, 1972. 2. Duine, J. A., Frank, J., Izn, and Jongejan, J. A., Enzymology of quinoproteins, Adv. EnzymoL, 59, 169, 1986. 3. Veenhuis, M., Van Dijken, J. P., and Harder, W., The significance of peroxisomes in the metabolism of one-carbon compounds in yeasts, Adv. Microbiol. Physiol., 24, 1, 1983. 4. Wegner, G. H. and Harder, W., Methylotrophic yeasts -1986, in Microbiol Growth on Cl Compound, Van Verseveld, H. W. and Duine, J. A., Eds., Martinus Nijhoff Publishers, Dordrecht, 1987, 131. 5. Borst, P., Peroxisome biogenesis revisited, Biochim. Biphys, Acta, 1008, 1, 1989. 6. Bormann, C. and Sahm, H., Degradation of microbodies in relation to activities of alcohol oxidase and catalase in Candida boidinii, Arch. Microbiol., 117, 67, 1978. 7. Veenhuis, M., Zwart, K. B., and Harder, W., Degradation of peroxisomes after transfer of methanolgrown Hansenula polymorpha into glucose-containing media, Fed. Eur. Microbiol. Soc. Microbiol. Lett., 3, 21, 1978. 8. Ogata, K., Nishikawa, H., and Ohsugi, M., A yeast capable of utilizing methanol, Agric. Biol. Chem., 33, 1519, 1969. 9. Ledeboer, A. M., Edens, L., Maat, J., Visser, C., Bos, J. W., Verrips, C. T., Janowitz, J., Eckart, M., Roggenkamp, R., and Hollenberg, C. P., Molecular cloning and characterization of gene coding for methanol oxidase in Hansenula polymorpha, Nucl. Acids Res., 13, 3063, 1985. 10. Nicolay, K., Venenhuis, M., Douma, A. C., and Harder, W., A 31P NMR study of the internal pH of yeast peroxisomes, Arch. Microbiol., 147, 37, 187. 11. Veenhuis, M., Suiter, G., Van der Klei, I., and Harder, W., Evidence for functional heterogeneity among microbodies in yeast, Arch. Microbiol., 151, 105, 1989. 12. Van der Klei, L, Veenhuis, M., Nicolay, K., and Harer, W., In vivo inactivation of peroxisomal alcohol oxidase in Hansenula polymorpha by KCN in an irreversible process, Arch. Microbiol., 151, 26, 1989. 13. Hansen, H., Veenhuis, M., and Roggenkamp, R M Peroxisomal protein import in a methylotrophic yeast, J. Cell. Biochem. Suppl., 14, (Part C), 26, 1990. 14. Roggenkamp, R., Didion, T., and Kowallik, K. V., Formation of irregular giant peroxisomes by overproduction of the crystalloid core protein methanol oxidase in the methylotrophic yeast Hansenula polymorpha, Moi Cell Biol., 9, 988, 1989. 15. Giuseppin, M. L. F., Van Eyk, H. M. J., and Bes, B. C. M., Molecular regulation of methanol oxidase activity in continuous cultures of Hansenula polymorpha, Biotechnol. Bioeng., 32, 577, 1988. 16. Shimizu, S., Ishida, M., Tani, Y., and Ogata, K., Flavin changes of Kloeckera sp. No. 2201 during adaptation to methanol, Agric. Biol. Chem., 41, 423, 1977. 17. Shimizu, S., Ishida, M., Kato, N., Tani, Y., and Ogata, K., Derepression of FAD pyrophosphorylase and flavin changes during growth of Kloeckera sp. No. 2201 on methanol, Agric, Biol. Chem., 41, 2215, 1977. 18. Eggeling, L., Sahm, H., and Wagner, F., Induction of FMN adenylyltransferase in methanol utilizing yeast Candida boidinii, FEMS Microbiol. Lett., 1, 205, 1977. 19. Brooke, A. G., Dykhuizen, L., and Harder, W., Regulation of flavin biosynthesis in methylotrophic yeast Hansenula polymorpha, Arch. Microbiol., 145, 62, 1986. 20. Tani, Y., Kato, N., and Yamada, H., Utilization of methanol by yeasts, Adv. Appl. Microbiol. 24, 165, 1978. 21. Hansen, H. and Roggenkamp, R., Functional complementation of catalase-defective peroxisomes in a methylotrophic yeast by import of the catalase A from Saccharomyces cerevisiae, Eur. J. Biochem., 184, 173, 1989. 22. Guiseppin, M. L. F., Van Eyk, H. M. J., Bos, A., Verduin, C., and Van Dyken, P., Utilization of methanol by a catalase-negative mutant of Hansenula polymorpha, Appl. Microbiol. Biotechnol., 28, 286, 1988. 23. Janssen, F. W., Kerwin, R. M., and Ruelius, H. W., Alcohol oxidase, a novel enzyme from a basidiomycetes, Biochem, Biophys. Res. Commun., 20, 630, 1965. 24. Janssen, F. W. and Ruelius, H. W., Alcohol oxidase, a flavoprotein from several basidiomycetes species. Crystallization by fractional precipitation with polyethylene glycol, Biochim. Biophys. Acta, 151, 330, 1968. 25. Bringer, S., Sprey, B., and Sahm, H., Purification and properties of alcohol oxidase from Poria contigua, Eur. J. Biochem., 101, 563, 1979. 26. Tani, Y., Miya, T., Nishikawa, H., and Ogata, K., The microbial metabolism of methanol. I. Formation and crystallization of methanol-oxidizing enzyme in a methanol-utilizing yeast, Kloeckera sp. No. 2201, Agric. Biol. Chem., 36, 68, 1972.

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Chemistry and Biochemistry of Flavoenzymes

21. Tani, Y., Miya, T., and Ogata, K., The microbial metabolism of methanol. II. Properties of crystalline alcohol oxidase from Kloeckera sp. No. 2201, Agric. Biol. Chem., 36, 76, 1972. 27a. Hopkins, T. R., Unpublished results. 28. Kato, N., Omori, Y., Tani, Y., and Ogata, K., Alcohol oxidases of Kloeckera sp. and Hansenula polymorpha. Catalytic properties and subunit structures, Eur. J. Biochem., 64, 341, 1976. 29. Sahm, H. and Wagner, F., Microbial assimilation of methanol. The ethanol- and methanol-oxidizing enzymes of the yeast Candida boidinii, Eur, J. Biochem., 36, 250, 1973. 30. Sahm, H., Oxidation of formaldehyde by alcohol oxidase of Candida boidinii, Arch. MicrobioL, 105, 179, 1975. 31. Fujii, T. and Tonomura, K., Oxidation of methanol, formaldehyde and formate by a Candida species, Agric. Biol. Chem., 36, 2297, 1972. 32. Fujii, T. and Tonomura, K., Oxodiation of methanol and formaldehyde by a system containing alcohol oxidase and catalase purified from Candida sp. N-16, Agric. Biol. Chem., 39, 2325, 1975. 33. Yamada, H., Shin, K.-C., Kato, N., Shimizu, S., and Tani, Y., Purification and characterization of alcohol oxidase from Candida 25-A, Agric. Biol. Chem., 43, 877, 1979. 34. Couderc, R. and Baratti, J., Oxidation of methanol by the yeast, Pichia pastoris. Purification and properties of the alcohol oxidase, Agric. Biol. Chem., 44, 2279, 1980. 35. Patel, R. N., Hou, C. T., Laskin, A. J., and Derelanko, P., Microbial oxidation of methanol. Properties of crystallized alcohol oxidase from a yeast, Pichia sp., Arch. Biochem. Biophys., 210, 481, 1981. 36. Nishida, A. and Eriksson, K. E., Formation, purification and partial characterization of methanol oxidase hydrogen peroxide-producing enzyme in Phanerochaete chrysosporium, BiotechnoL Appl. Biochem., 9, 325, 1987. 37. Hopkins, T. R. and Miiller, F., Biochemistry of methanol oxidase, in Microbial Growth on C} Compounds, Van Verseveld, H. W. and Duine, J. A., Eds., Martinus Nijhoff Publishers, Dordrecht, 1987, 150. 38. Sakai, Y. and Tani, Y., Simple purification and improvement of stability of alcohol oxidase from a methanol yeast, Candida boidinii S2, by forming an enzyme-azide complex, Agric. Biol. Chem., 52, 227, 1988. 39. Guiseppin, M. L. F., Van Eyk, H. M. J., Hellendorn, M., and Van Almkerk, J. W., Cell wall strength of Hansenula polymorpha in continuous cultures in relation to recovery of methanol oxidase, Appl. Microbiol. BiotechnoL, 27, 31, 1987. 40. Hopkins, T. R., Alcohol oxidase from Pichia-type yeast, U.S. Pat. 4,540,668, 1985; U.S. Pat., 4,619,898, 1986. 41. Boys, C. W. G., Hill, D. J., Stockley, P. G., and Woodward, J. R., Crystallization of alcohol oxidase from Pichia pastoris, J. Mol. Biol., 208, 211, 1989. 42. Vonk, J. and Van Bruggen, E. F. J., Electron microscopy and image analysis of two-dimensional crystals and single molecules of alcohol oxidase from Hansenula polymorpha, Biochem. Biphys. Acta, 1038, 74, 1990. 43. Ghisla, S. and Massey, V., Mechanisms of flavoprotein-catalyzed reactions, Eur. J. Biochem., 181, 1, 1989. 44. Hopkins, T. R., Red absorbing combination of alcohol oxidase and an azide compound, U.S. Patent, 4,430,427, 1984. 45. Geissler, J. and Hemmerich, P., Yeast methanol oxidases: an unusual type of flavoprotein, FEBS Lett., 126, 152, 1981. 46. Matheson, I. B. C., O'Kane, D. J., and Lee, J., Free radical participation in bacterial bioluminescence, Free Rad. Res. Comms.t 2, 1, 1986. 47. Ehrenberg, A., Miiller, F., and Hemmeich, P., Basicity, visible spectra and electron spin resonance of flavosemiquinone anions, Eur. J. Biochem., 2, 286, 1967. 48. Massey, V. and Palmer, G., On the existence of spectrally distinct classes of flavoprotein semiquinones. A new method for the quantitative production of flavoprotein semiquinones, Biochemistry, 5, 3181, 1966. 49. Palmer, G., Muller, F., and Massey, V., Electron paramagnetic resonance studies on flavoprotein radicals, in Flavins and Flavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 123. 50. Mincey, T., Tayrien, G., Mildvan, A. S., and Abeles, R. H., Presence of a flavin semiquinone in methanol oxidase, Proc. Natl. Acad. Sci. U.S.A., 77, 7099, 1980. 51. Kurreck, H., Bock, M., Bretz, N., Eisner, M., Kraus, H., Lubitz, W., Muller, F., Geissler, J., and Kroneck, P. M. H., Fluid solution and solid-state electron nuclear double resonance studies of flavin model compounds and flavoenzymes, J. Am. Chem. Soc., 106, 737, 1984. 52. Choong, Y. S. and Massey, V., Stabilization of lactate oxidase flavin anion radical by complex formation, J. Biol. Chem., 255, 8672, 1980. 53. Blankenhorn, G., Ghisla, S., and Hemmerich, P., Model studies on flavin-dependent carbonyl-activations: reduction of carbonyl compounds by flavosemiquinone, Z. Naturforsch., 27b, 1038, 1972. 54. Sherry, B. and Abeles, R. H., Mechanism of action of methanol oxidase, reconstitution of methanol oxidase with 5-deazaflavin, and inactivation of methanol oxidase by cyclopropanol, Biochemistry, 24, 2594, 1985.

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55. Geissler, J., Ghisla, S., and Kroneck, P. M, H., Flavin-dependent alcohol oxidase from yeast. Studies on the catalytic mechanism and inactivation during turnover, Eur. J. Biochem., 160, 93, 1986. 56. Fall, H. H. and Petering, H. G., Metabolite inhibitors. I. 6,7-Dimethyl-9-formylmethylisoalloxazine, 6,7-dimethyl-9-(2'-hydroxyethyl)-isoalloxazine, J. Am. Chem. Soc,, 78, 377, 1956. 57. Muller, F. and Dudley, K. H., The synthesis and borohydride reduction of some alloxazine derivatives, Helv. Chim. Acta, 54, 1487, 1971. 58. Bystrykh, L. V., Romanov, V. P., Steczko, J., and Trotsenko, Y. A., Catalytic variability of alcohol oxidase from the methylotrophic yeast Hansenula polymorpha, Biotechn. Appl. Biochem., 11, 184, 1989. 59. Schoen, K. and Gordon, S. M., Water-soluble methylol derivatives of riboflavin, Arch. Biochem., 22, 149, 1949. 60. Cromartie, T. H., Sulfhydryl and histidinyl residues in the flavoenzyme alcohol oxidase from Candida boidinii, Biochemistry, 20, 5416, 1981. 61. Porter, D. J. T., Voet, J. G., and Bright, H. J., Direct evidence for carbanions and covalent N(5)-flavin carbanion adducts as catalytic intermediates in the oxidation of nitroethane by D-amino acid oxidase, J. Biol. Chem., 248, 4400, 1973. 62. Massey, V., Ghisla, S., and Kieschke, K., Studies on the reaction mechanism of the flavoenzyme lactate oxidase. Formation of two covalent flavin-substate adducts on reaction with glycolate, /. Biol. Chem., 255, 2796, 1980. 63. Dawson, A. P. and Thorne, C. J. R., The reaction of mitochondrial L-3-glycerophosphate dehydrogenase with various electron acceptors, Biochem. J., 114, 35, 1969. 64. Cromartie, T. H., Irreversible inactivation of the flavoenzyme alcohol oxidase by cyclopropanone, Biochem. Biophys. Res. Commun., 105, 785, 1982. 65. Sanders, J. K. M., NMR spectroscopy in the study of Cl-metabolism, in Microbial Growth on Cl Compounds, Van Verseveld, H. W. and Duine, J. A., Eds., Martinus Nijhoff Publishers, Dordrecht, 1987, 113. 66. Roa, M. and Blobel, G., Biosynthesis of peroxisomal enzymes in the methanotrophic yeast Hansensula polymorpha, Proc. Natl. Acad. Sci. U.S.A., 80, 6872, 1983. 67. Roggenkamp, R., Janowiez, Z., Stanikowski, B., and Hollenberg, C., Biosynthesis and regulation of the peroxisomal methanol oxidase from methanotrophic yeast Hansenula polymorpha, Mol. Gen. Genet., 194, 489, 1984. 68. Ellis, S. B., Brust, P. F., Kuntz, J. J., Waters, A. F., Harpold, M. M., and Gingeras, T. R., Isolation of alcohol oxidase and two other methanol regulatable genes from the yeast Pichia pastoris, Mol. Cell. Biol., 5, 1111, 1985. 69. Cregg, J. M., Barringer, K. J., Hessler, A. Y., and Madden, K. R., Pichia pastoris as a host system for transformations, Mol. Cell Biol., 5, 3376, 1985. 70. Cregg, J. M., Genetics of methylotrophic yeasts, in Microbial Growth on Cl Compounds, Van Verseveld, H. W. and Duine, J. A., Eds., Martinus Nijhoff Publishers, Dordrecht, 1987, 158. 71. Patel, R. N., Hou, C. T., Laskin, A. L, and Derelanko, P., Microbial oxidation of methanol: properties of crystallized alcohol oxidase from Pichia sp., in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Eds., Elsevier/North-Holland, New York, 1982, 196. 72. Visser, A. J. W. G., Grande, H. J., Muller, F., and Veeger, C., Intrinsic luminescence studies on the apoenzyme and holoenzyme of lipoamide dehydrogenase, Eur. J. Biochem., 45, 99, 1974. 73. Hopkins, T. R., A multipurpose enzyme sensor based on alcohol oxidase, Amer. Biotechn. Lab., 3, 32, 1985. 74. Smith, V. J., Green, R. A., and Hopkins, T. R., Determination of aspartame using an alcohol oxidase enzyme electrode, J. Assoc. Off. Anal. Chem., 72, 30, 1989. 75. Becker, K., Hopkins, T. R., and Schirmer, R. H., Hypothesis: antimalarial activity of the ethanol/alcohol oxidase system in vitro, Free Rad. Res. Comms., 9, 33, 1990.

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Chapter 5

LIPOAMIDE DEHYDROGENASE, GLUTATHIONE REDUCTASE, THIOREDOXIN REDUCTASE, AND MERCURIC ION REDUCTASE — A FAMILY OF FLAVOENZYME TRANSHYDROGENASES Charles H. Williams, Jr.

TABLE OF CONTENTS I.

Introduction A. Definition of the Family B. Other Reviews and Related Reviews in This Series

123 123 123

II.

Comparisons Among Family Members A. Mechanistic Similarities B. Structural Similarities 1. Amino Acid Sequence Homology 2. X-ray Crystal Structures

123 123 127 127 134

III.

Lipoamide Dehydrogenase A. General Properties 1. Metabolic Roles 2. Distribution 3. Physical Properties B. Characteristics of Two-Electron Reduced Enzyme 1. Spectral Characteristics 2. Reaction of EH2 with Arsenite 3. Oxidation-Reduction Potentials of EH2 C. Kinetics of the Forward and Reverse Reactions 1. Presteady-State Kinetics 2. Steady-State Kinetics 3. Alternate Electron Acceptors D. Distinct Functions of the Nascent Thiols E. Electron Transfer Between the Nascent Dithiol and FAD 1. Extreme Negative Cooperativity 2. NAD + — An Effector in the Lipoamide Dehydrogenase Reaction F. Acid-Base Chemistry in the Catalytic Cycle G. Special Properties of the E. coli Enzyme 1. Factors Affecting the Amount of Charge-Transfer Complex 2. Kinetics of the Forward Reaction 3. Site-Directed Mutagenesis

135 135 135 137 137 138 138 139 139 139 139 141 142 142 143 144

Glutathione Reductase A. Metabolic Roles and Distribution B. Distinct Functions of the Nascent Thiols C. Characteristics of Two-Electron Reduced Enzyme

153 154 155 156

IV.

144 145 147 147 151 152

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Chemistry and Biochemistry of Flavoenzymes 1, 2, 3, 4, D. E.

V.

VI.

VII.

Spectral Characteristics Presteady-State Kinetics Steady-State Kinetics Inactivation of Glutathione Reductase by NADPH and NADH 5, Redox Potential Acid-Base Chemistry in the Catalytic Cycle 1, Studies Utilizing Absorption Spectral Properties 2. Studies Utilizing Kinetic Properties Inhibitors and Chemical Modification

156 156 157 159 161 161 161 162 164

Thioredoxin Reductase A. Metabolic Roles and Distribution B. Comparisons and Contrasts with the Other Members of the Family 1, Redox Potentials and Evidence for an Active-Site Base 2. Kinetics of the Reduction of Thioredoxin by NADPH 3, Formation of a C(4a)-Thiolate Adduct with 1-deazaFAD 4. Photoreduction of Thioredoxin Reductase C. Distinct Functions of the Nascent Thiols D. Catalytic Activity of TRR(Serl35 Cysl38) and TRR(Cysl35 Serl38)

165 165

Mercuric Ion Reductase A. Metabolic Role and Distribution B. Kinetics 1. Reduction of the Enzyme by NADPH 2. Steady-State Kinetics C. Active Site Thiols 1. The Active-Site Disulfide — Cys-135, Cys-140 2. The Auxiliary Thiols — Cys-558, Cys-559 D. Electron Transfer Via a Thiol to C(4a)-FAD Adduct and Differential Reactivity of the Subunits

178 178 179 179 180 180 180 181

Other A. B. C. D. E. F.

189 189 191 191 191 192 192

Pyridine Nucleotide-Disulfide Oxidoreductases Trypanothione Reductase Bis-^-glutamylcystine Reductase Cystine Reductase Asparagusate Dehydrogenase Sulfhydryl Oxidase NADH Peroxidase

168 169 172

172 174 174 175

186

Acknowledgments

195

References

195

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123

I. INTRODUCTION A. DEFINITION OF THE FAMILY The pyridine nucleotide-disulfide oxidoreductases form a family of homo-dimeric flavoenzymes having a redox-active disulfide and a FAD in each monomer. The well characterized members of this group are lipoamide dehydrogenase [EC. 1.8.1.4], glutathione reductase [EC. 1.6.4.2], thioredoxin reductase [EC. 1.6.4.5], and mercuric ion reductase [EC. 1.16.1.1] (not technically a disulfide reductase). Lipoamide dehydrogenase and glutathione reductase are convenient misnomers which are more properly dihydrolipoamide dehydrogenase and glutathione disulfide reductase. Other members of the family include trypanothione reductase [EC.1.6.4.8], CoA-glutathione reductase [EC.1.6.4.6], cystine reductase [EC.1.6.4.1], asparagusate reductase [EC.1.6.4.7], bis-^-glutamylcystine reductase and sulfhydryl oxidase. With the exceptions of mercuric ion reductase and sulfhydryl oxidase, all enzymes in this family catalyze electron transfer between NAD(P)(H) and a disulfide/ dithiol. NADH peroxidase, while not containing a redox-active disulfide, should be considered with this family. B. OTHER REVIEWS AND RELATED REVIEWS IN THIS SERIES Three recent reviews offer excellent collateral coverage of the general area of the present review.1"3 Earlier reviews of these enzymes contain still relevant material which can not be fully covered in modern reviews, and they add a historical perspective.4"6 Attention is called to four reviews in this series which overlap with this chapter.7"10 Another recent review touches on this family of enzymes in the broader context of thiol-disulfide exchange.11 A brief review on flavoprotein mechanisms touches on this enzyme family. 12

II. COMPARISONS AMONG FAMILY MEMBERS A. MECHANISTIC SIMILARITIES This discussion will be confined to the four best studied members of the family. The flow of electrons in these enzymes is from pyridine nucleotide to FAD to active center disulfide to substrate disulfide or to mercuric ion.13"18 Physiologically, lipoamide dehydrogenase operates in the opposite direction but it also catalyzes the reverse reaction, reduction of lipoamide by NADH, at a high rate over a broad range of pH.18"19 Electron transfer between the flavin and the active center disulfide probably involves a covalent thiolate adduct at the C(4a) position of FAD.20"23 Lipoamide dehydrogenase exhibits "ping-pong" kinetics13-24 as does glutathione reductase at physiological substrate levels;6-25"27 rapid displacement of NADP + by NADPH complicates the kinetic picture in the latter enzyme.28 Although these enzymes are capable of being reduced with four electrons per subunit, the catalytically active species is twoelectron reduced enzyme, EH2.13 It is convenient then to think in terms of the two half reactions—Eox to EH2 and EH2 to Eox, where the EH2 state is stable under anaerobic conditions. In the EH2 state the active-site disulfide is reduced and the FAD is oxidized, but there is charge-transfer between one nascent thiol anion and the FAD, giving rise to a characteristic absorption band with a maximum at 530 to 540 nm.16"18-29"31 (A "nascent thiol" is one produced by reduction of the active site disulfide.) In this review, the term EH2 will refer to the mixture of two-electron reduced species. In the case of lipoamide dehydrogenase, glutathione reductase, and mercuric reductase this is primarily the charge-transfer complex. The EH2 state of thioredoxin reductase is a near equal mixture of enzyme having FAD and dithiol and of enzyme having FADH2 and disulfide.32 Figure 1 shows the spectrum of lipoamide dehydrogenase at the Eox, EH2, and EH4 levels; analogous spectra of glutathione reductase and mercuric ion reductase are very similar. The spectral properties of the FAD

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 1. Spectra of lipoamide dehydrogenase. Oxidized enzyme, solid line; two-electron reduced enzyme, dot dash line; four-electron reduced enzyme, dashed line. The reductant was dithionite.

in EH2 depend on its environment, in particular on the juxtaposition of the thiolate which is the donor in the charge-transfer and on those groups which influence the thiolate pKa value. The schemes in Figures 2 and 3 show analogous mechanisms for lipoamide dehydrogenase and glutathione reductase.33'34 They derive from earlier proposals of Sanadi5 and Massey4 and they are useful as working hypotheses. The physiological direction of catalysis is clockwise in both cases emphasizing the fact that while lipoamide dehydrogenase catalyzes the reduction of NAD + , NADPH is oxidized by glutathione reductase. Referring to Figure 2, EH2, in the absence of NAD + , is an equilibrium mixture of species 3 and 4 when species 4 is the charge-transfer complex.30 The rate limiting step is the reduction of Eox by dihydrolipoamide;13 subsequent reoxidation of EH2 by NAD + is very fast. In glutathione reductase the rate limiting step is less clear, but appears to be the reduction of glutathione disulfide by EH2.6>35'36 There is at least one base in the active sites of lipoamide dehydrogenase, glutathione reductase and thioredoxin reductase.31"33'37 The evidence for this will be discussed separately for each enzyme. The identification of the base in erythrocyte glutathione reductase as His467' hydrogen bonded to Glu-472' comes from the crystal structure and the amino acid sequence (Section II.B). 3741 His-444' and Glu-449' are the analogous residues in E. coli lipoamide dehydrogenase.42 The histidine residue was identified by a novel procedure in which a bifunctional arsenoxide was first reacted with dihydrolipoamide bound to the transacetylase and then cross-linked to the histidine in lipoamide dehydrogenase.43"45 Table 1 is a list of important residues in glutathione reductase38 with the corresponding residue in lipoamide dehydrogenase and mercuric reductase.42 The schemes in Figures 2 and 3 indicate that the nascent thiols have distinct functions. The nascent thiolate alkylated by iodoacetamide and interacting directly with the disulfide substrate (lower sulfur, referred to as the interchange or distal thiol) is that nearer the Nterminus of the protein in lipoamide dehydrogenase,23'34'46'47 glutathione reductase,33'34'37'39'40 and mercuric reductase;48 it is distinct from the charge-transfer donor thiolate (upper sulfur, proximal). The population of molecules with the upper thiol deprotonated can exhibit chargetransfer while those molecules with the lower thiol deprotonated can be alkylated or interchange with glutathione disulfide to initiate the second half-reaction in glutathione reductase

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FIGURE 2. Scheme for the mechanism of lipoamide dehydrogenase. FAD, F; active-site base, B; redox-active disulfide, S-S. Charge transfer is indicated by a dashed arrow. Species 2 is a mixed disulfide between the interchange thiol and dihydrolipoamide. Dihydrolipoamide, Lip(SH)2; lipoamide, Lip(S)2. (From Arscott, L. D., Thorpe, C. and Williams, C. H., Jr., Biochemistry, 20, 1513, 1981. With permission.)

FIGURE 3. Scheme for the mechanism of glutathione reductase. Symbols are as in Figure 2. Species 8 is a mixed disulfide between the interchange thiol and glutathione. G-S-S-G, glutathione disulfide; G-S-H, glutathione.

125

126

Chemistry? and Biochemistry of Flavoenzymes TABLE 1 Important Residues in Glutathione Reductase with Homologues in Lipoamide Dehydrogenase and Mercuric Reductase

Residue

Group of atom

Cys-2 Val-25 Ile-26 Gly-27 Gly-29 Gly-29 Gly-31 Gly-32 Gly-32 Leu-33 Ala-36 Arg-37 Gly-43 Glu-50 Ser-51 Gly-55 Gly-56 Thr-57 Thr-57 Cys-58 Cys-63 Lys-66

Side chain Side chain Side chain

Lys-67 Cys-90 Tyr-114 Lys-120 Gly- 128 His- 129 Ala- 130 Ala- 130 Ala- 155 Thr-156 Gly- 157 Gly-158 He- 192 Val-193 Gly-194 Gly-196 Tyr-197 Ala- 199 Val-200 Glu-201 Ala-203 Arg-218 His-219 Arg-224 Gly-27 1 Arg-291

Side chain Side chain Side chain Side chain

Pro-293 Asn-294 Gly-304 Gly-311

— — Amide NH Amide NH

—

Amide NH Side chain Side chain Side chain

— Side chain Amide NH

— — Side chain O Amide NH Side chain Side chain Side chain

— Side chain Amide NH Carbonyl O Carbonyl O Carbonyl O

— — Side chain Side chain

— — Side chain Side chain Side chain Side chain Side chain Side chain Side chain Side chain

— Side chain

— — — —

Function or contact On flexible extension Apolar Apolar Bend Bend H-bond to pyrophosphate through HOH H-bond to pyrophosphate Tight fit of its helix to sheet H-bond to pyrophosphate through HOH Apolar Conserved GSSG binding — ion-pair Bend Binds tightly to ribose hydroxyls H-bond to adenine N-3 Bend Bend H-bond to pyrophosphate H-bond to pyrophosphate Thiol-disulfide interchange Electron transfer to the FAD Increase FAD electrophilicity, ion-pair with Glu201, proton acceptor from NADPH GSSG binding — ion-pair Intersubunit disulfide GSSG contact GSSG binding — ion-pair Bend H-bond to adenine NH 2 H-bond to adenine N-l H-bond to adenine NH2 H-bond to pyrophosphate through HOH H-bond to pyrophosphate through HOH End of FAD domain Beginning of NADP domain Apolar Apolar Bend Bend Protects FAD from water (- NADP) Gly in most nucleotide sites Apolar Ion pair with Lys-66 Conserved Ion pair with 2'-phosphoryl NADP Ion pair with 2'-phosphoryl NADP Ion pair with 2'-phosphoryl NADP Bend Increase FAD electrophilicity, ion pair with Asp-331 via HOH End of NADP domain Beginning of central domain Bend Bend

Residue in LipDH

Residue in MR

None Val-11 He- 12 Gly- 13 Gly- 15 Gly- 15 Gly- 17 Gly- 18 Gly- 18 Tyr-19 Ala-22 Ile-23 Gly-29 Glu-36 Lys-37 Gly-42 Gly-43 Thr-44 Thr-44 Cys-45 Cys-50 Ser-53

Asp-80 Val-103 Ile-104 Gly- 105 Gly- 107 Gly- 107 Ala- 109 Ala- 110 Ala-110 Met- 111 Ala- 114 Leu-115 Gly- 121 Glu-128 Arg-129 Gly- 133 Gly- 134 Thr-135 Thr-135 Cys-136 Cys-141 Ser-144

Lys-54

Lys-145

— IIe-103 Lys-108 Gly- 117 Tyr-118 Gly- 119 Gly- 119 Ala- 146 Thr-147 Gly- 148 Ser-149 Val-182 He- 183 Gly- 184 Gly- 186 Val-187 Gly- 189 Leu- 190 Glu-191

— Tyr-194

—

Phe-208 Leu-209 Pro-214 Gly-263 Arg-279

Gly-209 Glu-210 Ala-211 Ala-211 Ala-239 Thr-240 Gly-241 Ala-242 Val-275 Ile-276 Gly-277 Ser-279 Val-280 Ala-282 Leu-283 Glu-284 Ala-286 Arg-301 Ser-302 Arg-308 Gly-350 Arg-363

Pro-281 Phe-282 Gly-292 Gly-299

Pro-365 Asn-366 Gly-376 Gly-383

—

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127

TABLE 1 (continued) Important Residues in Glutathione Reductase with Homologues in Lipoamide Dehydrogenase and Mercuric Reductase Residue

Group of atom

Val-329 Gly-330 Asp-331

Carbonyl O

Asp-331 Gly-334 Thr-339 Thr-339 Arg-347 Tyr-364 Asn-365 Val-370 Gly-437 Gly-439 Gly-454 His-467' His-467' His-467' His-4677 Glu-472' GIu-473'

Amide NH

— Side chain

— Side chain Amide NH Side chain

— — Carbonyl O

— — — Imidazole N-3 Imidazole Imidazole N-l Carbonyl O Side chain Side chain

Function or contact H-bond to pyrophosphate through HOH Bend Ion pair with Arg-291 through HOH, H-bond to ribityl hydroxyls-3,5 H-bond to pyrophosphate Bend H-bond to charge-transfer thiolate H-bonds to FAD N-l and C-2-O GSSG binding — ion-pair End of central domain Beginning of interface domain H-bond to pyridinium CONH2 Bend Bend Bend Ion pair with redox thiols, proton donor to GS~ Increase FAD electrophilicity (EH2) Strong H-bond to Glu-472' H-bond with NH-3 of FAD Strong H-bond to His 467' GSSG binding — ion-pair with both ammonium groups

Residue in LipDH

Residue in MR

Ile-317 Gly-318 Asp-319

Ala~401 Gly-402 Asp-403

Asp-319 Gly-323 Ala-327 Ala-327 Ile-335 Tyr-350 Asn-351 Val-356 Gly-421 Gly-423 Gly-438 His-451 His-451 His-451 His-451 Glu-456 Ala-457

Asp-403 Gln-407 Val-411 Val-411 Thr-419 Leu-434 Thr-435 Val-440 Ala-505 Glu-507 Arg-522 Tyr-535 Tyr-535 Tyr-535 Tyr-535 Glu-540 Gly-541

Note: LipDH, lipoamide dehydrogenase consensus sequence, Figure 5; MR, mercuric reductase consensus sequence, Figure 7; GSSG, glutathione disulfide; GS~, glutathione anion.

FIGURE 4. Scheme for the formation and breakdown of the thiolate to C(4a)-FAD adduct. (From Thorpe, C. and Williams, C. H., Jr., Biochemistry, 20, 1507, 1981. With permission.)

catalysis. The same differential functions have been shown in thioredoxin reductase.49 Spectral properties of lipoamide dehydrogenase having the interchange thiol alkylated, of thioredoxin reductase substituted with 1-deaza-FAD, and of mercuric reductase at low pH indicate that the charge-transfer donor thiolate (nearer the C-terminus) can form a covalent bond with the C(4a) position of the flavin and it has been suggested that this is the mechanism of electron transfer (Figure 4).20-23-47 B. STRUCTURAL SIMILARITIES 1. Amino Acid Sequence Homology Lipoamide dehydrogenase, glutathione reductase, mercuric reductase and trypanothione

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Chemistry and Biochemistry of Flavoenzymes

reductase have a high degree of homology in amino acid sequence which extends to all domains.42-50"57 The structural similarity was first observed in peptides containing the redox active disulfide of lipoamide dehydrogenase and glutathione reductase.58"64 This is also seen in mercuric reductase.48 The region of the active site disulfide in thioredoxin reductase was clearly different.63-65'66 However, homology in the FAD-binding region and in the pyridine nucleotide binding region was a common feature in all members of the family.67 The complete amino acid sequences of lipoamide dehydrogenase from six species are known and four of these have been compared.68 A true consensus sequence among the three prokaryotes, £. coli,42 Pseudomonas putida,69 and Azotobacter vinelandii™ (all Gram negative), and the three eukaryotes, human,51-53 pig,51 and yeast71*72 is not possible due to the very early divergence of the bacterial enzyme. The six sequences are shown in Figure 5 together with a "consensus" sequence. There is identity among at least four sequences at 47% of the positions and there is homology among all six sequences at 46% of the positions. An identity matrix is shown in Figure 6. The highest concentration of invariant positions outside the active site is not, as might be guessed, in the FAD or NAD nucleotide binding folds, but rather in the interface domain. There are 25 invariant positions in the interface domain: 6 Gly, 4 Ala, 6 Glu, 3 Pro, and one each of Phe, Asp, Val, Leu, Arg and His. These include His-451', the active site base, and Glu-456', which interacts with His-451'. Returning to the sequence around the active site cystine, the very impressive identity among the six species of lipoamide dehydrogenase extending over 17 residues from Leu-40 (E coli sequence, Figure 5) through Leu-56 and broken by only two conservative changes, tempts one to speculate on its necessity. An important exception to this "rule" is found in the sequence from Peptococcus glycinophilus where an Arg replaces Val-47 — hardly a conservative change (see also Section V.A and Reference 333).73 Such a finding should be attributed to experimental error except for the recent finding in Spirulina sp. glutathione reductase in which He and Arg replace Asn-60 and Val-61 at the homologous positions (Figure 7).74 These could be thought of as compensating changes since neither is conservative, but there is no change at Asn-46 in the Peptococcus glycinophilus sequence. Another exception to a rule that is not a rule is the presence of Phe at position 449 in the 9-residue essentially invariant sequence extending to Ala-457 in the A. vinelandii lipoamide dehydrogenase sequence which includes the active center base, His-451 (Figure 5); the other five species have a second His at this position. It is unlikely that thiols in lipoamide dehydrogenase, aside from the nascent thiols, have any important function since the A. vinelandii enzyme has only one other Cys and this is homologous only with one (of 7) of the Cys of P. putida enzyme. Figure 7 gives an alignment of the four homologous members of the family using the consensus sequences for lipoamide dehydrogenase and mercuric reductase. The alignment was done manually starting with several regions of high homology and working out in both directions. Figure 6 gives an identity matrix for the family which includes both known glutathione reductase sequences.54-55 The level of identity between the four enzymes varies between 24 and 40%; the level of homology is very impressive. The level of identity between prokaryote and eukaryote sequences is higher in the case of glutathione reductase than with lipoamide dehydrogenase. As expected, trypanothione reductase is more closely related to glutathione reductase than to the other enzymes and is more closely related to human than to E. coli glutathione reductase.57 The mercuric reductase identity matrix shows the already noted relationship between mercuric reductase derived from plasmids harbored by Gram positive (RC607 and pI258) and Gram negative (Tn501 and Tn21) organisms.75"77 Clearly homology within this enzyme family extends throughout the molecule (Figure 7). All four sequences are identical at 12% of the positions and a further 12% are similar. The level of homology is lowest from the middle to just before the end of the FAD domain where only two positions show identity. Aside from the active site and the nucleotide binding folds,

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FIGURE 5. Amino acid sequences of (top to bottom) lipoamide dehydrogenase from pig adrenal medula (heart),46 human liver,46*53 yeast,71 Escherichia coli,42 Azotobacter vinelandii,70 Pseudomonasputida,™ and a "consensus" sequence. Positions lacking identity among at least three sequences are underlined. The pig sequence is the default. Positions homologous with glutathione reductase at breaks between the FAD, pyridine nucleotide (PN), central (Cen) and interface (Inf) domains are shown.54 The A. vinelandii enzyme contains an insert: K E G K T A, indicated by a slash.

129

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 5.

(continued).

there are five areas of very impressive homology between the five (or four) sequences: one at the beginning of the pyridine nucleotide domain (residues 153 to 158 in the glutathione reductase sequence), one at the interface between the pyridine nucleotide and central domains (285 to 305), one in the central domain (326 to 333) and two in the interface domain (363 to 386 and 430 to 451). From the crystal structure discussed below, it can be seen that carbonyls of the polypeptide chain at the end of the FAD domain form hydrogen bonds through water molecules to the pyrophosphate of the FAD (Table I).38 The region at the end of the pyridine nucleotide domain contains an Arg thought to be important in modulating the redox properties of FAD.78 Homology in the interface domain emphasizes the importance of the dimeric structure to be discussed later. Three segments of the thioredoxin reductase sequence are also shown in Figure 7 and these are considered in the family identity matrix of Figure 6.79 Since the flavin nucleotide binding domain of the four homologous enzymes is near the N-terminus, the first thioredoxin reductase segment—the N-terminal 42 residues—is aligned with that region. The second segment (residues 105 to 206) is aligned with the pyridine nucleotide domain of the other enzymes. The third segment, comprising the 56 C-terminal residues, is aligned with the central domain, referring to glutathione reductase. There is homology between these three segments of thioredoxin reductase and at least two of the other four sequences at 82 of the 200 positions considered (43, 46, and 30% homology in the three segments). The active center disulfide is in the second segment between two highly conserved stretches, but in an immediate area where the other sequences are not homologous. The alignment is diagrammed in Figure 8. The levels of identity between the three segments of thioredoxin reductase and

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FIGURE 6. Identity matrices for the family. Human glutathione reductase, H-GR; E. coli glutathione reductase, E-GR; lipoamide dehydrogenase consensus sequence, C-LD; Trypanosoma congolense trypanothione reductase, T-TpR; mercuric reductase consensus sequence, C-MR; E. coli thioredoxin reductase, E-TrR. Azotobacter vinelandii and Pseudomonas putida lipoamide dehydrogenases, A. vin. and P. put. respectively. The mercuric reductase designations are as follows: chromosomal Bacillus sp., strain RC607; pI258 in Staphylococcus aureus; Tn501 on pVSl in Pseudomonas aeruginosa; and Tn21 on R100 in Shigellaflexnerii.

the other members of the family average about 23%. Thus, it would seem that while thioredoxin reductase has acquired its nucleotide binding regions by divergent evolution, its active site has evolved convergently. The third segment — the C-terminal 56 residues — while less homologous on the above basis with the other sequences, is identical with mercuric reductase at 27% of the positions leaving little doubt of a relationship. This segment includes

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 7. Sequence homology between (top to bottom) human erythrocyte glutathione reductase,54 lipoamide dehydrogenase "consensus" sequence (Figure 5), Trypanosoma congolense trypanothione reductase," the final 480 residues of mercuric reductase consensus sequence and three sequence segments of Escherichia coli thioredoxin reductase.79 Identity or homology in the four sequences (excluding thioredoxin reductase) are designated by asterisk or ampersand, respectively. Positions in the thioredoxin reductase sequence showing homology with any two of the other sequences have been underlined. Inserts in the trypanothione reductase sequence are indicated by a slash: V H G P P F F A after position 38 and Q K N V V T V T E G after position 130. The mercuric reductase consensus sequence was derived from a comparison of four sequences using Tn501 as the default. 75 Breaks between the FAD, pyridine nucleotide (PN), central (Cen), and interface (Inf) domains of glutathione reductase are shown.54

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FIGURE 7

(continued).

FIGURE 8. Areas of amino acid sequence homology between thioredoxin reductase and the other members of the family as represented by glutathione reductase using the domain boundaries of glutathione reductase. The three solid portions of the thioredoxin reductase sequence represent the three segments of the sequence compared in Figure 7. The position of the active site is indicated in each case.

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Chemistry and Biochemistry of Flavoenzymes

a seven-residue stretch that is highly homologous in all five sequences. This is of interest since, like all other members of the family, thioredoxin reductase is a dimer, and one might have expected its C-terminal segment to be homologous with the interface domains of the other enzymes, rather than with the central domain. Again, this suggests an independent evolution of the domains; independent evolution of the catalytic and NAD + -binding domains of glyceraldehyde-3-phosphate dehydrogenase has been suggested.80 2. X-ray Crystal Structures The advent of a three-dimensional structure of an enzyme often forms a unique turning point in the course of research on that enzyme; such was certainly the case with the structure of human erythrocyte glutathione reductase.37'39'40 When combined with the amino acid sequence, the complete structure resulted.54-64'81 This was the first structure of a FADcontaining enzyme and the first structure of a "full-sized" flavoenzyme. The structure gave a clear picture of the catalytic events as outlined in Section II.A, and of both substrate binding sites. In addition to confirming the presence and juxtaposition of the active site cystine and the base, it revealed, and predicted functions for several groups not previously detected by any other means. Finally, it gave a rationale for the dimeric structures of this enzyme family and for the negative cooperativity observed in lipoamide dehydrogenase. Because the enzyme was active in the crystal, diffusion of substrates, products, and inhibitors into the crystal have provided difference maps rich in information.78'82'83 Refinement of the structure has revealed further important details.38 Although this structure will be reviewed in a separate chapter,7 it will perhaps be useful to insert a short description of the active site here both for completeness and to give the point of view of one who works primarily on mechanism. The domain boundaries are given in Figure 7. The active site contains key elements from the FAD, pyridine nucleotide and central domains of one polypeptide chain and from the interface domain of the other chain. Thus, the structure predicts that only dimeric enzyme should be active. The active site is divided by the isoalloxazine ring into a pyridine nucleotide compartment on the re side and a dithiol-disulfide interchange compartment on the si side — a ping-pong active site complete with a net. Cys-63 provides the nascent thiol interacting directly with the flavin as predicted. The sulfur of Cys-58 faces out into the glutathione disulfide binding pocket thus providing solid evidence for the suggestion that this nascent thiol would interact with the substrate;33 indeed, the mixed disulfide can be seen in difference maps of crystals soaked in glutathione.78 The base, His-467' is oriented with N(3) toward the active center disulfide, somewhat closer to the sulfur of Cys-58 than to Cys-63, and also close to the binding position of one sulfur of glutathione disulfide. (Residues designated with a prime come from the other polypeptide chain.) N(l) of the imidazole is oriented toward Glu-472' with which it forms a short, i.e., strong, hydrogen bond. In Eox, Glu-472' and His-467' are thought to share a single proton. There is a water molecule hydrogen bonded to N(3) of His-467'38 suggesting an analogy to chymotrypsin where the His-57 is hydrogen bonded to Asp-102 via N(l) and to Ser-195 via N(3). Such hydrogen bonding at N(3) serves to keep the His-Asp or His-Glu diad unprotonated in the resting enzyme. The water may also serve as a proton source in the formation of EH2, thus: N3-H-O-H-H-OH + H-Buffer *± N3H + H-O-H-H-O-H + Buffer One of the protons in EH2 is presumed to be on N(3) of His-467' and the other shared between the nascent thiols. Cys-58 is stabilized as a thiolate by ion-pair interaction with the imidazolium ion of His-467'. A long helix runs from Cys-58 to Pro-88; the dipole of this helix could provide further stabilization for negative charge in the area of the nascent thiols.84

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The charge-transfer interaction can stabilize Cys-63 as a thiolate. The fact that the extinction coefficient of the charge-transfer is high suggests that Cys-63 is largely present as a thiolate. A very exciting discovery of the structure is the fact that the isoalloxazine ring is surrounded by two ion-pairs in Eox, Lys-66/Glu-201 and Arg-291/Asp-331 (via a water), and a third ion-pair in EH2, His-467'/Glu-472'. Although the polypeptide chain including Lys-66 is on the si side, the methylene chain extends under the isoalloxazine ring placing the amino group on the re side, 0.33 nm from the flavin N(5); Glu-201 is then distal on the re side. In each case it is the positive end of the ion-pair that is closer to the flavin, leading to the very attractive suggestion that this influences the redox potential of the FAD.78 Glutathione disulfide binds asymetrically across the monomer-monomer interface; the half molecule which forms the mixed disulfide with Cys-58 is designated GS-I and the half molecule which leaves first as GS-II. The zwitterionic end of GS-II appears to be the most important determinant in binding. The imidazolium group of His-467' is well positioned to protonate the leaving glutathione,2'83 NADPH binds in an extended conformation in a long trough with many contacts at the adenosine end including a constellation of positive charges that confer specificity for that pyridine nucleotide. Indeed when NADH binds, a phosphate ion from the buffer is trapped at that locus. The incoming NADPH must displace the side chain of Tyr-197 which shields the pyridinium end of the site, and thus the FAD, from solvent. The pyridinium ring adopts a stacked configuration with respect to the re side of the isoalloxazine ring for good overlap of the TT orbitals. There is a very close approach of the pyridinium N(4) to the Lys-66/Glu201 ion-pair (0.25 nm) and to the flavin N(5) (0.35 nm). In the mechanism proposed by Pai and Schulz, Glu-201 is the proton acceptor implying that two electrons, rather than a hydride ion pass to the flavin and requiring the uptake of two protons on the si side; thus their species 2 is an EH3+ rather than an EH2. The resulting unpaired Lys-66 then serves to expel the NADP+.75-82 Since the structure defines the absolute stereochemistry of the flavinpyridine nucleotide interaction for this enzyme, glutathione reductase has been used in a novel way to determine the stereospecificity of other flavoenzymes in which the flavin can be replaced.85 The three-dimensional structure of Azotobacter vinelandii lipoamide dehydrogenase has proven a very difficult one to solve and required comparison to the glutathione reductase structure as well as isomorphous replacement techniques.86 A preliminary comparison had shown that the two structures showed remarkable similarity reflecting their mechanistic similarity.87 The homologies shown in Table 1 show that most crucial residues, aside from those involved in specificity, are identical. The yeast enzyme is also being studied using a similar combination of techniques.88

III. LIPOAMIDE DEHYDROGENASE A. GENERAL PROPERTIES h Metabolic Roles Lipoamide dehydrogenase functions as part of several mitochondrial multi-enzyme complexes. While its role in each case is the reoxidation of dihydrolipoic acid and reduction of NAD + , this function is required in four distinct metabolic settings: in the conversion of pyruvate to acetyl-CoA, in the conversion of a-ketoglutarate to succinyl-CoA, in the oxidation of the a-ketoacids resulting from the transamination of the branched chain amino acids, and in the conversion of glycine to A^Af10-methylenetetrahydrofolate, CO2 and NH^.89 In the complexes, the lipoic acid in all cases is bound in amide linkage to a lysine residue. This lysine residue is contained within a distinct domain of the transacylase component of the pyruvate and a-ketoglutarate dehydrogenase complexes, but is on a small carrier protein in the glycine cleavage system.90"93 Recent studies have shown that only the

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 9. Model for interactions of proteins of glycine decarboxylase with those of glycine reductase in Eubacterium acidaminophilum. PI, pyridoxal phosphate-containing decarboxylase protein; P2, lipoamide containing hydrogen carrier protein; P3, lipoamide dehydrogenase; P4, transferase protein; PA, selenoprotein; GR, glycine reductase; THF, tetrahydrofolate; CH2 = THF, N5,N10-methylenetetrahydrofolate; and cm, cytoplasmic membrane. (From Freudenberg, W., Mayer, F. and Andreesen, J. R., Arch, Microbiol, 152, 182, 1989. With permission.) The glycine cleavage and glycine reductase are as follows: Glycine + THF + R(S)2 -> CH2=THF + CO2 + NH3 + R(SH)2 CH^THF + 2NAD(P) + ADP + ?, -» THF + CO2 + 2NAD(P)H + ATP Glycine + R(SH)2 + ADP + P; -» Acetate + NH3 + ATP + R(S)2

R-enantiomer is utilized in the a-ketoacid dehydrogenase systems.94 Acetylation of reduced lipoamide in the a-ketoacid complexes is at the 8-position,95-96 rather than at the 6-position as previously thought.97 It is not known which lipoamide position is utilized in the mixed disulfide with lipoamide dehydrogenase (Figure 2). The a-ketoacid and glycine cleavage systems are further distinguished by the cofactor of the decarboxylase component, thiamine pyrophosphate or pyridoxal phosphate respectively, and by the final acceptor, CoA or tetrahydrofolate respectively.89'93'98*99 The glycine cleavage system from three sources, chicken liver, Peptostreptococcus glycinophilus, and Eubacterium acidaminophilum has been studied in some detail.93-98-99 The P. glycinophilus is interesting in that this is the first strict anaerobe from which lipoamide dehydrogenase was isolated and characterized.100-101 The reducing equivalents produced are presumably used to reduce more glycine via the Se-dependent glycine reductase in a reaction coupled to ATP synthesis (Figure 9).102 105 The physiological electron donor of glycine reductase is not known.102-103 The fact that dithiothreitol can serve as the electron donor for glycine reductase,102 suggests that the physiological donor may be 2-electron reduced lipoamide dehydrogenase (see Section III. A. 2). The involvement of a flavoprotein had been suggested.102

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A single gene codes for the lipoamide dehydrogenase functioning in the pyruvate and a-ketoglutarate complexes of E. coii.106 However, galactose- and maltose-stimulated lipoamide dehydrogenase activity has been detected in mutant E. coll lacking the Ipd gene product.107 Lipoamide dehydrogenase has been detected in the plasma membranes of several organisms and its potential role has been discussed.108 Sensitive methods for the detection of lipoic acid have been developed in this connection.109 A single gene is also suggested in humans from study of lactic acidosis.110-111 In Pseudomonas putida more than one gene has been demonstrated.112'113 The LPD1 gene of yeast has also been characterized.114 If only one gene codes for lipoamide dehydrogenase in mammals, the isozymes observed upon electrophoresis or anion exchange chromatography of the pig heart enzyme could be due to posttranslational modifications and/or to experimental artifact.6 2. Distribution Lipoamide dehydrogenase is ubiquitous in aerobic organisms including prokaryotes, eukaryotes, and some archaebacteria. Its presence in anaerobic organisms has not been widely tested, but as already mentioned, it is present in the obligate anaerobe Peptostreptococcus glycinophilus, and has recently been isolated from other glycine fermenting bacteria Clostridium cylindrosporum and Clostridium sporogenes.115 An enzyme referred to as atypically small lipoamide dehydrogenase has been isolated from another glycine fermenting anaerobe, Eubacterium acidaminophilum.n6 Unlike the lipoamide dehydrogenase from Peptostreptococcus glycinophilus which is normal in most of its properties,100-115 the enzyme from E. acidaminophilum has a subunit molecular weight of 35,000 rather than 50,000, prefers NADP + , and appears to be unusually difficult to reduce (see Section V.A).116 It has been shown that E. acidaminophilum, an atypically small lipoamide dehydrogenase, is associated with glycine reductase at the plasma membrane rather than with the glycine cleavage enzymes in the cytoplasm as shown in the scheme of Figure 9.117 In this position it can serve to transfer electrons both from the oxidative decarboxylation of glycine as well as the subsequent NAD(P)-dependent oxidation of Af5,Af10-methylenetetrahydrofolate. The typical lipoamide dehydrogenase of Peptostreptococcus glycinophilus may similarly be associated with glycine reductase and, as suggested above, serve directly as the electron donor of glycine reductase (Section III.A.I). Peptostreptococcus glycinophilus is known from its 16S RNA to be an ancient bacterium.118 It is likely that lipoamide dehydrogenase has always been present in these anaerobes, since the lipoic acid-dependent breakdown of glycine is central to the metabolism of these organisms.104 This argues against this organism having acquired lipoamide dehydrogenase adventitiously, late in its evolution.50 The list of sources of purified lipoamide dehydrogenase given in an earlier review6 may be supplemented with the following: Pseudomonas putida,69'112'113 Pseudomonas aeruginosa PAO,119 Pseudomonas fluorescens,120 Bacillus stearothermophilus,121 the parasitic roundworm, Ascaris lumbricoides var. suum,122 Ascaris suum muscle mitochondria,123 the thermophilic fungus Malbranchea pulchella var. sulfurea,124 and the halophilic archaebacterium Halobacterium halobium,125 as well as Eubacterium acidaminophilum, Clostridium cylindrosporum, and Clostridium sporogenes cited above.115-116 The Trypanosoma brucei enzyme has been studied in the purified plasma membrane.126 3. Physical Properties Lipoamide dehydrogenase as isolated from most sources is a relatively stable fluorescent protein. Unless noted, the properties described will be those of the pig heart enzyme. The spectrum shown in Figure 1 has a well resolved visible peak at 455 nm indicative of flavin in an apolar environment.127 The extinction coefficient at 455 nm is 11.3 mM~lcm~l. The ratio of the visible to the near-uv peak is 1.3 and the ratio of the peak at 273 nm to that at 455 nm, an excellent indicator of purity, is 5.3. The peak to trough ratios are 3.5 (455 nm/

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Chemistry and Biochemistry of Flavoenzymes

392 nm) and 2.8 (354 nm/310 nm). The trough at 310 nm fills in when the enzyme turns over aerobically with NADH as can happen in elution from affinity columns. New fluorescent emission peaks suggest that breakdown of histidine residues may be involved.128 The fluorescence excitation spectrum of lipoamide dehydrogenase is essentially that of the absorbance spectrum. The fluorescence emission maximum is at 520 nm and 530 nm for enzyme bound and free FAD respectively. The ratios of the emission at the respective maxima for enzyme bound and free FAD are 4.5 at 5°C, 2.7 at 25°C, and 1.6 at 45°C.4 Pulse fluorometry studies indicated that the flavin centers are not identical,129'130 and this could be related to observation of extreme negative cooperativity.23 The triplet yield is 14%. The circular dichroism of apo- and holo-lipoamide dehydrogenase has been carefully investigated.131'132 The data show that the FAD does not contribute significantly to the farultraviolet bands (190 to 240 nm). Thus, circular dichroism in this region will be a good indicator of protein structure in enzyme modified by site directed mutagenesis. The negative band near 370 nm in the circular dichroism spectrum of free FAD is replaced by a positive doublet and a broad shoulder extending to 290 nm. However, the same set of gaussian bands account satisfactorily for the spectrum in this region for both free and bound FAD suggesting that no new transitions are generated upon binding. The circular dichroism of the reduced enzyme will be discussed later. Magnetic circular dichroism studies have been reported.133 B. CHARACTERISTICS OF TWO-ELECTRON REDUCED ENZYME Two-electron reduced lipoamide dehydrogenase is an equilibrium mixture of several species. These are shown in Figure 2, species 2 to 8. The designation EH2 will refer to the mixture which is stable anaerobically. The oxidized enzyme is reduced rapidly and almost stoichiometrically by the substrate, dihydrolipoamide.18 The primary product is the chargetransfer complex, species 4, in which the thiolate nearer the C-terminus is the donor and FAD is the acceptor.46 The enzyme can also be reduced to EH2 by NADH and this process is very rapid and stoichiometric. The product is again the charge-transfer complex but with NAD + bound.13-18 Complete reduction of EH2 to EH4 requires dithionite (Figure 1) but EH2 is partially reduced by NADH at low pH; the degree of this over-reduction is species dependent (see Section III.G.I). 1. Spectral Characteristics The spectrum of EH2, which is predominantly the charge-transfer complex, is given in Figure 1. The two flavin bands are diminished somewhat and blue shifted. The new chargetransfer band extends beyond 600 nm imparting a red color; this can be monitored sensitively at 530 nm where oxidized enzyme and EH4 have no absorbance. The description of this species as a charge-transfer complex has been discussed.17'29-30'134 EH2 forms spectrally distinguishable complexes with both NAD + and NADH. Titration of EH2 (preformed by reduction with dithionite) with NADH causes an enhancement of the charge-transfer band and further diminution of the visible peak; changes in the near ultraviolet peak are much less than additive for additions of NADH up to one equivalent/FAD, that is EH2-NADH has less absorbance at 340 nm than EH2 + NADH.135 The spectral changes are quite similar to those observed earlier on titration of glutathione reductase EH2 with NADPH.28 Titration of EH2 with NAD + causes a shift of the charge-transfer band to longer wavelengths and further diminution of the visible peak. The resolution of the shoulders of the visible band, characteristic of the charge-transfer complex, is changed to that characteristic of the oxidized enzyme (Figure 1). There is an increase in the near ultraviolet peak at pH 6.3, but no change in this band at pH 5.8 indicating that a small amount of NADH is formed at pH 6.3. EH2 is of course reoxidized by NAD + at higher pH values (see Section III.C). However, the fact that the spectrum of enzyme reduced by stoichiometric NADH is distinct

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from that of enzyme reduced by dihydrolipoamide or dithionite indicates that an EH2-NAD + can form at pH 7.6. The changes in the spectrum of EH2 upon titration with NAD + are associated with an apparent KD of 280 jjiM at pH 6.3.135'137 The circular dichroism of lipoamide dehydrogenase reduced with equimolar NADH is different from that of enzyme reduced by dihydrolipoamide.132 The former would be EH2 - NAD + while the latter would be EH2 (see above). 2. Reaction of EH2 with Arsenite Inhibition of lipoamide dehydrogenase catalytic activity by arsenite or Cd2 + only when the enzyme is prereduced was the major clue to the identity of the second prosthetic group. Thus, it is EH2 that reacts with these reagents. 17>18 The specificity of these reagents for vicinal dithiols, together with the spectral characteristics of EH2, led to the realization that the second redox active group was a disulfide. In the reaction of EH2 with arsenite the spectrum returns to that of oxidized enzyme since the charge-transfer donor is taken into the arsenite complex. The exact chemistry of this reaction has been disputed,138 but this does not affect the conclusion that arsenite inhibits only prereduced enzyme.18

3. Oxidation-Reduction Potentials of £H2 The redox potential, E2, for the couple oxidized enzyme/EH2, measured by equilibration with lipoamide/dihydrolipoamide mixtures or with mixtures of oxidized and reduced azine dyes, is -280 mV at pH 7. The redox potential, E1; for the couple EH2/EH4, measured from the extent of dismutation of EH2 to mixtures with oxidized enzyme and EH4, is - 346 mV at pH 7.31 Thus, the relative potentials explain the stability of EH2 in the presence of excess substrates at neutral pH; the redox potential of the NADVNADH couple is -315 mV (per corrections in Clark, p. 490) and that of lipoamide/dihydrolipoamide is -287 mV 139-141 Because the pH dependence of E, is -60 mV/pH unit from pH 5.5 to pH 7.6 and that of NAD+/NADH is -30 mV/pH unit, reduction of EH2 to EH4 becomes more facile at low pH as shown in Figure 10.3I AE2/ApH is also -60 mV; these slopes show that two protons, as well as two electrons, are involved in the reductions at neutral pH. This is discussed further in Section III.F below. Another consequence of the relative potentials is that the EH2-NAD+ complex, described above, is stable at low pH. The effect of NAD + on the redox potential at pH 6 has been estimated.142 C. KINETICS OF THE FORWARD AND REVERSE REACTIONS 1. Presteady-State Kinetics The limiting rate of reduction of pig heart lipoamide dehydrogenase by dihydrolipoamide as monitored by the appearance of the charge-transfer band (555 s~ ! ), is equal to the steadystate turnover number (575 s"1) at 25°C, pH 7.6.13 This indicates that this half reaction is rate limiting in catalysis, and that species at the EH2 reduction level are the intermediates. As expected then, reoxidation of EH2 by NAD + is very rapid (>800 s"1).13 The question of whether the charge-transfer complex per se is an intermediate in catalysis has not been addressed directly. By analogy with thioredoxin reductase, where charge-transfer is seen only under special conditions, it would seem that the charge-transfer complex may not be a necessary intermediate. In the reverse reaction, reduction of the oxidized enzyme to EH2 by excess NADH is too rapid for accurate measurement (>800 s"1) and even at equimolar NADH, the pseudo-first-order rate is 140 s'1 at 25°C, pH 6.3.13 Thus, the reduced flavin

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 10. Effect of pH on the oxidation-reduction potentials of lipoamide dehydrogenase. (O), data obtained from titrations of enzyme with dihydrolipoamide; (•), value obtained from titrations of EH2 with lipoamide; (0), data obtained from titrations of phenosafranine and enzyme with dithionite; (4), value obtained from titration of EH2 and reduced phenosafranine with potassium ferricyanide; (V), data obtained from titrations of Eox and safranine T with dithionite. (n), theoretical points for E! obtained from determined values for E2 interpolated from the solid line at that pH and the relationship E2 - E, - (29.6)log K, where K - [EH2]2/[EH4][EOX]. The dashed lines are curves relating Em to pH for the NADH/NAD+ couple (9); and the dihydrolipoamide/lipoamide couple (7). (From Matthews, R. G. and Williams, C. H., Jr., 7. Biol Chem., 251, 3956, 1976. With permission.)

intermediate, E-FADH~ in the scheme of this reductive half reaction (Figure 11) has not been observed. The limiting rate of reoxidation of EH2 by lipoamide is approximately 800 s ~ ' . This rate cannot be compared directly with the steady-state turnover in the direction of lipoamide reduction (Table 2) due to the complex role of NAD + in increasing the concentration of EH2 in the steady state (Section HI. G.I). 135136 The pH dependence of the limiting rates of reduction of oxidized enzyme by dihydrolipoamide and of reoxidation of EH2 by lipoamide have been determined.136'143 The former is 830 s" 1 and is independent of pH from 5.5 to 8.0. The latter rate is 880 s"1 at pH 6.2

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FIGURE 11. Reduction of glutathione reductase by NADPH or lipoamide dehydrogenase by NADH. PNH and PN+, reduced and oxidized pyridine nucleotide, respectively; E and EH", oxidized and two-electron reduced enzyme, respectively. (From Matthews, R. G., Ballou, D. P., and Williams, C. H., Jr., J. Biol. Chem,, 254, 4974, 1979. With permission.)

TABLE 2 Kinetic Parameters for Mammalian Lipoamide Dehydrogenase Kinetic parameter

Rat liver enzyme

Pig heart enzyme

Yeast

490 nM 520 |xM 345

300 \LM 200 |xM 550

700 M-M 400 \LM 430

62 \LM

—

840 M-Af 124 37°C, pH 8.0 24

— 530 25°C, pH 7.6 13, 136

30M-A* — — 25°C, pH 7.6 144, 145

Forward reaction K-dihydrolipoamide

KNAD

Turnover number3 Reverse reaction KNADH *^lipoamide

Turnover number Assay conditions Reference a

3

Moles substrate x mole FAD"1 x sec"1

and is dependent on the protonation of a residue on the EH2-lipoamide complex with pKa of 7.9. It is argued that the residue affecting the rate of reoxidation of EH2 is the base. Further discussion of these results is in Section III.F below. 2. Steady-State Kinetics Mammalian and yeast lipoamide dehydrogenase show parallel line reciprocal plots when [NAD+] is varied at several levels of dihydrolipoamide,13-24'144'145 and the same pattern is observed (with the pig heart enzyme) in the reverse reaction at pH 8 where reduction of EH2 to EH4 is minimal (see Section III.G.l).13-24 The rate of isotope exchange and the lack of effect of lipoamide on that rate together with product inhibition patterns have been determined.24 These results are consistent with a bi bi ping-pong mechanism.13'24 The kinetic constants are summarized in Table 2. The pH dependence of the turnover numbers in the forward and reverse reactions reveals pKa values at 6.3 and 7.9 respectively.136 NAD + plays a unique role in the reverse reaction in addition to being a product. In the absence of added NAD + , the oxidation of NADH by lipoamide shows a pronounced, pH dependent lag. 18 - 145 ~ 147 The maximum stimulation is shown at the pH optimum of the reaction, ^ 3 135,147 However, NAD + does not effect the rate of reoxidation of EH2 by lipoamide and thus can not be exerting its effect by increasing the affinity of EH2 for lipoamide. Rather, NAD + increases the [EH2] in the steady state by reoxidation of EH4; NADH over-reduces the enzyme at low pH (see Section III.G.I). 135 Figure 12 gives a scheme for the kinetic mechanism of lipoamide dehydrogenase.

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 12. Model depicting the catalytic cycle and dead-end complexes in catalysis by Hpoamide dehydrogenase. A, B, P, and Q are NADH, Hpoamide (Lip(S)2NH2), NAD, and dihydrolipoamide (Lip(SH)2NH2), respectively. (From Wilkinson, K. D. and Williams, C. H., Jr., /. Biol. Chem., 256, 2307, 1981. With permission.)

3. Alternate Electron Acceptors The enzyme later identified as Hpoamide dehydrogenase was first purified as a so-called diaphorease, an enzyme transferring electrons to an artificial acceptor such as dichlorophenolindophenol or ferricyanide.148 It was shown that the low diaphorase activity of the native enzyme could be greatly enhanced and the natural activity virtually eliminated by treatment with Cu(II).149"151 Native Hpoamide dehydrogenase also transfers electrons to oxygen very slowly. The oxidase activity is enhanced by potassium iodide or ammonium sulfate and under conditions of high salt the pH optimum is 5.5.152 Other chemical modifications may lead to alterations in the ratios of activities.147'152"156 D. DISTINCT FUNCTIONS OF THE NASCENT THIOLS The two nascent thiols produced in the reduction of pig heart Hpoamide dehydrogenase by substrate are Cys-45 and Cys-50 (Figure 5). Their distinct functions as shown in Figure 2 are known.23'34'46-47-86 Cys-50 (upper thiol in Figure 2) interacts directly with the FAD; Cys-45 (lower thiol in Figure 2) interchanges with the dithiol substrate. Similar results are seen with glutathione reductase (Figure 3))33'34.37,39,78 an(j mercuric reductase (Section VI.C.I). 48 The interchange thiol is probably also the site of reaction with maleimide spin labels.153 The functional assignments are based on chemical modification studies. The two thiol moieties exhibit widely different reactivities toward iodoacetamide, allowing a monolabeled derivative of EH2 to be prepared which is homogeneously alkylated on Cys-45.46 Alkylation involves the loss of a proton from EH2 and thus, the monolabeled derivative is given the designation EHR.47 Alkylation disrupts the charge-transfer interaction in EH2, and EHR

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FIGURE 13. Spectral changes induced by titration of EHR with NAD + . EHR (curve 1) was mixed with increasing concentrations of NAD+ (curves 2 to 6 respectively). Equal concentrations of NAD + were added to the reference cuvette. The dashed line is the spectrum extrapolated to infinite NAD + by using the 1/Ae448 intercept obtained from a double reciprocal plot (inset). The dotted line is an estimate of the spectrum of the modified flavin species after correction from the contribution of residual unmodified flavin. (From Thorpe, C. and Williams, C. H., Jr., Biochemistry, 20, 1507, 1981. With permission.)

exhibits a spectrum of oxidized, bound flavin blue shifted by 7 nm. Although alkylation on Cys-45 interferes with normal charge-transfer, it is clear from several properties of EHR that Cys-50 still interacts with the FAD. Charge-transfer is restored by the addition of 3aminopyridine adenine dinucleotide (AAD + ) a nonreducible NAD + derivative. Furthermore, a covalent interaction between Cys-50 of EHR and the FAD is induced by NAD + ; this will be covered in Section III.E. While EHR is a two-electron reduced derivative, it is air stable and inactive toward lipoamide substrates, since alkylation of a single residue prevents reformation of the disulfide bridge. The FAD of EHR, as expected, is fully reduced by 1 mol of dithionite per FAD or by excess NADH. EHR retains a catalytically competent binding site for pyridine nucleotides as indicated by its interactions with NADH, AAD+ , and NAD+ and by the fact that it has transhydrogenase activity at least as high as that of the native enzyme.23-46-47 The alkylation studies just cited make it clear that it is Cys-50 that interacts with the FAD. Based on the fact that EHR was inactive with lipoamide, it was suggested that the other sulfur (Cys-45) participated in interchange with dihydrolipoamide. This was reasonable given that the more reactive thiol should be exposed for efficient interchange. However, this required the assumption that the two functions were carried out by different thiols; it was possible that alkylation of Cys-45 simply blocked interchange by the other thiol. It remained for the X-ray crystal structure, first of homologous glutathione reductase, and subsequently of lipoamide dehydrogenase to clarify this point showing clearly that Cys-45 is the interchange thiol (distal to the flavin) and to confirm that Cys-50 is the FAD interacting thiol (proximal).37-39-78-86 E. ELECTRON TRANSFER BETWEEN THE NASCENT DITHIOL AND FAD The spectral changes observed when NAD+ is added to two-electron reduced lipoamide dehydrogenase monoalkylated on Cys-45, EHR, suggest the formation of a covalent bond between the sulfur of Cys-50 and the C(4a) position of the flavin. Figure 13 shows the

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Chemistry and Biochemistry of Flavoenzymes

spectral changes during an NAD + titration.23'47 In addition to the partial bleaching of the 448 nm band there is an increase in absorbance at 390 nm. Extrapolation of the observed changes to infinite NAD + gives the dashed spectrum in which approximately half the absorbance at 448 nm has been lost. This suggested that the changes were occurring at only one of the two flavins in the dimeric protein. Indeed, binding studies using the gel filtration method,157 showed that 1.2 mol of NAD + were bound per 2 mol of FAD. The changes extrapolated to full formation on both monomers revealed an estimate of the spectrum of the adduct having a single band at 380 nm and an extinction coefficient of 7 mAf^crn" 1 . Comparison of this spectrum with that of model compounds of known structure suggests that it represents a covalent flavin adduct at the C(4a) position. The spectral changes are complete within 3 ms and the changes are fully reversible. The apparent K D for NAD + dissociation decreases from 370 |xM at pH 6.8 to 35 |xM at pH 8.8.47 The fact that covalent bond formation between the thiol and FAD is rapid and reversible suggests that the adduct could be an intermediate in the movement of electrons from the thiols to the FAD in catalysis. This is supported by the proximity of Cys-50 to the flavin C(4a) position.78'86 The scheme in Figure 4 gives a proposed mechanism for the formation and breakdown of this proposed intermediate in the native enzyme. The charge-transfer thiolate (upper) attacks the C(4a) position of the isoalloxazine ring to generate the covalent adduct. Nucleophilic reaction by the interchange thiolate completes the electron transfer reducing the flavin and regenerating the disulfide. The C(4a) adduct is not stabilized in the native enzyme and thus is not observed. However, with the interchange thiol alkylated, completion of electron transfer is blocked at the adduct. The covalent C(4a) adduct had also been proposed as a catalytic intermediate in lipoamide dehydrogenase on the basis of model studies. 158162 Similar spectral changes have been observed in thioredoxin reductase in which the FAD has been replaced by 1-deaza-FAD (Section V.B.3) and in mercuric reductase titrated with NADP+ at low pH (Section VI.D). 2021 1. Extreme Negative Cooperativity The extreme negative cooperativity or half-the-sites reactivity observed in the spectral changes induced in EHR by NAD + is perhaps reminiscent of the effect of NAD + on the spectrum of native lipoamide dehydrogenase.147 The data with EHR suggest that NAD + binding in one subunit induces adduct formation in that subunit and promotes changes in the other subunit which preclude tight binding of a second molecule of NAD + .47 Functional cooperativity in lipoamide dehydrogenase and glutathione reductase is reasonable in light of their structures which show intimate contract between the subunits, with elements of each active site drawn from both polypeptide chains.38'86 Similar spectral changes are induced by thionicotinamide adenine dinucleotide and nicotinamide hypoxanthine dinucleotide but not by acetylpyridine adenine dinucleotide or NADP + . In contrast, binding studies show that the quite distinct spectral changes brought about by the NAD + analogue, AAD + , namely charge-transfer, are associated with the binding of one molecule per FAD.47 Examples of extreme negative cooperativity have been observed in thioredoxin reductase in which the FAD has been replaced by 1-deaza-FAD (Section V.B.3) and in mercuric reductase under several conditions (Section VI. D).20'22-23 2. NAD + — An Effector in the Lipoamide Dehydrogenase Reaction The redistribution of electrons between dithiol and FAD in EHR may indicate an effector function of NAD + in addition to its substrate function as the electron acceptor in the forward reaction catalyzed by lipoamide dehydrogenase.47 A second line of evidence also suggests such an effector function. The spectral perturbation upon binding NAD + to native EH2 at low pH was mentioned in Section III.B.I. A spectrum similar to that of EH 2 -NAD + is observed transiently (in the 2 ms dead-time of the rapid reaction spectrophotometer) when

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FIGURE 14. Scheme for the reduction of lipoamide dehydrogenase by dihydrolipoamide. Species 1 is the oxidized enzyme with dihydrolipoamide bound and species 5 is EH2 with lipoamide bound. (From Matthews, R. G., Ballou, D. P., Thorpe, C., and Williams, C. H., Jr., J. Biol. Chem., 252, 3199, 1977. With permission.)

EH2 is mixed with nicotinamide hypoxanthine dinucleotide or acetylpyridine adenine dinucleotide at neutral pH.136 A similar spectrum is observed in about 12 ms upon reduction of the enzyme with reduced nicotinamide hypoxanthine dinucleotide at neutral pH. Detection of intermediates in the reduction of the enzyme by NADH or in the reoxidation of EH2 by NAD + are not possible with current technology since the reactions are very rapid. However, such intermediates have been detected with glutathione reductase and mercuric reductase.35'163 These results imply that the binding of oxidized pyridine nucleotide to EH2 should influence the equilibrium distribution of electrons between the flavin and the dithiol increasing the amount of reduced flavin.136 Structural studies have shown that the pyridinium ring of NADPH bound to glutathione reductase EH2, following reduction of the enzyme by excess NADPH, is oriented roughly parallel to the isoalloxazine ring of the flavin such that there should be good overlap of the TT orbitals.78-82 Assuming that NAD + binds to lipoamide dehydrogenase in a similar fashion, then the resulting stacked ring system might well be expected to have an altered redox potential. The relative redox potentials of disulfide/dithiol and FAD/FADH" are discussed in the next section;31-136-143 because the two electrons of EH2 are predominantly on the thiols the redox potential of disulfide/dithiol must be considerably more positive than that of FAD/ FADH". However, the binding of NAD + to EH2 at neutral pH results in an exchange of electrons between the two carriers and reduction of the NAD+ . F. ACID-BASE CHEMISTRY IN THE CATALYTIC CYCLE The presence of a base in the active site is supported by two lines of evidence. First, the oxidation-reduction potential of the EOX/EH2 couple varies by 60 mV/pH unit showing that the reduction involves two protons as well as two electrons. Since EH2 is a chargetransfer complex in which the donor is a thiolate, only one proton can be shared between the two thiols. Thus, the second proton must be taken up by another residue — the base.31-143 The base is His-451 hydrogen bonded to Glu-456 (Table l).51-86 Second, the rate of reduction of lipoamide dehydrogenase by dihydrolipoamide is independent of pH over the range of pH from 5.5 to 8.1. The invariance of the rate of reduction at pH values above 5.5 indicated that the base has a pKa below 5.5 in the oxidized enzyme and that above this pKa the base can deprotonate dihydrolipoamide making it a good nucleophile in thiol-disulfide interchange.136-143 The pKa values of the thiols of dihydrolipoamide are 9.35 and 10.65.143 The reductive half reaction can be pictured as in the scheme of Figure 14 where species 1 is oxidized enzyme with dihydrolipoamide bound and species 5 is EH2 with lipoamide bound. The two protons are shared between three groups at each stage as required for activation of the successive thiol-disulfide interchanges.143

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 15. (a) Scheme for the deprotonation of EH3+ to prototropic tautomers of EH2. (b) Scheme for the protonation and deprotonation of the ion-pair in papain. (From Sahlman, L. and Williams, C H., Jr., J. Biol. Chem., 264, 8033, 1989. With permission.)

EH2 is an equilibrium mixture between several species (Figure 2). The evidence for this will be more fully developed in Section III.G. Two of these species (3 and 4, Figure 2) are shown in the scheme of Figure 15a indicating that each thiol can exist as a thiolate. The third prototropic tautomer, with the base deprotonated, is not thought to be favored over the thiolate-containing species. This is based on analogy with work on papain and glyceraldehyde-3-phosphate dehydrogenase where the thiolate-imidazolium ion-pair is favored over the thiol-imidazole in an apolar active site. In these enzymes the pKa of the thiol is lowered and the pKa of the imidazole is raised; for example in papain, Cys-25 has a pKa of 3.3 and His-159 has a pKa of 8.5 as shown in the scheme of Figure 15b.164>165 The left-hand species in Figure 15a is the charge-transfer complex and is presumably favored in the equilibrium. The right-hand species is the form of the enzyme that reacts with iodoacetamide or with lipoamide in the reverse reaction. The pH dependence of the rate of mono-alkylation of EH2 reveals a pKa of 7.9 which has been ascribed to the base. This same deprotonation has been shown to govern the maximal rate of reoxidation of EH2 by lipoamide.143 Thus, the pKa of the base shifts upward in EH2 relative to Eox. The association of macroscopic pKa values with the ionization of specific groups in the protein must be regarded as a working hypothesis since a macroscopic pKa value, determined by a specific method, may reflect the ionization of more than one group. The importance of this caveat will become clear below. The pKa values (Section III.C and this section) are summarized in Table 3. The charge-transfer band is a valuable reporter group of the ionization state of the electron transfer thiolate. Macroscopic pKa values associated with the influence of pH on the visible spectrum of two-electron reduced pig heart lipoamide dehydrogenase and of yeast glutathione reductase have been determined by monitoring changes in the principal flavin band near 460 nm and of the charge-transfer band at 540 nm.31-33-166 In these experiments,

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TABLE 3 Pig Heart Lipoamide Dehydrogenase pKa Values Experiment Charge transfer (e530 nm) Charge transfer (€455 nm) Vmax (reverse) Vmax/KLipoamide

Vmax (forward) K-DHLA

Rate of alkylation Redox potential

Enzyme species

EH2 EH2 EH2*Lipoamide

EH2

EH2-NAD+

EH2 EH2 EH2 — EH-

pK,

PK2

4.4 3.9 — — —

— 7.0 —

— — —

— 6.3 5.2 — —

pK3 — — 7.9 8.3 — 8.4 8.0 >7.6

PK4

Ref.

8.7 9.3 —

166 166 136 143 136 143 143 31

— — — — —

the enzyme is reduced to the EH2 stage anaerobically and mixed in the rapid reaction spectrophotometer with anaerobic buffers having a range of pH values. The ionization of at least four active site amino acid side chains can influence the spectrum of EH2 over the range of pH studied: the two active site thiols and the His-Glu diad. Other groups, such as the two ion-pairs proposed as moderators of the flavin redox potential,78 are ignored in the analysis and this is an obvious over-simplification. As the pH is decreased, the extinction of the charge-transfer band diminishes until, at pH 3.6, the spectrum is essentially that of the oxidized enzyme.166 The changes take place in two distinct phases; in the upper pH range there is an isosbestic point at 499 nm, while in the lower pH range isosbestic points at 432 and 486 nm characterize the titration. This suggests that three species are involved in the titration and this is confirmed by two pKa values, at 4.4 and 8.7. By analogy with the less complex papain system, the pKa at 4.4 has been associated with ionization of Cys-50 and the pKa at 8.7 with His-451, more properly with the diad of His-451 and GIu-456. Recalling the caveat made above concerning assignment of pKa values to specific groups, it should be pointed out that, while the charge-transfer thiol is being directly monitored, it shares a single proton with Cys-45 (Figure 15a). Therefore, the ionization of the two active site thiols is probably reflected in the pKa at 4.4. The pKa associated with the base (8.7) is significantly higher than that determined from the rate of reoxidation of EH2 by lipoamide or from the alkylation of EH2 (8.0). The situation is made more complex when the data from the flavin band at 455 nm are considered. Here the results are better fit to three pKa values: 3.9, 7.0, and 9.3. Clearly, the flavin band reflects the titration of a somewhat different set of residues than does the charge-transfer band. It seems safe to conclude only that the lower pKa values reflect ionization of the thiol(s) and that the upper values reflect the base, but the microscopic pKa values of these residues are not obtainable from these data at present. These broad assignments are made clearer by work on monoalkylated glutathione reductase (EHR) (Section IV.D). G. SPECIAL PROPERTIES OF THE E. COLI ENZYME Lipoamide dehydrogenase from E. coli has been studied in some detail. This is of particular interest both for the understanding it gives of species at the EH2 level in addition to the charge-transfer complex, and because it is amenable to site-directed mutagenesis.167~171 The enzyme reduced by one mol of dithionite per FAD is a pH-dependent mixture of at least five species — three of these are at the EH2 level plus the oxidized enzyme and a four-electron reduced enzyme.167 This is a consequence of small changes in several properties from those of the pig heart enzyme and is not thought to result in a change of the mechanism (Figure 2). 1. Factors Affecting the Amount of Charge-Transfer Complex Reduction of the E. coli enzyme with excess substrates leads to almost complete 4electron reduction even at pH 7.5 (Figure 16), conditions which lead to no reduction beyond

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 16. Scheme for the reduction of lipoamide dehydrogenase by NADH or dihydrolipoamide. Physiological catalysis proceeds counterclockwise from E in the upper triangle. The numbers under the species refer to the same species in Figure 2; species 4 is the charge transfer complex and species 5 is the C(4a)-adduct.

the EH2 stage with the pig heart enzyme.172 However, when the E. coli enzyme is reduced with excess dihydrolipoamide in the rapid reaction spectrophotometer, the charge-transfer complex is fully formed in the 3 ms dead time; this intermediate decays slowly to the fully reduced enzyme.6 Moreover, titrations with up to 4 mol of dihydrolipoamide show that some charge-transfer complex is stabilized at equilibrium.173 Careful analysis of both absorbant and fluorescent changes shows that significant amounts of two species are in equilibrium with the charge-transfer species in reductive titrations of the E. coli enzyme. The scheme shown in Figure 17 shows the proposed structures of five enzyme forms at the EH2 reduction level.167 The three forms in the central column predominate at neutral pH where the central form is the charge-transfer species which is nonfluorescent and favored by high pH. The upper form is now pictured as the prototropic tautomer of the charge-transfer complex; it has 68% of the fluorescence of the oxidized enzyme and is favored by low pH. The lower form of EH2 in the central column has the electrons on the FAD; it is somewhat favored by low pH. All three of these forms are presumed intermediates in catalysis (Figure 16). The scheme in Figure 17 is based on data from reductive titrations from pH 5.8 to 8.0 in which the data from the linear portions of plots of fluorescence and e 530nm vs. reductant were extrapolated to an intersection to correct for dismutation and to give the properties of fully formed EH2. Plots of extrapolated fluorescence vs. extrapolated € 530nm> and extrapolated e 455nm vs. extrapolated e 530nm , for the experiments across the range of pH have two linear portions, suggesting that more than two spectrally distinct species are in equilibrium. This is shown in Figure 18a where any pair of vertically aligned points represents data from a reductive titration at a given pH. Extrapolation of the lower data set to 0 e530nm gives the fluorescence of the upper species in the scheme (68%) while extrapolation to 0 fluorescence gives the e 530nm of the charge-transfer species (3300 M^cm" 1 , equal to that of the pig heart enzyme). Extrapolation of the upper data set to 0 e530 nm gives the € 455 nm of the upper species in the scheme (10,500 M~lcm~l) while extrapolation to an e530 ^ of 3300 M~lcm~l gives the e 455nm of the charge-transfer species (7,800 M^cm" 1 ). With these properties the proportion of each form contributing to the observed spectrum could be

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FIGURE 17. Model for the pH dependence of the spectroscopic properties of E. coli lipoamide dehydrogenase at the EH2 level. The three protonation states are represented by the columns labeled A, B, and C. The three spectrally distinct species are represented by rows and are labeled I, II, and III. The arrows of forms IIB and IIC signify the charge transfer interactions between the thiolates and the FAD. (From Wilkinson, K. D. and Williams, C. H., Jr., J. Biol. Chem., 254, 863, 1979. With permission.)

calculated.167 This analysis was clarified by similar titrations carried out in the presence of 0.2 M guanidinium chloride since the reduced flavin species is not significant under this condition; plots such as those in Figure 18a are linear throughout (Figure 18b).168 Thus, in 0.2 Af guanidinium chloride, the charge-transfer complex and the fluorescent species are the only EH2 forms present. This nondenaturing concentration of guanidinium chloride was known to perturb the absorption spectrum, catalytic activities, and the reactivity of a single thiol in oxidized pig heart lipoamide dehydrogenase.174 The presence of the two additional EH2 species is the first of two reasons for the observation of less charge-transfer complex in reductive titrations of the E. coli enzyme. The other major difference between the E. coli and pig heart lipoamide dehydrogenases has to do with the redox potentials, E2 and El (as defined for the pig heart enzyme, Section B.3) — both their absolute values and the difference between them. E2 is — 264 mV and -280 mV, and E L is -317 mV and -346 mV for the E. coli and pig heart enzymes, respectively, at pH 7.31-167 Thus, both potentials are more positive in the E. coli enzyme and the difference between them is less, making the E. coli enzyme more easily reducible especially to the 4-electron stage and further accounting for the diminished charge-transfer complex at equilibrium. The slopes of plots of these midpoint potentials as a function of pH are 60 mV/pH unit as they are in the pig heart enzyme showing that two protons as well as two electrons are involved. The kinetic manifestation of over-reduction of the E. coli enzyme is product inhibition by NADH (Figure 12), and it has been suggested that this may have physiological significance in control of the bacterial enzyme where the phosphorylation/dephosphorylation control mechanism of the mammalian enzyme is absent.89-169 As shown in the scheme of Figure 12, NADH inhibits both by reducing EH2 to EH4 and by binding to EH2. This inhibition is relieved by NAD + but only partially since NAD + also inhibits by binding both to oxidized

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 18. (a) Relationship of fluorescence and 455 nm extinction to 530 nm extinction of E. coli lipoamide dehydrogenase at the EH2 level. Each pair of values at 530 nm extinction represented a dithionite titration at a different pH. The pH values were from left to right: 5.8, 6.3, 6.5, 7.1, 7.6, and 8.0; (A), fluorescence of EH2 relative to Eox vs. the 530 nm extinction of EH2; (O), 455 nm extinction of EH2 vs. the 530 nm extinction of EH2. (b) Relationship of fluorescence and e455 to the e530 of E. coli lipoamide dehydrogenase at the EH2 level in the presence of 0.2 M guanidinium chloride. Each pair of data points vertically aligned was determined in a separate dithionite titration; the pH values were from left to right, 5.7, 6.1, 6.5, 7.1, and 7.6. The relevant values were determined for the fully formed EH2; e453 (O), and fluorescence relative to the oxidized enzyme (A) were plotted versus the €530. (From Wilkinson, K. D. and Williams, C. H., Jr., J. Biol. Chem,, 254, 852, 1979 and Wilkinson, K. D. and Williams, C. H., Jr., J. Biol. Chem., 254, 863, 1979. With permission.)

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enzyme and to EH2. From the steady-state rate equations, it is apparent that the degree of inhibition will depend on the ratio of NADH to NAD + and on the redox potential, E } . Therefore, the wide variation in the severity of NADH inhibition between the E. coli and pig heart enzymes is explained by quantitative differences leaving the basic lipoamide dehydrogenase mechanism unchanged. 2. Kinetics of the Forward Reaction The kinetics of the reaction catalyzed by the E. coli enzyme in the forward direction are technically very difficult to measure because of the severe product inhibition, and because the reaction is very fast.172 NAD + + dihydrolipoamide ±5 NADH + H + + lipoamide An enzyme assay using 3-acetyl pyridine adenine dinucleotide was developed for monitoring purification, but it is not a physiological substrate, nor does it imitate the binding properties of NAD + . 173 Therefore, kinetic parameters for the forward reaction were measured in the rapid-reaction spectrophotometer with 30 nM enzyme FAD and 100 fiM dihydrolipoamide from pH 5.5 to 8.O.170 (The apparent KM for dihydrolipoamide was determined to be 12 (juM at pH 5.78 with a constant concentration of NAD + of 4.2 mAf, which later was shown to be four times KM for NAD + at that pH. Correcting for the NAD + concentration, the KM for dihydrolipoamide was 16 \LM.) The data were best fit using the equation: v = Vmax([S]/KM + [S]2/aK2M)/(l + 2[S]/KM + [S]2/aK2M) in which it is assumed that the binding of substrate to one subunit changes the dissociation constant for substrate in the second subunit with a factor a, that is, there exists cooperative binding.175 The Hill coefficient of 1.1 to 1.4 indicated that there was only modest positive cooperativity. The KM(NAD)+ was 230 |xM and kcat was 420 s" 1 at pH 7.5.170 The kinetics of the E. coli enzyme in the forward direction can also be estimated by conventional spectrophotometry at pH 7.6. The value of Vmax was 770 s"1 — higher than that obtained by rapid mixing. The KM for dihydrolipoamide was 70 to 80 (JiAf and for NAD + , 390 ^A/.176 The KM values for dihydrolipoamide differed by fivefold in the two methods; both values are much lower than those determined for the eukaryotic enzymes (Table 2). The pH dependence of kcat revealed an apparent pKa of 6.7 on the EH 2 -NAD + complex and the pH dependence of kcat/KM(NAD+) gave an apparent pKa of 7.4 on the free EH2.170 The kinetics determined over a range of NADH concentration showed a mixed pattern as would be expected given that this reaction product also reduces active EH2 to inactive EH4. The percentage of enzyme in the EH4 state, calculated for each combination of NADH and NAD + (using data of Reference 167) relating E^ (the oxidation-reduction potential of the EH2/EH4 couple) to the potential of the NADH/NAD+ couple, varied from less than 11% at pH 7.55 to as high as 50% at pH 6.5.17° Experiments in which two-electron reduced enzyme was mixed rapidly with buffers of various pH values revealed pKa values of 6.4 and 7.1 for data taken at 529 and 448 nm, respectively.170 While the data could not be analyzed at the extreme pH values, it was obviously different from the pig heart enzyme. The pKa of 6.4 was much higher than 4.35 observed with the pig heart enzyme. The higher pKa suggested a less tight interaction between the charge-transfer thiolate and the imidazolium ions in the E. coli enzyme compared with the pig heart lipoamide dehydrogenase. However, it was also noted that a pKa below pH 5.8 would not have been detected in the E. coli enzyme since at that pH the charge-transfer species which was being monitored constitutes only 23% of the EH2 and that percentage decreases as the pH decreases.167 The pKa of 6.4 is somewhat higher than that of 5.7 obtained

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Chemistry and Biochemistry of Flavoenzymes

in the earlier study, 167 It seems likely that this is due to the differences in the methods used. In the later study, extinctions observed 1 to 9 s after a pH change were used while the earlier work used extinctions extrapolated to full EH2 obtained in protracted dithionite titrations at various pH values. A second (higher) pKa of 8.2 was estimated but this was outside the range of the actual measurements.167 Macroscopic pKa values determined from the kinetics of the forward reaction and from changes in the charge-transfer absorbance all lie in the neutral range of 6.4 to 7.4 in contrast to the much lower pKa ascribed to the thiols in the pig heart enzyme. It was suggested however, that these pKa values near 7.0 to a greater or lesser degree reflect the ionization of the nascent thiols.170 3. Site-Directed Mutagenesis The cloning of the Ipd gene coding for lipoamide dehydrogenase in E. coli has made site-directed mutagenesis of the enzyme possible.42 A split-gene technique for mutagenesis of the gene was developed in order to overcome the instability problems encountered when attempting to mutagenize the intact gene.171 The Ipd gene was dissected into two fragments which were separately subcloned into M13 vectors for in vitro mutagenesis followed by reconstitution in the vector, pJLA504, for over expression under the transcriptional control of the XPR and XPL promoters and a temperature-sensitive X represser.177 After thermoinduction E. coli cells transformed with the plasmid carrying the reconstituted gene contained 4 to 5 times more lipoamide dehydrogenase activity than is normally found in the wild-type organism. This strategy has been used to engineer a number of mutations.171-178 Mutations have been designed to effect three distinct types of changes: (1) remove one of the titratable residues from the active site (Cys-44 to Ser, Cys-49 to Ser and His-444' to Gin), (2) moderate the rate of the pyridine nucleotide half reaction (Lys-53 to Arg, Glu188 to Asp, the double mutation, and Ile-184 to Tyr), and (3) replace a residue with the residue present at the homologous position of glutathione reductase (Ile-184 to Tyr, and Ser-52 to Lys). The Ile-184 to Tyr mutation will also test the effect of that aromatic residue on the properties of the FAD. In the first group, the Cys-44 to Ser enzyme was red as isolated. This was fully expected since, with the interchange thiol present as Ser, the electron transfer thiol is present without the necessity of reduction, and so participates as the charge-transfer donor. Physiological activity could not be detected in this enzyme since the thiol interchange half reaction was interrupted, but as expected, it had normal diaphorase activity implying that the pyridine nucleotide half reaction was unaffected.179 These results were identical to those obtained with the analogous mutation of mercuric reductase.180 Enzymes in which the base, His-444', had been altered to Gin had approximately 0.4% activity.181 This was also as expected. Given that the pH of the assay is 1.7 units below the first pKa of dihydrolipoamide,143 it would be expected that the activity would be diminished by 50 to 100 times, since one function of the base is to deprotonate the substrate. A second postulated role of the base is the stabilization of the electron transfer thiol as a thiolate making it a better nucleophile for attack on the C(4a) position of the flavin during electron transfer. The observation of almost no charge-transfer stabilized at equilibrium, in contrast to the wild type enzyme, indicates that the base is also important in this latter role and probably accounts for the additional fivefold loss of activity.182 Glutathione reductase having the analogous mutation likewise has about 0.5% activity (Section IV.D).183 The side chain of Tyr-197 in glutathione reductase must move in order for NADPH to bind.78-82 The homologous residue in lipoamide dehydrogenase is He. The visible band of glutathione reductase is red shifted relative to lipoamide dehydrogenase and the fluorescence of glutathione reductase is quenched compared to that of lipoamide dehydrogenase. In lipoamide dehydrogenase having Ile-184 changed to Tyr (I184Y) both properties take on the values associated with glutathione reductase.184 Forms of glutathione reductase in which

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TABLE 4 Absorbance Spectral Properties of Glutathione Reductase

Peak Peak Peak Peak a

b

c

to to to to

Enzyme source

Human erythrocytes

Yeast

E. co/i

t(mM~l cm"1)3 A1% (280 nm)b peak absorbance ratioc peak absorbance ratioc trough absorbance ratioc peak absorbance ratioc

10.5 [463 nm] 14.7 6.7 [280 nm/463 nm] 7.3 [274 nm/463 nm] 2.3 [463 nm/405 nm] 1.12 [463 nm/377 nm]

11.3 [460 nm] 15.4 8.5 [280 nm/462 nm] 8.9 [275 nm/462 nm] 2.3 [462 nm/405 nm] 1.14 [462 nm/380nm]

nd nd 7.5 [280 nm/462 nm] 8.1 [274 nm/462 nm] 2.45 [462 nm/406 nm] 1.12 [462 nm/380 nm]

The values are based on A1% at 463 nm = 2.01,190 and Mr = 52,400,54 as calculated for the human enzyme,186 and for the yeast enzyme, on comparison of the absorbance at 460 nm of enzyme bound FAD with the absorbance at 448 nm of the free FAD after removal from the enzyme using €448 am = 11.3 mM~ ! cm~' for free FAD.25 Protein was determined using the ultracentrifuge as a refractometer and in the case of the yeast enzyme, bovine albumin was the standard.190-191 Data for the human enzyme were taken from Reference 190 and are in fair agreement with earlier work;193 data for the yeast enzyme were averages for several preparations and were taken from Reference 192; and data for the E. coli enzyme were taken from Reference 196.

Tyr-177 is mutated to Phe, Ser, and Gly have quantum efficiencies ~~ 40% that of FAD in free solution and these levels are 25-fold higher than the wild-type enzyme.183 These results provide a very clear demonstration of what has long been an unproven assumption, that quenching of the flavin fluorescence commonly found with flavoproteins is due in large part to juxtaposition of the flavin to aromatic amino acid residues. The reduction of wild type lipoamide dehydrogenase by NADH to the EH2 stage is very fast and only the further reduction to EH4 can be observed in the rapid reaction spectrophotometer. Reduction of I184Y by NADH, on the other hand, can be resolved into three distinct phases: reduction of the FAD, k^ = 1000 s" 1 ; electron transfer to the disulfide, k2 = 50 s"1; and reduction to EH4, k3 — 3s" 1 . The scheme in Figure 16 shows the steps involved. The second phase, electron transfer to the disulfide, is shown as two steps. The fact that thiolate to C(4a)-FAD adduct is not observed, indicates that its breakdown is faster than its formation. Both these steps are very fast in the reverse direction as indicated by flavin reduction with dihydrolipoamide. The impressive effects of the He-184 to Tyr mutation hold out the hope that other mutations at this position will lead to changes that will allow further understanding of the NADH half-reaction.184

IV. GLUTATHIONE REDUCTASE Homogeneous glutathione reductase isolated from yeast, human erythrocytes, and E. coil has been studied in some detail. The X-ray crystal structure of the human enzyme was described in Section II.B.2 and the similarities with lipoamide dehydrogenase were touched on in Section II.A. Glutathione reductase from most sources is a dimer, the monomers having a Mr = 52,000. The enzyme from the Cyanobacterium, Spirulina maxima, apparently exists in a pH dependent dimer-tetramer equilibrium.185 The FAD is tightly bound to the yeast and E. coli enzymes, but the binding is less tight to the erythrocyte and liver enzymes, a property which has been exploited to replace the FAD with various flavin derivatives.186"188 Circular dichroism studies indicate that the structure of the apoenzyme is not altered significantly from that of the haloenzyme and that the apoenzyme reconstituted with FAD is identical with the native enzyme.189 The flavin fluorescence is largely quenched. The spectral properties of the enzyme are given in Table 4. Both values for the extinction coefficient near 463 nm are subject to errors inherent in the methods used.54-190'192 The somewhat lower 274 nm/463 nm ratio for the human enzyme is partially due to its lower

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Chemistry and Biochemistry of Flavoenzymes

Tyr content; correction following the method given in Reference 6, p. 101, the value of 7.3 would be 8.1 if the Tyr contents were equal. The higher value for the yeast enzyme may reflect its purity. A. METABOLIC ROLES AND DISTRIBUTION Glutathione reductase catalyzes the reduction of glutathione disulfide by NADPH: NADPH + GSSG + H + ±5 NADP + + 2GSH where GSSG is glutathione disulfide and GSH is glutathione, 7-glutamylcysteinylglycine. As already mentioned in the discussion of the crystal structure, the proton is probably donated by a fixed water molecule.38 The human enzyme is 4 to 6 times as active with NADPH as with NADH, 1 9 3 195 but the yeast and E. coli enzymes are quite specific for NADPH.28-196 20° The enzyme from erythrocytes, yeast, and liver is effectively retained on 2',5'-ADP-Sepharose 4B.201-203 In comparing activities, the ionic strength is crucial since there is activation by salt which is pronounced in the erythrocyte enzyme.193'195'204 Reported activation energies at high ionic strength vary from 11.1 to 14.4 kcal/mol;193"195 at low ionic strength there is a break in the Arrhenius plot at 17°C.195 A value of 16.7 kcal/mol can be calculated for the yeast enzyme from the data given in Section IV.C.2. The pH optimum for the reaction under standard assay conditions is 6.8 to 7.3 with NADPH and 6.2 to 6.4 with NADH for the erythrocyte enzyme,193-194-204 6.7 to 7.4 with NADPH for the yeast enzyme,25 and 7.3 for the E. coli enzyme.200'205 The oxidation-reduction potential of GSSG/2GSH couple is taken as -241 mV at pH 7 and 25°C. This number is an average of five determinations based on equilibrium with couples of known oxidation-reduction potential: NADP + /NADPH, two indicators and cystine/cysteine (Clark, p. 486).139'206-209 The latter values have been corrected using -222 mV as EQ for the cystine/cysteine couple rather than -350 mV which is incorrect.139'206 The value based on the NADP^/NADPH couple has been corrected using -327 mV as Eo,210 (I = 0.2) rather than - 320 mV.206 Two values have not been included in the average since possible artifacts in these studies have been cited. 194-206>211 The redox potential difference of 86 mV is equivalent to an equilibrium constant of approximately 800 M (AE£ = 29.61ogKeq):

This large redox potential difference and the normally high NADPH/NADP + ratio in the cell predicts that reduction of glutathione disulfide will be greatly favored, and indeed the estimated glutathione/glutathione disulfide ratio is high. It has been suggested however, that one reason that the reaction does not approach equilibrium in the cell is the fact that the K M for GSSG, 65 |xM, is much higher than the predicted equilibrium concentration of GSSG. Moreover, the high KM value is the basis for the putative roles of the GSSG concentration in many cellular processes.1'2 Another important factor influencing the glutathione disulfide/ glutathione ratio, in addition to the concentration of glutathione reductase, is the flux through reactions which utilize glutathione (see below).11 Glutathione reductase is a cytoplasmic enzyme. Since the NADPH/NADP+ ratio is high in the cytoplasm, the reasonable assumption has been made that the enzyme is present largely in the EH2 state.1-2 Because glutathione reductase maintains a high GSH/GSSG ratio in the cell, estimated to be in the range 20 to 1000, the metabolic roles of glutathione are the raison d'etre of the reductase. Glutathione is the specific substrate of the enzyme glutathione peroxidase (EC. 1.11.1.9), 212 of a broad group of detoxification enzymes referred to as thiol

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S-transferases (EC.2.5.1.18),213 and of several transhydrogenases such as microsomal glutathione-protein disulfide oxidoreductase (EC. 1.8.4.2).214-215 It is of interest that glutathione reductase is induced in rat liver by rrans-stilbene and other known inducers of drug-metabolizing enzymes.216 The less specific role of glutathione in maintaining the thiol/disulfide poise of the cell is effected directly in some cases and via glutaredoxin, a small protein containing a redox active disulfide, in others.]' The specificity of systems requiring reduction by a thiol is not clear; glutathione, reduced glutaredoxin or reduced thioredoxin are potential thiol or dithiol reductants (Section V.A). E. coli mutants having very low residual glutathione reductase activities maintain their glutathione pools largely in the reduced state, presumably via interchange, and thus, the enzyme is probably not essential.217 Glutathione in mitochondria, comprising 10 to 15% of cellular glutathione, is thought to originate in the cytoplasm.218 22° However, a mitochondrial and plastidic glutathione reductases have been reported.221'222 The glutathione disulfide binding site of erythrocyte glutathione reductase accommodates other substrates such as lipoate and the mixed disulfide of glutathione and coenzyme A (CoASSG).83 Both of these compounds are poor substrates for the erythrocyte enzyme.193-223 Evidence for a separate enzyme, specific for CoASSG and having a pH optimum of 5,5, has been presented, but this has been disputed.223"227 Glutathione reductase is widely but not universally distributed in aerobic organisms. Its absence in trypanosomatids is of interest since they contain trypanothione [TV1 ,A^-bis-(glutathionyl)spermidine] and trypanothione reductase (see Section VILA). Glutathione reductase has been isolated from many sources and these have been listed.2 The following may be added to this list: germinated peas,211 pea chloroplasts (Pisum sativum L),228 rice kernel,229'230 Thiobacillus thiooxidans,231 hepatopancreas of the sea mussle (Mytilus edulis L),232'233 the sea urchin egg,234-235 and the mycelium of Phycomyces blakesleeanus.236 Rather little is known about structural differences between glutathione reductases from various species. Not surprisingly, antibodies raised against the rat liver enzyme cross-react with the enzyme from other mammalian species (human and pig). However, there is no cross-reactivity with the enzyme from yeast, spinach or Rhodospirillum rubrum. The immunological determinant in the rat liver enzyme appears to be in the pyridine nucleotide domain.237 B. DISTINCT FUNCTIONS OF THE NASCENT THIOLS The two nascent thiols produced in the reduction of yeast glutathione reductase by substrate are Cys-58 and Cys-63 (Figure 5, using the numbering of the human enzyme). Their distinct functions as shown in Figure 3 are known.33-34-37'39 Cys-63 (upper thiol in Figure 3) interacts directly with the FAD; Cys-58 (lower thiol in Figure 3) interchanges with the disulfide substrate. Similar results were seen with lipoamide dehydrogenase (Section III.D),34'36 and mercuric reductase (Section VI.C.I). 48 The two thiol moieties in yeast glutathione reductase exhibit widely different reactivities toward iodoacetamide, allowing a monolabeled derivative of EH2 to be prepared which is homogeneously alkylated on Cys-58.33'34 Alkylation was assumed to involve the loss of a proton from EH2 as is the case with lipoamide dehydrogenase, and thus, the monolabeled derivative is given the designation EHR.47 While EHR is a two-electron reduced derivative, it is air stable and inactive toward glutathione disulfide. Alkylation of a single residue prevents both reformation of the disulfide bridge and interchange with glutathione disulfide. Interchange would have been detected because the charge-transfer interaction of EH2 (Section IV.C.5) is unaffected by alkylation, and formation of a mixed disulfide between Cys-63 and glutathione disulfide would have abolished charge transfer. EHR retains a catalytically competent binding site for pyridine nucleotides as indicated by its interactions with NADPH, AADP+, and NADP+ and by the fact that it has transhydrogenase activity at least as high as that of the native enzyme.33'34 The alkylation studies just cited make it clear that it is Cys-63 that interacts with the FAD. Based on the fact that EHR was inactive with glutathione

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Chemistry and Biochemistry of Flavoenzymes

disulfide, it was suggested that the alkylated sulfur normally participated in interchange with the disulfide substrate. This was reasonable given that the more reactive thiol should be exposed for efficient interchange. This required the assumption that the two functions were carried out by different thiols; it was possible that alkylation of Cys-58 simply blocked interchange by the other thiol. However, the X-ray crystal structure of human glutathione reductase had shown clearly that Cys-58 is the interchange thiol (distal to the FAD) and that Cys-63 is the FAD interacting thiol (proximal).37-40'78 C. CHARACTERISTICS OF TWO-ELECTRON REDUCED ENZYME The spectral properties of the oxidized enzyme are given in Table 4. Two-electron reduced glutathione reductase is an equilibrium mixture of several species. These are shown in Figure 3, species 2 to 8. The designation EH2 will again refer to the mixture which is stable anaerobically. The charge-transfer complex predominates with glutathione reductase. By analogy with lipoamide dehydrogenase, the donor is a nascent thiolate and the acceptor is FAD. The resonance Raman spectrum indicates that the FAD is oxidized in EH2*NADPH.238 A preliminary 13C-NMR study of A, vinelandii lipoamide dehydrogenase and erythrocyte glutathione reductase at the Eox, EH2, and EH4 levels detects signals from C(2), C(4), C(4a), and C(10a). The data indicate that the flavin in the EH2 forms of the two enzymes is strongly polarized and that higher IT electron density is stabilized at the C(4a) position in glutathione reductase than in lipoamide dehydrogenase. The data also show that in EH4 of both enzymes the flavin is anionic at pH 7.239 The oxidized enzyme is reduced rapidly by the product GSH (Section IV.C.5). The enzyme can be reduced to EH2 by NADPH and this process is very rapid and stoichiometric. The product is the charge-transfer complex but with NADP + bound in the case of stoichiometric NADPH and with NADPH bound if excess reductant is used.6-28 Complete reduction of EH2 to EH4 requires dithionite.6 1. Spectral Characteristics The spectrum of two-electron reduced yeast glutathione reductase is very similar to that of pig heart lipoamide dehydrogenase at the same reduction level.240 There are significant spectral differences between the enzyme reduced with excess NADPH and that reduced with excess NADH in that the charge-transfer band is more intense with the physiological substrate.197 Thus, EH2 forms a tight complex with NADPH - KD = 2.1 |xAf. The formation of this complex is very rapid;28 a KD of 11 |xM is reported for the E. coli enzyme.6 The enzyme from both sources forms a less tight complex between NADP + , and EH2 have lower absorbance at 540 nm and higher absorbance at 590 nm.6'197'241 With both the yeast and E. coli enzymes, treatment with stoichiometric NADPH in the presence of an excess of NADP + leads to the slow, partial conversion of EH2*NADP+ to the semiquinone anion, and the process is enhanced at high pH. 6>28>241 The semiquinone state must be readily accessible since glutathione reductase is reduced by the one-electron reductant methyl viologen and is reoxidized by one-electron acceptors such as ferricyanide and p-benzoquinones. Since the methylviologen and p-benzoquinone reactions are unaffected or stimulated respectively by NADP+ or its analogues, these electron carriers must react via another site (Section IV. E).242-243 2. Presteady-State Kinetics The reductive half-reaction of yeast glutathione reductase has been elegantly investigated.35 Three steps are spectrophotometrically distinguished using rapid reaction kinetics at 5°C, pH 7.6 (Figure 11). The first, complete in the dead-time of the apparatus (5 ms), is attributed to the formation of a charge-transfer complex in which NADPH is the donor and FAD is the acceptor. The spectrum of this species has a peak at 460 nm (e = 8.3m M ~ v cm"1), a shoulder at 490 nm and a broad peak centered at 650 nm (e = 1.0 m M~lcm~l). The second step, detected as an increase at 420 nm, has a limiting rate constant

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TABLE 5 Steady State Kinetic Parameters for Glutathione Reductase Kinetic parameter KNADPH KGSSG Turnover number1 Temperature Phosphate buffer Reference a

Erythrocyte

Erythrocyte

Yeast

13 \LM

9.5 |xM 19 (xM 128 25°C 0.03 M, pH 7.0 195

3.8 \LM 55 pM 250 25°C 0.1 M, pH7.6 25

125 IJLM 238 25°C 0.3 A/, pH 7.0 195

Yeast

£. coli

13 \LM

11 |xM 125 M-M 420 25°C 0.078M, pH7.6 192

125 \LM

33 5°C 0.1 M, pH 7.6 192

Moles substrate x mole FAD~' x sec~ f . Ping pong mechanism assumed.

of 153 s^ 1 showing saturation kinetics (KD = 8.3 jxAf). This step displays a kinetic isotope effect of 2.7 with (4S)-[2H]NADPH which indicated that C-H bond breakage in NADPH takes place at this step, presumably reducing the FAD. However, the increase at 420 nm and the lack of change at 460 nm, neither of which are consistent with flavin reduction, suggests that formation or breakdown of a C(4a) adduct was being observed (Figures 11 and 13), but neither of these would be expected to show an isotope effect. The third kinetically detectable step, forming the 540 nm absorbance characteristic of EH2, has a limiting rate constant of 68 s™ 1 and is thus rate limiting in the overall half-reaction. In this step, electron transfer from FADH~ to the disulfide is followed by NADP + dissociation and NADPH binding so that the final product is EH2*NADPH. The third step is associated with a kinetic isotope effect of 1.8 which is shown to be due to the fact that the second step becomes partially rate limiting when (4S)-[2H]NADPH is the substrate. It was suggested that, since charge-transfer interaction appears to precede hydride transfer, the transition state for flavin reduction involves extensive transfer of negative charge.35 The same suggestion could be made for lipoamide dehydrogenase operating in the direction of pyridine nucleotide reduction since the step preceding flavin reduction again involves a charge-transfer interaction. The presence of a kinetic isotope effect of 1.7 in the steady-state kinetics at 25°C, using (45)-[2H]-NADPH, led to the suggestion that the reductive half-reaction, and specifically enzyme disulfide reduction, was rate-limiting in the overall reaction,35 but this may not be the case. In a separate study of the yeast enzyme,6 the limiting rate of the reductive halfreaction was determined as 88 s ~ ] at 5°C, pH 7.6. The limiting steady-state turnover number was 33 mol substrate x s" 1 x mol FAD"1 which is distinctly slower than the rate of the reductive half-reaction determined in either study.635 The rate of reoxidation of EH2 (produced by borohydride reduction and anaerobic dialysis) was 48 s ~ l . Applying the equation for a bi bi ping-pong mechanism, Vmax = k3 x k7/(k3 + k7) (where k3 and k7 are the rates of the two half-reactions) to these data, the turnover number was calculated as 31 s" 1 in good agreement with the steady-state value. It was concluded that the enzyme followed this mechanism and that the reoxidation of EH2 was rate limiting.6 In any case, the much slower relative rate of the reductive half-reaction is a major difference between glutathione reductase and lipoamide dehydrogenase.35 3. Steady-State Kinetics The steady-state kinetic parameters, assuming a ping-pong mechanism, are given in Table 5 for glutathione reductase from three species. The dependence of KGSSG on ionic strength is of interest remembering that the predicted cellular equilibrium concentration of GSSG is less than K^^ and the measured range of [GSSG] is 5 to 200 jxM (Section IV.A).1-195 The assumption of a ping-pong mechanism holds except at high [GSSG]; the anomaly has led to the proposal of a branched mechanism.26'195-244'246 The kinetics of the reverse reaction, for which the pH optimum is about 8, have been determined.247-248 The kinetics of glutathione reductase in reverse micelles have been studied.249

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 19. Branching steady-state kinetic mechanism for glutathione reductase. Catalysis from E is counterclockwise for the ping-pong mechanism and clockwise for the sequential mechanism.

The structure of glutathione reductase, showing distinct binding sites for GSSG and NADPH,39 allows for complications in a simple ping-pong mechanism, which does not require independent sites. Thus, since GSSG can react with EH2-NADP+, EH2 or EH2«NADPH and since the relative concentrations of these enzyme species will vary with the substrate concentrations, purely kinetic descriptions such as ping-pong and ordered sequential are inadequate.6-28 It has been suggested that deviations from classical ping-pong kinetics are to be expected and precedents have been cited.28 The gene coding for E. coli glutathione reductase has been cloned, sequenced and overexpressed.250"253 The human gene has also been cloned and sequenced.254 The use of sitedirected mutagenesis of the E. coli enzyme, to change a key residue in the pyridine nucleotide binding pocket, has yielded very interesting results and, inevitably, further complications.183 In this study, Tyr-177 (homologous with Try-197 in the human enzyme) was changed to Phe (Y177F), Ser (Y177S), and Gly (Y177G). While the steady-state kinetics of Y177F were like those of wild-type enzyme (ping-pong), the kinetics of Y177S and Y177G were altered and could be analyzed as an ordered sequential mechanism. If, as seems highly likely, the rate of flavin reduction by bound NADPH has been significantly slowed, GSSG could bind to E-NADPH as in an ordered sequential mechanism as shown in the scheme of Figure 19. The limiting turnover number of Y177F was changed only slightly but the limiting turnover numbers of Y177S and Y177G were 137 and 4.7 s"1 respectively (cf. Table 5). Favoring of the ordered sequential mechanism was also shown to be a result of dramatic lowering of KGSSG in Y177S, 2 \LM, and Y177G, 5 |xM, compared to 53 jxM for wild-type enzyme.183 The binding of NADPH to glutathione reductase requires that the side chain of Tyr-197 swing through about 90°C.82 It has been suggested that this side chain protects the isoalloxazine ring from solvent and thereby from oxygen.78 This hypothesis was tested in the engineered enzymes just described by measuring the half-life of borohydride reduced enzyme. Since the reoxidation rates of Y177F, Y177S, and Y177G were similar and were only about 2.4 times as fast as the wild-type rate, the authors concluded that protection is derived more from the buried position of the flavin than from the nature of the side chain at position 177.183 Altering other carefully selected combinations of residues in the NADPH binding site decreases kcat/KM for NADPH 250-fold and raises kcat/KM for NADH 72-fold.255 The intrinsic, primary kinetic isotope effects for the spinach, yeast, and E. coli glutathione reductases were significantly different at pH 8.1: DV/KNADH = 4.3, 2.7, and 1.6 and

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T

V/K NADH = 7.4, 4.2, and 2.2, respectively. These were measured from steady-state kinetics and from rapid reaction kinetics of the reductive half-reaction. It was suggested that this reflects differences in the transition-state structures for the hydride transfer step as catalyzed by the three enzymes.256 Several synthetic glutathione disulfide analogues have been examined both as substrates and in their mode of binding by difference crystallography. The analogues had fewer charged groups than the natural substrate. Their Vmax/KM values ranged down from 18 to 0.004. It could be concluded that the carboxyl at Glu-I which is a direct salt bridge is less important for binding than that at Glu-II which is an indirect salt bridge, and that the amino group at Glu-I, an indirect salt bridge, is less important for binding than the amino group at Glu-II, a direct salt bridge.257 4. Inactivation of Glutathione Reductase by NADPH and NADH Glutathione reductase is inactivated by preincubation with its substrate NADPH. The phenomenon has received a good deal of attention due to the possibility that it could form the basis of metabolic control. The extent and rate of the inactivation as well as the reversibility of the process are different for the enzyme isolated from various sources. Moreover, the extent and rate of inactivation, and its reversibility, depend on the conditions for the enzyme from any one species. Inactivation of the erythrocyte enzyme in preincubations with NADPH was noted in early studies.193 Inactivation appeared to be associated with aggregation in the erythrocyte enzyme,204 but it is noted that inactivation by NADPH was studied at low enzyme concentration while aggregation promoted by NADPH was observed in the ultracentrifuge at 5 mg/ ml. Aggregation was also observed, but to a lesser extent, in the absence of NADPH, and thiols protected under this condition.258 Inactivation by 1 mM NADPH was maximal in 5 min with 50% activity remaining, and was completely reversed by GSH in approximately 10 min. Reversal by GSH led to the suggestion that inactivation involved intermolecular thiol oxidation.204 In this connection it should be recalled that the erythrocyte enzyme has two thiols not found in the E. coli enzyme, Cys-2 on a flexible arm not visualized in the crystal structure, and Cys-90 which forms a disulfide at the monomer-monomer interface.40'81 Some characteristics of the inactivation were measured as a lag time in the assay and thus were quantitated under conditions of regain of activity.204 The inactivation (lag time) was first order with respect to the GSSG in the assay but less than first order with respect to NADPH. It was suggested, in order to explain aggregation, that the latter result was consistent with NADPH reducing one active site while blocking another intermolecularly.204 The finding would also be consistent with extreme negative cooperativity within a single dimer molecule remembering that both lipoamide dehydrogenase, thioredoxin reductase, and mercuric reductase exhibit negative cooperativity under some conditions.20>22>23 Inactivation by NADPH and NADH has also been studied in the enzyme from yeast,259'260 andE. coli,261 and has been noted for the mouse liver enzyme.262 In general, the characteristics of the inactivations determined were similar to those seen with the erythrocyte enzyme, but given the species variability, each of these studies should be considered separately. Inactivation of 0.6 |iAf yeast glutathione reductase by 50 fiM NADPH or NADH was complete in 60 min with half-times of 8 and 10 min respectively in a biphasic process. Protection was afforded by high concentrations of NADP + , GSH, GSSG, and dithiothreitol. Substantial reactivation was observed with GSH and ferricyanide following removal of the NADPH. Partial reactivation by GSSG or dithiothreitol was shown. The time course of inactivation was similar to that of the loss of charge-transfer absorbance (530 nm) but only half the latter disappeared as the activity fell to 3.5%. It was proposed that the process involved the formation of an intramolecular disulfide between one of the active center thiols and another thiol. While it was indicated that the enzyme should be fully protected under average cellular

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Chemistry and Biochemistry of Flavoenzymes

conditions, it was suggested that variations in the GSSG concentration could lead to inactivation acting as a physiological control.259-260 A study of the inactivation of E. coli glutathione reductase by NADPH and NADH has shown that the process was inversely dependent on the enzyme concentration.200 The effect of protein concentration has been noted in an earlier publication.261 Inactivation was rapid and monophasic with 1 |xM NADPH and 1 nM enzyme FAD giving a t1/2 of 1 m i n - Complex formation between NADPH and the two-electron reduced enzyme (EH2) at higher levels of NADPH protected against rapid inactivation. NADP + , produced in a side reaction with oxygen, also protected by forming a complex with EH2. These complexes with NADP(H) made analysis of the concentration dependence of the inactivation process difficult.200 NADH and NAD + do not form significant complexes with EH2. The relationship between the final level of inactivation with NADH and the enzyme concentration indicated that inactivation was due to dissociation of the normally dimeric enzyme. The kinetics of the GSH promoted reactivation were second-order, monomer-monomer, over 75% of the reaction with an apparent association rate constant of 13 x 106 Af~ I min~ 1 . Thus, the position of the dimer-monomer equilibrium between an active dimeric two-electron reduced species and an inactive monomeric 2-electron reduced form appeared to determine the enzyme activity.200 Although it is difficult to envisage such a dimer-monomer equilibrium in light of the extensive noncovalent interaction between monomers revealed by the crystal structure, the possibility of dimer dissociation under cellular conditions of high GSH concentration has been suggested previously.40 Inability to separate monomer and dimer was ascribed to the inherent dynamics of gel chromatography when attempting to separate a kinetically controlled dimer-monomer system.263 The conditions promoting rapid, reversible and extensive inactivation of the E, coli enzyme are far removed from those present in the cell. At the concentration of glutathione reductase in £. coli (880 nM),196 and using the apparent KD of 220 nM determined for the dimer-monomer equilibrium, the enzyme would be less than 30% dissociated. Moreover, inactivation was quite slow at the NADPH concentration reported in bacterial cells (150 (jiM),264 thus it was concluded that it was unlikely that this phenomenon was significant as a physiological control.200 It should be recalled that an enzyme concentration dependent dimer-monomer equilibrium has been suggested for lipoamide dehydrogenase which loses normal (but not diaphorase) activity simply upon dilution; the same conditions lead to lowering of the molecular weight as estimated by light-scattering measurements.265-266 Conditions affecting the monomer-dimer equilibrium have been studied.267'268 The erythrocyte enzyme has a Cys residue at position 90 which forms a disulfide across the twofold symmetry axis between the subunits, at least in isolated oxidized enzyme.40 An alteration was engineered in E. coli glutathione reductase in which Thr-75 (at the homologous sequence position) was changed to Cys (T75C).269 It was shown that the T75C enzyme did not dissociate to monomer in sodium dodecyl sulfate as did the wild-type control unless reducing conditions were used, indicating that an intersubunit disulfide had formed in the mutant enzyme. Neither was any dissociation observed if the mutated enzyme was treated with NADPH, both at high concentration. It was also shown that both wild-type and T75C enzyme followed the same thermal stability curve with 50% activity remaining at ~ 37°C in the presence of a high concentration of NADPH and at ~ 67°C in its absence. Almost no inactivation by NADPH was seen at 25°C,269 and under these conditions (high NADPH concentration) none would have been predicted based on the earlier study.200 It will be of interest to see if the inactivation of T75C enzyme shows the same protein concentration dependence and if the reactivation gives the same monomer concentration dependence as does the wild-type enzyme, or if it gives the ambiguous dependence observed in this laboratory with the erythrocyte enzyme.192 Given that the disulfide and the base in the active sites of glutathione reductase and lipoamide dehydrogenase are derived from separate polypeptide chains (Section II.B.2),39'40'86

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dimer dissociation would be expected to lead to extensive inactivation of normal catalysis. Indeed, site directed mutagenesis of the base of either enzyme leads to greater than 99% inactivation (Section III.G.3). 181 - 183 Thus, the hypothesis for E. coli glutathione reductase, that dimer dissociation leads to the inactivation observed upon incubation with NADPH, is an attractive one. While it is consistent with the data for the E. coli enzyme, it is possibly at variance with the experiment in which the spectrum of the yeast enzyme was followed during treatment with NADPH with loss of only half the charge-transfer absorbance.260 It is possible however, that some charge-transfer occurs in the monomer even without stabilization by the base (following putative dissociation). A definite dependence on yeast enzyme concentration has been observed but its quantification is made difficult by rapid reactivation by NADP + formed in the assay, subsequent to preincubation with NADPH.192 In the erythrocyte enzyme, a different mechanism probably applies. 5. Redox Potential A plot of the redox potential of lipoamide dehydrogenase (EOX/EH2) vs. pH, from pH 5.5 to 7.6, has a slope of — 60 mV/pH unit showing that two protons as well as two electrons are involved in the reduction. Above pH 7.6 the data suggest a shift to a -30 mV slope which would apply to a one proton reaction i.e., E OX /EH~. 31 The redox potential of glutathione reductase has been determined from pH 5.5 to 9.8 using NADH in the presence of NAD + , 192 since, unlike the natural substrate, NADPH, NADH does not form complexes with either Eox or EH2 and reacts quickly.197-200 The redox potential at pH 7, 20°C is - 235 mV and the slope of plots of the potential as a function of pH changes from — 52 to — 39 mV/pH unit at pH 7.4. These slopes deviate significantly from those expected for a twoelectron reduction involving two protons (58 mV/pH, 20°C) or one proton (29 mV/pH, 20°C).139 However, below pH 7.4, the two-electron reduction involves essentially two protons (Eox to EH2). The deviation from theory at higher pH indicates the presence of other dissociable groups whose pKa values are linked to the redox state of the enzyme.32'139 Acetytlpyridine adenine dinucleotide has been used to confirm the NADH/NAD+ data; it reacted rather slowly but its potential was ideal being very close to that of the enzyme.192 Glutathione is another possible reductant, however, there may be a potentially interesting complication. Glutathione reductase in which the FAD has been replaced by 6-SCN-FAD was titrated with GSH. Treating the reaction as a binding and plotting l/[charge transfer] vs. 1/GSH (not 1/[GSH]2) gave a straight line suggesting that the reaction was blocked at the mixed disulfide stage (Figure 3).270 A similar observation has been made with enzyme containing FAD, even at pH values as high as 8.4 (Section IV.D).25'192 If it can be shown that the reaction is indeed blocked at the mixed disulfide, it should be possible to measure the redox potential of the EOX/ESSG couple. D. ACID-BASE CHEMISTRY IN THE CATALYTIC CYCLE 1. Studies Utilizing Absorption Spectral Properties Two-electron reduced glutathione reductase contains at least three interacting groups which are protonatable, the nascent thiols and the base (Figure 15a). This system is analogous to, but more complex than, those in glyceraldehyde-3-phosphate dehydrogenase and papain where a single thiol and a histidine residue in a relatively apolar milieu form a thiolateimidazolium ion-pair which is favored over the thiol-imidazole prototropic tautomer (Figure 15b). Thus, the activation of a potential nucleophile by a juxtaposed base is a common theme in enzymology. In an effort to more nearly mimic the papain titrations, the macroscopic pKa values were determined on yeast glutathione reductase that had been reduced and alkylated selectively on the interchange thiol by monitoring both the charge-transfer and the flavin absorbances (Figure 3). Like papain and glyceraldehyde-3-phosphate dehydrogenase, alkylated glutathione reductase showed two macroscopic pKa values, 3.7 and 9.1, and by

162

Chemistry and Biochemistry of Flavoenzymes

analogy these were associated primarily with the thiol (the charge-transfer donor) and the imidazole respectively.33-166 Results with the native enzymes understandably are more complex.166 Glutathione reductase had apparent pKa values at 4.8, 7.1 and 9.2. Lipoamide dehydrogenase had apparent pKa values at 3.9 to 4.4, 7.0, and 8.7 to 9.3. It seems safe to state on the basis of the papain analogy and the relative charge-transfer absorbance in the two titration phases, that the lowest macroscopic pKa in each case, reflects deprotonation of the thiol to an anion that can act as the donor in the charge-transfer complex, and that the highest macroscopic pKa observed in these enzymes reflects the ionization of the His residue. This conclusion is clear for alkylated glutathione reductase where the analogy to papain can be made and it is logical, too, for the native enzymes. Since the base is unprotonated in the oxidized enzyme, its pKa changes markedly upon reduction as is the case in lipoamide dehydrogenase. In the physiological direction then, it can serve as the base in the oxidation of dihydrolipoamide (Figure 2, species 1 to 2) and as the acid in the reduction of GSSG (Figure 3, species 7 to 8 and 8 to 1). The observed pKa values near pH 7 in native lipoamide dehydrogenase at 455 nm and in native glutathione reductase at 540 nm can not be assigned with any confidence as reflecting specific ionizations.166 The complexities of the system are made much clearer in a recent report which became available only after this chapter was written (see also Reference 7).271 Stabilization of Cys63, the flavin interacting thiol, as a thiolate, comes from its favorable position for hydrogen bonding with Thr-339-OGl (Table 1) and with the C2'-oxygen of the ribityl side chain as well as the charge-transfer interaction with the flavin. The position of Cys-63 in EH2 is not favorable for interaction with the base.271 The importance of the base has been further demonstrated by site directed mutagenesis of His-439' to a glutamine residue (H439Q).183 The activity of H439Q was found to be less than 1% that of wild type enzyme. As will be discussed shortly, the base serves to protonate the first dissociating glutathione thiolate preventing the unproductive nucleophilic attack on the mixed disulfide.27 Thus, this function confers more than 100-fold rate enhancement on catalysis. The expected magnitude of this enhancement has been discussed for lipoamide dehydrogenase (Section III.G.3). Mutation of His-439' lowered the KM for NADPH.183 The identity of alternate bases in H439Q has been explored in a subsequent study: Y99F had essentially full activity.272 2. Studies Utilizing Kinetic Properties The pKa values of groups whose ionization affects the catalytic activity have been determined for both the yeast and erythrocyte enzymes and these important studies allow a far more complete interpretation than was possible with the spectral data discussed above and the redox potential data (Section IV.C.4).27-273 Two pKa values were calculated from plots of log(V/KGSSG) vs. pH, 8.4 and 8.8 for the yeast enzyme and 7.8 and 8.6 for the erythrocyte enzyme. Activity was dependent on both groups being protonated. These were attributable to groups on free EH2 and free GSSG and it was suggested that the group having the higher pKa was the a-amino group of the 7-glutamyl moiety of GSSG and that a group on EH2 had the lower pKa. Two pKa values were calculated from plots of log(V) vs. pH, 6.2 and 9.2 for the yeast enzyme; from similar data for the erythrocyte enzyme, that covered the range of 6.6 to 10.8, one pKa of 9.0 was calculated. These pKa values were associated with groups on the EH2-GSSG complex and were attributed to the base (9.0 to 9.2) and to the interchange thiol (6.2).27-273 The pKa attributed to the base in the erythrocyte enzyme was seen to vary slightly with temperature, and the enthalpy of ionization was 3.2 kcal/mol. This value was low for the ionization of a histidine residue and it was noted that this emphasizes that the base is in fact the His-467'-Glu-472' diad. A pKa of 9.1 to 9.3 was also derived from plots of log(V) vs. pH where NADPH, 2',3'-cyclic-NADPH or NADH were the varied substrate with the erythrocyte enzyme.36

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Solvent kinetic isotope effects (SKIE) have been determined in order to detect possible rate-limiting protonation steps.27 SKIE were observed at pH 8 on V but not on V/K when either NADPH or GSSG was the varied substrate at a saturating level of the second substrate. However, when either of the mixed disulfides, glutathione~5-thio-2-nitrobenzoic acid or glutathione-2-thio-4-nitropyridine were used as the varied substrate, solvent kinetic isotope effects were observed on both V and V/K using the erythrocyte enzyme. These were shown to be good substrates with KM values of 70 \iM and 13 jutM, respectively, and maximal rates of 62% and 57% of the GSSG maximal rate, respectively. Since the SKIE seen with these substrates, 3.4 and 3.2, respectively, were similar to that observed with GSSG, 2.6, it was suggested that the same rate-limiting proton transfer was being observed. Further, it was argued that since a very similar solvent kinetic isotope effect of 2.5 was observed when NADPH was the varied substrate, this protonation was rate-limiting in the overall reaction. The identification of the rate-limiting step with the protonation of the first dissociating glutathione anion rests on two points. First, recalling the crystallographic definition of the GSSG subsites (Section II. B. 2), the mixed disulfide is formed in Site I and the GS" occupying Site II departs first.78 In catalysis with GSSG as substrate at pH 8, where it is known that the enzyme is in the EH~ state (Section IV.C.5),192 the high pKa of the glutathione thiol dictates that the first departing glutathione anion be protonated. This suggests, given the similarity of the SKIE, that the mixed disulfides are bound such that the glutathione half is in Site II and departs first as the thiol. This is supported by the fact that the inhibitor, S(2,4-dinitrophenyl)-glutathione binds in this orientation.274 Second, for the SKIE to be observed on V/K, the first thiol/disulfide interchange, in which the substrate disulfide is reduced, must be reversible. However, it was argued that if the mixed disulfides were bound in the opposite orientation, the proton-independent release of the thionitroaromatic function might be expected to be an irreversible step (since the thionitroaromatics are poor nucleophiles) and no SKIE on V/K would be seen.27 The schemes in Figure 20 give the steps in the first thiol/disulfide interchange for GSSG on the left and for the thionitroaromatics on the right.27 It was proposed that the rate-limiting step was the protonation of the first glutathione thiolate (k3 in Figure 20). The failure to observed a solvent kinetic isotope effect on V/K where GSSG was the variable substrate was explained logically on the basis of the partitioning of the thiolate between protonation (forward reaction) and nucleophilic attack on the mixed disulfide (back reaction). The high pKa of this thiolate favors protonation, that is the equilibrium in the back reaction is unfavorable (k3 ^> than k2). With the thionitroaromatic bound as shown on the right, a solvent kinetic isotope effect on V/K is observed presumably because k3 is approximately equal to k2. Table 6 summarizes all of the pKa values derived from the three types of data. Figure 21 shows all possible species given three ionizing residues, and the middle row contains the species thought to predominate at any given pH. Thus, the pKa values are: FAD interacting thiol (Cys-63) < interchange thiol (Cys-58) < base (His-467')- The lowest pKa is assigned to the thiol interacting with flavin on the basis of the development of charge-transfer as the pH of two-electron reduced enzyme is raised from three and on the basis of the papain analogy (Section III.F). In two-electron reduced enzyme monoalkylated on the interchange thiol (EHR) this assignment is clear. In the neutral pH range, pKa values are detected by all three methods (Table 6) and the redox potential data indicates that the EH2 to EH~ transition is in this region (Section IV.C.4). Finally, activity is lost and charge-transfer reaches a maximum as the base is deprotonated, and again, this is in conceit with the papain analogy. This interpretation indicates that EH2 and EH~ are catalytically active in contrast to lipoamide dehydrogenase where only EH2 is active (Section III.F). The scheme in Figure 3 is an Eox to EH2 to Eox mechanism. At high pH it would be modified by the second glutathione molecule dissociating as the thiolate. Failure of the redox potential experiments

164

Chemistry and Biochemistry of Flavoenzymes

FIGURE 20. Proposed mechanism of the first thiol-disulfide interchange, including the rate determining proton-transfer step, in the oxidative half-reaction of human glutathione reductase. Column on the left shows glutathione and the column on the right shows the asymmetric disulfide with its binding orientation. (From Wong, K. K., Vanoni, M. A., and Blanchard, J. S., Biochemistry, 27, 7091, 1988. With permission.)

TABLE 6 Yeast Glutathione Reductase pKa Values Experiment

Enzyme species

pKt

PK2

pK3

pK4

Ref.

Charge transfer complex absorbance Charge transfer complex absorbance V^ (vary GSSG, NADPH saturating) V^/KGSSO Redox potential

EH2 EHR

4.8 3.7 — — __

7.1 — 6.2

— — — 8.4 —

9.2 9.1 9.2 — —

166 166 273 273 192

EH2-GSSG

EH2 EH2 — EH-

— 7.4

to detect the pKa at 9.2 indicates that the model with only three dissociating species is too simple. E. INHIBITORS AND CHEMICAL MODIFICATION The finding of irreversible inhibition of reduced yeast glutathione reductase by 2-chloroethylisocyanate derived from the antimalerial-antineoplastic drugs of the l,3-bis(2-chloroethyl)-l-nitrosourea (BCNU) type has generated great interest in the possibility of chemotheropeutic intervention via this enzyme.275 The effects were presumed to be due to inhibition of the glutathione reductase/glutathione peroxidase antioxidant system.276 This presumption now has experimental support,277 including the novel observation that erythrocytes pretreated with BCNU to lower their glutathione reductase activity and transferred to a drug-free medium do not support the growth of the malarial parasite, Plasmodium falciparum, when their

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FIGURE 21. Forms of 2-electron reduced glutathione reductase as a function of pH.

glutathione reductase activity is low.277'279 A high resolution structure of the BCNU-modified erythrocyte enzyme shows clearly that the reagent reacts exclusively at the interchange thiol of EH2 giving rise to a S-carbamoyl derivative.280 The structure of the enzyme reacted with the related compound, l-(2-chloroethyl)-3-(2-hydroxyethyl)-l-nitrosourea is consistent with the presence of a 2-hydroxyethyl group on the interchange thiol and chemical evidence for this is referred to. Thus, the mechanism of reagent activation must be different with this reagent.280 The availability of NADPH is also a factor in the inhibition of glutathione reductase in v/vo.281'282 Mitomycin-C and 5-nitrofurans have been treated as reversible inhibitors of glutathione reductase.282'283 Perhaps the most interesting reversible inhibitor is 2,4,6-trinitro-benzenesulfonate.284-285 It inhibits the GSSG reductase activity and is itself reduced by four electrons. In the presence of oxygen it is reoxidized giving rise to an apparent oxidase activity. This oxidase activity is stimulated by NADP + as are all presumed one-electron acceptors including 2,6-dichlorophenolindophenol and ferricyanide.284'285 This stimulation by NADP+ suggests that these compounds interact with the enzyme at another site. Indeed, the azine dye safranin and menadione bind in the large interface cavity between Phe-78 and Phe-78', and His-75 and His-75', well away from the flavin.83 It was suggested that binding at this site could influence the active center disulfide via the helix formed by residues 63 to 80.40 Does this imply tunneling? If so, what better tunnel than a helix. A variety of other protein modifying reagents have been used to inactivate glutathione reductase. In some cases the site of reaction was suggested by the known reagent specificity but in no case has the residue(s) been identified in the sequence.286293 In some studies, the simultaneous loss of GSSG reductase activity and transhydrogenase activity and/or diaphorase activity was used to suggest the nature and location of the residue modified.286'290

V, THIOREDOXIN REDUCTASE A. METABOLIC ROLES AND DISTRIBUTION Thioredoxin reductase catalyzes the reduction of thioredoxin by NADPH:15 NADPH + H + + TR(S2) ±5 NADP + + TR(SH)2 where TR(S2) is thioredoxin and TR(SH)2 is reduced thioredoxin, a small (Mr = 12,000) protein of known structure containing a redox active disulfide.294'295 The flow of electrons

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Chemistry and Biochemistry of Flavoenzymes

in catalysis by thioredoxin reductase is the same as in glutathione reductase: from NADPH to the FAD, from reduced FAD to the active site disulfide, and finally from the active site dithiol to the disulfide of thioredoxin by two sequential thiol-disulfide interchange reactions.3-15 The equilibrium constant ([TR(SH)J[NADP + ]/[TR(S2)][NADPH]) of the reaction for the E. coli enzyme is pH dependent having values of 44 at pH 7, 5 at pH 8, and 0.7 at pH 9.15 The pH optimum under a standard set of assay conditions is centered at about pH 7.7 in both phosphate and Tris but the absolute activity is approximately twofold higher in phosphate.15 The extinction coefficient of the FAD at 456 nm is 11.3 mM~ 1 cm~ 1 and the spectral ratios are A(271 nm):A(456 nm) - 5.8; A(380 nm):A(456 nm) - 1.03.296 Glutaredoxin also has a redox active disulfide.3'297'298 Whereas thioredoxin is reduced by NADPH via its reductase, thioredoxin reductase, glutaredoxin is reduced by glutathione.3 Overlap between the role of reduced thioredoxin in metabolism and the roles of glutathione and reduced glutaredoxin is not totally clear.3-298"301 Mutants of E. coli lacking either glutaredoxin or thioredoxin are viable, but the double mutant could not be obtained by PI transduction without the addition of cysteine to the medium, indicating that either thioredoxin or glutaredoxin is essential.302-303 Together however, they maintain the thiol/disulfide redox poise of the cytoplasm and serve as electron sources for systems such as ribonucleotide reductase,15'294'301'304-305 methionine sulfoxide reductase,306 and PAPS (adenosine 3'-phosphate-5'-phosphosulfate) reductase.307 While the term "thioredoxin" was coined in connection with ribonucleotide reductase, this protein had been demonstrated in the latter two systems previously.306-307 E. coli thioredoxin is required for phage (f 1, M13, and T7) replication.308-309 Glutaredoxin is related structurally to T4 phage-encoded thioredoxin and to the cytoplasmic glutathione transferases; its primary sequence relationship to E. coli thioredoxin is tenuous, but since the overall 3-dimensional structures of E. coli and T4 thioredoxins are similar, it has been postulated that glutaredoxin has a similar structure.298*310'314 T4 thioredoxin is reduced by E. coli thioredoxin reductase; the apparent KM values for the two thioredoxins are similar.314 Spinach contains three thioredoxins: m, f, and h.315 Thioredoxin h is reduced by an NADPHdependent reductase as in E. coli.316 Thioredoxins m and f are reduced in a light-dependent process via ferredoxin and ferredoxin-thioredoxin reductase and function in the regulation of malate dehydrogenase and fructose-1,6-bisphosphatase respectively.315"319 Several interesting functions have been indicated for mammalian thioredoxin, including reductive deiodination of thyroid hormone and influence on the binding state of the glucocorticoid receptor.320-321 Four proteins, rat liver microsomal protein disulfide isomerase, p subunit of chicken prolyl 4-hydroxylase, rat form I phosphoinositide-specific phospholipase C, and human thyroid-hormone-binding protein have been shown to contain two internally homologous domains which are approximately 30% homologous withE. coli thioredoxin.215'322'324 Indeed, protein disulfide isomerase is a substrate for bovine thioredoxin reductase.325 Figure 22 shows the active site sequences,326 and Table 7 outlines the functions of representative thioredoxins and glutaredoxins. Pig liver cytosol thioltransferase shows 82% identity with calf thymus glutaredoxin.327 Glutathione peroxidase from bovine erythrocytes and two glutathione S-transferases (rat liver cytosol a and JJL) have modest homology with calf thymus glutaredoxin but do not contain the characteristic thioredoxin active site sequence.312 Two short sequences (residues 75 to 77 and 91 to 93 in the E. coli numbering) together with the two residues between the cysteines form a flat surface on one side of the disulfide projection which is thought to be involved in protein binding.298-313 Only the thioredoxin reductase of E. coli has been studied in detail but the enzyme from rat liver has been purified and its basic properties surveyed. While the enzyme from both sources is a dimer of identical subunits, each subunit of E. coli thioredoxin reductase has a Mr of only 35,400 while the liver enzyme has a subunit Mr of — 50,000, i.e., more typical of the other family members.328 The enzyme has also been purified from baker's yeast,329

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FIGURE 22. Amino acid sequences of thioredoxin, glutaredoxin, and related proteins around the redox active disulfide. The additional short sequences, together with the two residues between the cysteine residues, form a flat surface thought to be involved in protein binding.313 Charged residues thought to influence the redox properties of the disulfide have been underlined in the Escherichia coli thioredoxin and glutaredoxin and in the phage T4 thioredoxin sequences; K57 is also part of this cluster in E. coli thioredoxin.313 E. coli thioredoxin and glutaredoxin, and phage T4, yeast, calf liver and Corynebacterium nephridii thioredoxin sequences are aligned as in Reference 310. The spinach chloroplast type m and Anabaena cylindrica thioredoxin sequences were aligned with the others as in Reference 318. The spinach chloroplast type f thioredoxin was aligned with the others as in Reference 317. The calf thymus glutaredoxin sequence was aligned with the others as in Reference 311. The Anacystis nidulans R2 thioredoxin m sequence was taken from Reference 319. Complete sequences often thioredoxins have been compared.326 The pig liver thioltransferase sequence was taken from Reference 327. The sequences around the first active sites of rat liver microsomal protein disulfide isomerase, p subunit of chicken prolyl 4-hydroxylase, rat form I phosphoinositide-specific phospholipase C and human thy raid-hormone-binding protein were taken from References 215, 322, 323, and 324.

TABLE 7 Thioredoxin and Glutaredoxin Functions Species and/or type

Function(s)

Bacterial, spinach type h and yeast thioredoxin

Reduction of ribonucleotide diphosphates Reduction of PAPS to SO^ + PAP in cysteine biosynthesis Reduction of methionine sulfoxide Reduction of ribonucleotide diphosphates Reduction of PAPS to SOi~ + PAP in cysteine biosynthesis Reduction of methionine sulfoxide Reductive deiodination of thyroid hormone Binding state of the glucocorticoid receptor Regulation of chloroplast malate dehydrogenase Regulation of chloroplast fructose- 1,6-bisphosphatase Assembly of filmentous phage Essential subunit of its phage-encoded DNA polymerase Reduction of ribonucleotide diphosphates

Bacterial and mammalian? glutaredoxin Mammalian thioredoxin Spinach and cyanobacterial thioredoxin m Spinach thioredoxin f Phage fl or M13 infected E, coli Phage T7 infected E. coli Phage T4 thioredoxin

168

Chemistry and Biochemistry of Flavoenzymes TABLE 8 Phenotypes of Glycine Fermenting Anaerobes Species Peptostreptococcus glycinophilus Clostridium cylindrosporum Clostridium sporogenes Clostridium siicklandii Bacterium W6 Eubacterium acidaminophilum

t-Lpd

TrR

Tr-like-P

0.4% 0.5% 0.01% yes 0.01% no

no(?) yes(?) 0.5% 0.5% yes 1.4%

7 9 9 9

yes yes

Note: Data are from the laboratory of Prof. Dr. Jan R. Andreesen, der Georg-August-Universitat Gottingen: percent of cell protein calculated by enrichment factors. t-Lpd = typical lipoamide dehydrogenase; TrR = thioredoxin reductase — dimer of 35K subunits — also referred to as atypical lipoamide dehydrogenase and ET-FP, electron-transferring flavoprotein; Tr-like-P — thioredoxin-like protein.

Novikoff ascites tumor cells,330 spinach,316 and Rhodobacter sphaeroides Y.331 The yeast, spinach, and Rhodobacter sphaeroides enzymes are of the lower Mr E. coll type. Purification of the E. coli enzyme is greatly assisted by affinity chromatography.332 However, the enzyme should be eluted with salt rather than with NADPH since aerobic turnover leads to spectral changes (filling in of the trough at 305 nm) which may be connected with the appearance of new fluorescent emission peaks suggesting the breakdown of histidine residues.128 Recent evidence, including amino acid sequence data, indicates that the enzyme referred to as atypically small lipoamide dehydrogenase (see Section III.A.2) may be a thioredoxin reductase.333 Two phenotypes are possible as shown in Table 8: Peptostreptococcus glycinopilus utilizes a typical lipoamide dehydrogenase for glycine catabolism, Eubacterium acidaminophilum utilizes thioredoxin reductase (atypical lipoamide dehydrogenase), and the other species appear to have a mixed phenotype. The thioredoxin reductase of Eubacterium acidaminophilum differs from the E. coli enzyme in requiring a large excess of NADPH for reduction.115'117 B. COMPARISONS AND CONTRASTS WITH THE OTHER MEMBERS OF THE FAMILY It is difficult to decide whether to emphasize the differences or the similarities between E. coli thioredoxin reductase and the other members of this family. The discussion in Section II.B.I. leaves little doubt that the flavin and pyridine nucleotide binding domains of thioredoxin reductase evolved divergently from a common ancestor with the others, but that its active site has evolved convergently (Figure 7 and 8). In addition to the differences in Mr and the structure around the redox active disulfide another contrast is important: the redox potentials of the FAD/FAE>H2 and the disulfide/dithiol couples are not well separated as they are in the other enzymes and no charge-transfer complex is observed.15'32'334'336 Chargetransfer does appear to be stabilized by the rat liver enzyme.328 Since liver thioredoxin reductase seems to be more like the other enzymes it is likely that its active site structure will be of the glutathione reductase type. Returning to E. coli thioredoxin reductase, two important similarities between it and lipoamide dehydrogenase are the mode of electron transfer between the two redox centers via a thiol-C(4a)-FAD covalent bond, and the presence of a base in the active site.20-32 Since the FAD and disulfide redox potentials are similar, the two-electron reduced state of the enzyme is an equilibrium of at least two forms: an FADH2-disulfide form and an FADdithiol form.

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FIGURE 23. Scheme defining the redox potentials in thioredoxin reductase. The definitions are in Table 9. (From O'Donnell, M. E. and Williams, C, H., Jr., J. Biol. Chem,, 258, 13795, 1983. With permission.)

1. Redox Potentials and Evidence for an Active-Site Base The small separation between the FAD/FADH2 and disulfide/dithiol redox potentials is evident in anaerobic reduction of E. coli thioredoxin reductase by NADPH or dithionite. Titration proceeds in a single phase, monitoring the main flavin band at 456 nm, with the uptake of four electrons/FAD. No long wavelength band attributable to charge-transfer is observed, but, depending on the presence of trace oxygen, a small amount of neutral (blue) semiquinone is seen.334 The four redox potentials defined in Figure 23 are given in Table 9 and were determined by equilibration with the NAD/NADH system. (NADH and NAD + do not form detectable complexes with the enzyme). An additional experiment was required in which the amount of microform III was determined at approximately half reduction by observing FADH2 reoxidation as microform III was converted to microform II (a step in catalysis and therefore fast) by complexing the latter with phenylmercuric acetate.32 It should be emphasized that this method yields microscopic potentials, whereas the potentials given for the other members of the family, though given as for specific chemical groups, are in fact macroscopic potentials for EOX/EH2 and EH2/EH4 (see also Section III.B.3). Examination of the redox potentials over the pH range 5.5 to 8.5 showed slopes of approximately 60 mV/pH for the FAD/FADH2 couple and 52 mV/pH for the disulfide/dithiol couple. Therefore, flavin reduction involved two protons as well as two electrons, but disulfide reduction involved an apparently nonintegral number of protons (1.8) for a twoelectron reduction. This suggested the presence of a base whose pKa depended on the redox state of the sulfurs as shown in Figure 24. The pKa values and the equilibrium constants were determined by a novel procedure in which a plot of K t /K 2 (as defined in Figure 23) vs. pH gave pK6 (the ionization of a group on microform III) and a plot of K 2 /Kj vs. pH gave pK5 (the ionization of a group on microform II). The group ionizing on microform II was the base (pKa = 7.6). The group ionizing on microform II was pictured as one of the thiols (pKa = 7.0). This assignment was based on precedents of other enzymes in which thiols in a relatively apolar milieu form ion-pairs with closely juxtaposed bases (see above). The pKa of the base on microform II then would be higher than its pKa on microform III because of the stability of the ion-pair.32

-327mV

-317 mV

25°C 167

-346mV 25°C 31

Em(pH7) (FAD/FADH2)

d

c

b

a

FAD-enzyme. FADH2-enzyme. Disulfide-enzyme. Dithiol-enzyme.

25°C 192

yes -242 mV

yes -264mV

yes -280mV

Stable charge transfer complex Em(pH7) (S-S/(SH)2

Temperature References

Yeast glutathione reductase

E. colt lipoamide dehydrogenase

Pig heart lipoamide dehydrogenase

TABLE 9 Oxidation Reduction Potentials

32

no

- 254 mVa -271 mVb -243 mVc -260mV d 12°C

E. coli thioredoxin reductase

25°C 16

- 335 mV

yes -269mV

P. aeruginosa mercuric reductase

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FIGURE 24. Scheme showing the protonation of species II and species III from Figure 23. (From O'Donnell, M. E. and Williams, C, H., Jr., /. Biol. Chem., 258, 13795, 1983. With permission.)

FIGURE 25. Scheme showing the oxidative half-reaction of thioredoxin reductase. The lower thiol is the interchange thiol and the upper thiol is the electron transfer thiol. TR(S)2 and TR(SH)2 are thioredoxin and reduced thioredoxin, respectively. The protonated base which donates the protons to thioredoxin is not shown; see Figure 24 where the species at the top right is the prototrophic tautomer of the left-hand species in this figure.

Dithiol-disulfide interchange between reduced thioredoxin reductase and thioredoxin must be initiated by a thiol anion based on model reactions.337 The ion-pair is therefore the logical species to initiate this reaction. The protonated base is also important in the dithioldisulfide interchanges as proton donor to thioredoxin, first to the nascent thiol anion in the mixed disulfide, and finally to the second nascent thiol anion as reduced thioredoxin is released (Figure 25). Formation of the mixed disulfide increases the acidity of the base since it then lacks its ion-pair partner. Reprotonation of the base by the second thiol of the reductase following the first interchange is assumed in Figure 25, The base then, in this working hypothesis, interacts with both thiols. The pKa values for the thiol (7.0) is higher by about two pH units than the value for analogous residues in yeast glutathione reductase.33 The significance of this is not obvious but it should be recalled that the inherent nucleophilicity of a thiolate increases as its pKa increases.338'339 Countering this is the fact that the rate of nucleophilic attack increases as the pH increases (higher nucleophile concentration).340 Comparing thioredoxin reductase and glutathione reductase at a given pH, the concentration of attacking nucleophile is less but the inherent nucleophilicity of that thiol anion is greater in thioredoxin reductase.

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Chemistry and Biochemistry of Flavoenzymes

2. Kinetics of the Reduction of Thioredoxin by NADPH NADPH reacts with the re face of the flavin ring of thioredoxin reductase as with glutathione reductase.85 Reduction by a single concentration of NADPH observed in the rapid reaction spectrophotometer at 2°C proceeds in three phases.341 The first phase is complete in the dead time of the instrument (~ 3 msec) and results in a spectrum with enhanced absorption at long wavelengths and only about 20% decrease in absorption in the 450 nm region. This species decays to the four-electron-reduced enzyme in two phases, having rate constants of 2600 min" 1 and 300 min" 1 , The rate of the overall reaction at low temperature is 520 min" 1 . 6 - 341 The scheme shown in Figure 23 (replacing NADH and NAD + with NADPH and NADP + respectively) represents a minimal reaction mechanism for thioredoxin reductase. Catalysis is pictured as proceeding I to III to II to I when thioredoxin is in excess and where I represents the oxidized enzyme. The rapidly formed species is considered to be a charge-transfer complex between oxidized enzyme and NADPH. The spectrum of the species formed with a rate constant of 2600 min" 1 has not been determined, but it is assumed to be III as a charge-transfer complex in which NADP + , still bound, is the acceptor and FADH2 is the donor. If NADPH is in excess, catalysis might be pictured as proceeding from I to III to II in a '*priming" reaction and then II to IV to III to II. The slow reaction with rate constant of 300 min" 1 may represent the dissociation of NADP + from IV. 3. Formation of a C(4a)-Thiolate Adduct with 1-deaza-FAD It is well known that the properties of the flavin are markedly influenced by the surrounding protein.342 Given that the redox potential of the FAD in thioredoxin reductase is less negative than in the other disulfide reductases that form charge-transfer complexes, it was suggested that replacement of the FAD by a flavin with still more negative potential such as 1-deaza-FAD might favor charge-transfer formation.343 Reduction of 1-deaza-FAD thioredoxin reductase, in contrast with the native enzyme, proceeds in two distinct phases as is the case with the other disulfide reductases.20 The spectrum of two-electron reduced enzyme (Figure 26), having more absorbance at 414 nm than either the oxidized or the fully reduced enzymes, does not indicate a charge-transfer complex since the absorbance of the new band at 414 nm increased as the pH decreased while the absorbance of a charge-transfer complex in which thiol anion is the donor has the opposite behavior. Fluorescence excitation and emission spectra taken concurrently with the optical spectra suggested that there were two two-electron reduced species, one a nonfluorescent species having absorbance at 414 nm (referred to as the 414 nm species), and a fluorescent species having a spectrum similar to that of oxidized flavin (referred to as the 550 nm species). These species were shown to be in rapid equilibrium by reaction of half reduced enzyme with phenylmercuric acetate which reacted with both species leading to the breakdown of the 414 nm species. The ratio of the species was pH dependent. From the changes in the spectrum of half reduced enzyme with pH, it was possible to derive the spectrum of the intermediate absorbing at 414 nm as shown in Figure 26. The spectrum, with a single peak having high extinction, was very similar to those of adducts with 1-deazaFAD at the C(4a) position, Plots of the ratio of the concentration of the 414 nm species to that of the 550 nm species and the inverse vs. pH indicated a group with a pKa of 7,41 on the 414 nm species and a group with a pKa of 6.73 on the 550 nm species. Figure 27 gives an interpretation of these data. The 414 nm species is pictured as a thiol to 1-deaza-FAD adduct which would be nonfluorescent. The flavin is oxidized in the 550 nm species. The pKa on the 414 nm species is attributed to the free thiol. It is perhaps not wise to attempt a correlation of this pKa with any in the native enzyme since it is difficult to predict the effects of the adduct involving a thiol which would be free in the native enzyme. The pKa on the 550 nm species is that of the thiol forming the ion-pair. The 550 nm species is favored at all pH values in the range observed.20

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FIGURE 26. Effect of pH on partially reduced 1-deaza-FAD-tnioredoxin reductase. Curve 1, oxidized enzyme; curve 2, 1.26 mol of dithionite/mo) of 1-deaza-FAD. The enzyme was then titrated anaerobically with 0.5 M acetic acid, curves 3 to 10 from pH 8.2 to pH 5.8. The dashed spectrum is an estimate of the 414 nm absorbing 1deaza-FAD-thioredoxin reductase species obtained from the difference between spectra 2 and 9 after correcting each for an 11% contribution from neutral semiquinone; extinction coefficients of 6.8 and 0.6 n\M~l cm' 1 at 550 nm for the 550 nm species and the 414 nm species, respectively were used to scale the difference spectrum. (From O'Donnell, M. E. and Williams, C. H., Jr., J. Biol, Chem., 259, 2243, 1984. With permission.)

FIGURE 27. Scheme showing the pH dependent equilibria between two-electron reduced 1-deaza-FAD thioredoxin reductase species. (From O'Donnell, M. E. and Williams, C. H., Jr., J. Biol. Chem., 259, 2243, 1984. With permission.)

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Chemistry and Biochemistry of Flavoenzymes

4. Photoreduction of Thloredoxin Reductase The formation of a blue (neutral) semiquinone in high yield upon irradiation of thioredoxin reductase in the presence of a large excess of EDTA is observed over a period of 4 h.344 The semiquinone is further reduced to FADH2 at an even slower rate. In contrast to this very slow semiquinone production, enzyme reduced by NADPH in the dark and subsequently exposed to light is rapidly converted to the semiquinone. The rate depends on the amount of NADPH used in the reduction; with 0.5 mol NADPH per FAD the half-time is less than 0.5 min, and wii;h 2.0 mol NADPH per FAD the half-time is about 2 min. Thus, FAD catalyzes the comproportionation in keeping with the known photosensitizing nature of free flavins. The rate of free radical production (EPR) exactly parallels the rate of increase in absorbance at 580 nm. The exact spectral characteristics of the semiquinone depend on the state of oxidation of the disulfide-dithiol. In the dithiol form the maximum is at 578 nm while in the disulfide form the maximum is at 588 nm. That the spectral properties are determined by the redox state of the disulfide is indicated by three findings. If semiquinone is produced by irradiation following reduction of the enzyme by 0.5 mol reductant/FAD, the maximum is at 588 nm, while if the semiquinone is formed following reduction by 2.0 mol reductant/FAD, the maximum is at 578 nm. Oxidation of enzyme irradiated in the presence of excess EDTA requires ferricyanide stoichiometric with the observed semiquinone for irradiation times less than 4 h (i.e., one-electron reduced enzyme), but for longer times, up to 4 equivalents of ferricyanide are required. The semiquinone exhibiting a maximum at 578 nm, upon addition of thioredoxin, shifts its maximum to 588 nm while NADPH causes the opposite shift.344 The lack of reactivity of the semiquinone per se with either thioredoxin or NADPH shows that it cannot be involved in catalysis. The rapid production of semiquinone by irradiation of partially reduced enzyme is a light-activated comproportionation since it is totally dependent upon the presence of some oxidized enzyme. Enzyme fully reduced by dithionite forms no semiquinone, while enzyme partially reduced by dithionite rapidly forms semiquinone upon irradiation. The semiquinone is rapidly reoxidized by oxygen to yield an enzyme with unaltered spectral and catalytic properties.344 Similar reactions have been very briefly reported for lipoamide dehydrogenase; the dark reverse reaction is comparatively rapid, being complete in 30 min.345 C. DISTINCT FUNCTIONS OF THE NASCENT THIOLS Specific alkylation of one of the nascent thiols in the two-electron reduced state of lipoamide dehydrogenase and glutathione reductase has allowed the assignment of the thiol nearer the amino terminus as the thiol-disulfide interchange thiol and the other as the electrontransfer thiol in the mechanism of both these enzymes.33-46 Alkylation studies with twoelectron reduced thioredoxin reductase resulted in an equal alkylation of both nascent thiols, suggesting a more open active site in this protein relative to the other enzymes.346-347 Consequently, assignment of the particular catalytic roles to each thiol was not possible. The gene (trxE) encoding thioredoxin reductase in £. coli has been cloned into the filamentous phage f 1 and a plasmid expression vector and subsequently sequenced.79-348 The gene can be expressed from the plasmid, pPMR14, under the control of the trxE promoter in an E. coli cell line (A304) deficient in thioredoxin reductase.349 Cys-135 and Cys-138 form the active site disulfide.65'66'79 Site-directed mutations of these residues individually, TRR(Cysl35 Serl38) and TRR (Serl35 Cysl38), have allowed the assignment of unique catalytic roles to these thiols.49 Spectral analyses of TRR(Serl35 Cysl38) as a function of pH and ionic strength has revealed a strong dependence of the spectrum on both parameters. The e458 of wild type, TRR(Cysl35 Cysl38), and e453 of TRR(Cysl35 Serl38) are pH independent. A new band tentatively identified as revealing a charge-transfer complex (e530 = 1300 M ~' cm ~') unique to TRR(Ser 135 Cysl38), has been observed under conditions of high ammonium cation

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TABLE 10 Steady-State Kinetic Parameters of the Three Thioredoxin Reductase Proteins

Enzyme

Turnover number

NADPH

pM

nmol/min/nmol TRR(Cysl35 Cysl38) TRR(Serl35 Cysl38) TRR(Cysl35 Serl38) a

1320 (2000) 143 790a

KM Thioredoxin

1.25(1.2) 0.88 1.76a

2.89 (2.8) 2.70 2.73a

Apparent KM and turnover number values calculated using a line determined by the first two data points in Figure 28c.

concentration. These results suggest the assignment of Cys-138 as the FAD-interacting thiol in the reduction of thioredoxin by NADPH via thioredoxin reductase.49 Several additional lines of evidence support this assignment.350'351 First, when the FAD is replaced with 1-deaza-FAD in each mutated enzyme, only TRR(Serl35 Cysl38) forms a pH dependent flavin-C(4a) adduct.350 A similar pH dependent adduct (Figure 26) forms when 1-deaza-FAD wild type enzyme is 2-electron reduced.20 Second, the absorbance spectrum of 4-thio-FAD-TRR(Serl35 Cysl38) after 12 h at 4°C is indicative of a mixture of approximately half 4-thio-FAD and half FAD, suggesting a reaction between the 4 position of the flavin and Cys-138, perhaps only on one subunit. In contrast, 4-thio-FAD-TRR(Cysl35 Ser 138) resembles the spectrum of 4-thio-FAD. (The binding of 6-thiocyanato-FAD to the mutant apoproteins showed no evidence for a reaction between either of the thiols and the 6 position of the flavin.) Thus, at least three lines of spectral evidence support the placement of Cys-138 nearer the FAD (upper thiol in Figure 25). If, as with the other two members of this enzyme family, the two distinct catalytic functions are each carried out by a different nascent thiol, then Cys-135 would be the interchange thiol (lower thiol in Figure 25).49-350 There is a modest negative interaction between redox centers in wild type thioredoxin reductase, i.e., the potential of the FAD/FADH2 couple depends on the redox state of the disulfide/dithiol and vice versa.32 As expected, the redox potentials of these mutated proteins (both - 280 mV) are closer to that for the FAD/FADH2 couple on dithiol-containing wild type enzyme (- 260 mV) than to the potential of disulfide-containing enzyme (- 243 mV).351 A small amount of blue, neutral semiquinone (~ 5%) is produced in dark, reductive titrations of wild type thioredoxin reductase. TRR(Cysl35 Serl38) and TRR(Serl35 Cysl38) have shown stabilization of 40% and 20% semiquinone respectively. These results indicate that the Ser hydroxyl group at position 138 may stabilize the semiquinone through hydrogen bonding with N(5) of the FAD more efficiently than does the thiol group.351 Similar stabilization of semiquinone by hydrogen bonding has been supported by crystal structures of flavodoxin.352 D. CATALYTIC ACTIVITY OF TRR(Serl35 Cysl38) AND TRR(Cysl35 Serl38) Steady-state kinetic analyses of the altered proteins have revealed turnover numbers of 10% and 50% of the value of the wild type enzyme for TRR(Serl35 Cysl38) and TRR(Cysl35 Serl38), respectively, and no changes in the apparent KM values of thioredoxin or NADPH (Table 10),49 All three enzymes show parallel-line kinetics when thioredoxin is the varied substrate, indicating that the mechanism of activity is unchanged in the mutated enzymes. However, in Figure 28 showing the secondary plots where NADPH is the varied substrate (at infinite thioredoxin), the expected linear behavior of wild type enzyme and of TRR(Serl35

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 28. Secondary plots of steady-state analyses. Panel a, TRR(Cys-135, Cys-138); Panel b, TRR(Ser-135, Cys-138); Panel c, TRR(Cys-135, Ser-138). (From Prongay, A. J., Engelke, D. R . , and Williams, C. H., Jr., J. Biol. Chem., 264, 2656, 1989. With permission.)

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FIGURE 29. Reduction of thioredoxin by the reduced forms of TRR(Ser-135, Cys138) and TRR(Cys-135, Ser-138). The positions of possible steps responsible for the catalytic inefficiencies are indicated by asterisks. (From Prongay, A. J., Engelke, D. R., and Williams, C. H., Jr., J. Biol Chem., 264, 2656, 1989. With permission.)

Cysl38) is not observed with TRR(Cysl35 Serl38) indicating a change in mechanism in this species. Thus, the value of Vmax for TRR(Cysl35 Serl38) of 50% that of wild type enzyme would be an upper limit. The catalytic activity possessed by the two mutated proteins is very surprising. The proposed mechanism for this family of enzymes (Schemes in Figures 2 and 3), including thioredoxin reductase (Figure 25), involves sequential thiol-disulfide interchange reactions in which the interchange thiol (lower, Cys-135) of 2-electron reduced enzyme attacks the disulfide of the substrate forming a mixed disulfide. This mixed disulfide is then attacked by the electron-transfer thiol (upper, Cys-138) of 2-electron reduced enzyme, reforming the active site disulfide and releasing the dithiol form of the substrate. The removal of either of the thiols of this active site disulfide should, it was thought, completely remove the catalytic activity with the disulfide containing substrate. Indeed, the active site Cys to Ser mutations of mercuric reductase are inactive, except that the Cys-135 Ser-140 mutated protein (electron transfer thiol mutated) has 3% the activity of the wild type enzyme in a disulfide (DTNB) reductase reaction.180 This result strongly suggests that FADH2 can directly reduce the mixed disulfide between Cys-135 and nitrothiobenzoate and offers a reasonable explanation for the catalytic activity of the thioredoxin reductase mutants. It would appear then that the remaining thiol in either mutation of thioredoxin reductase can fairly readily form a mixed disulfide with thioredoxin and that either of these can be reduced by FADH2, as shown in Figure 29, in which the suggested partially blocked step is indicated by an asterisk.49 If the indication that Cys-138 normally interacts with the FAD is correct, then the activity of TRR(Cysl35 Ser 138) would be limited by inefficient electron transfer directly from FADH2 to the mixed disulfide, since this step would have to replace the normal thiol-disulfide interchange initiated by Cys-138. The fact that mutation of Cys135 to Ser results in substantial loss of activity suggests that both thiols have distinct functions, leading to a working hypothesis that Cys-135 is the interchange thiol. The activity then of TRR(Serl35 Cysl38) would be limited by inefficient thiol-disulfide interchange with oxidized thioredoxin initiated by Cys-138. Electron transfer to the resulting mixed disulfide could be normal. The suggested mechanisms for activity in each mutated enzyme reemphasize the more open active site required by thioredoxin reductase in order to accom-

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Chemistry and Biochemistry of Flavoenzymes

modate its protein substrate. However, it is not suggested that wild type enzyme is any less specific in dithiol-disulfide interchange than are the other members of this enzyme family. The X-ray crystal structures of the wild type and mutated enzymes, now well advanced, will help to better define these possibilities.353-354 The structure of glutathione reductase shows that a near linear arrangement of substrate disulfide and protein disulfide perpendicular to the plane of the isoalloxazine ring exists, with the interchange thiol located at a greater distance from the FAD than is the proximal thiol.38'40'78 The experimental results presented here, combined with the previous A^-ethylmaleimide modification studies346 and redox studies,32-334 appear to indicate that the stereochemical arrangement may be different in E. coli thioredoxin reductase.49-350 The arrangement in glutathione reductase would not allow for either electron transfer by FADH2 to a mixed disulfide involving Cys-135 or for interchange with oxidized thioredoxin initiated by Cys-138 as observed. Examination of the structure of the substrate thioredoxin, reveals that the disulfide is located towards the end of a projection from the main body of the protein and that it is surrounded on three sides.295 In order to position the more exposed sulfur of the substrate disulfide such that it can interchange with the active site dithiol, most of the thioredoxin disulfide cage would have to be accommodated in the active site. In explaining the two results just cited in structural terms, it may be easier to picture the enzyme disulfide axis as more nearly parallel to the flavin ring than perpendicular (Figure 29). In this way, interchange initiated by Cys-138 and electron transfer from FADH2 to a mixed disulfide involving Cys-135 might be possible.49-350 This prediction should be confirmed or rejected by the X-ray crystal structure.354

VI. MERCURIC ION REDUCTASE A. METABOLIC ROLE! AND DISTRIBUTION Mercuric reductase is a flavoenzyme catalyzing the reduction of Hg(II) by NADPH, and is an essential component of the organomercurial and mercuric salts detoxification system.355"358 Since the Hg(II) must be chelated, preferably with a thiol ligand, in order to be a good substrate,355 it is not clear just how different is the chemistry of the reaction catalyzed, from the reactions catalyzed by the other members of this enzyme family. NADPH + R-S-Hg(II)-S-R + H+ -» NADP+ + Hg(0) + 2R-SH If the reduced flavin attacks the Hg(II) directly, then this chemistry would be unique within this enzyme family (Section VI.D). Alternatively, FADH~ might attack the sulfur to which Hg(II) is liganded in chemistry more analogous to that functioning in other members of the family. From residue 78 on, mercuric reductase is homologous with glutathione reductase and lipoamide dehydrogenase, particularly around the redox active disulfide.56 Residues 1 to 78 form a domain containing a thiol pair, Cys-10 and Cys-13 (in the Pseudomonas aeruginosa sequence) giving mercuric reductase a Mr of 58,660 Da.56 Removal of this domain by proteolysis does not affect the mercuric reductase activity.48 Mercuric reductase has an additional pair of cysteine residues, Cys-558 and Cys-559, not found in the other enzymes and it was suggested that these function to bind mercury at the active site.56 Mercuric reductase shares many spectral properties with lipoamide dehydrogenase and glutathione reductase, and forms an EHR-type species upon reductive alkylation with iodacetamide.16-48 The P. aeruginosa enzyme has been most thoroughly studied, but thus far, only the enzyme from Bacillus sp. strain RC607 has yielded crystals suitable for structure determination.359 Mercuric reductase from Mycobacterium scrofulaceum does not appear to require exogenous thiols and has equal activity with NADH or NADPH.360 The gene coding for mercuric reductase, merA is part of the mer operon consisting

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typically of the following genes (gene product and function in parenthesis): merR (regulatory protein, normally represses transcription, but acts as an inducer in the presence of Hg[II]), merT (transport protein), merP (periplasmic Hg[II] binding protein), merC (function unclear), merA (mercuric reductase), merD (function unclear), merB (organomercury lyase).361 The concept of a "bucket brigade" of dithiol-containing proteins has developed whereby Hg(II) is sequestered by MerP, conducted across the membrane by MerT, accepted by the Cys-10/ Cys-13 dithiol of mercuric reductase and finally liganded by Cys-558, Cys-559, and Cys135 for reduction.56'361363 The presumed ability of Hg(II) to form multidentate ligands, as it passes from one dithiol to the next, allows this whole process to occur without Hg(II) ever being free in solution.364'365 The mer operon is usually found on a plasmid, but the Bacillus sp. enzyme(s) are coded for on the chromosomal DNA. It is a curiosity that the Bacillus and Streptomyces lividans have a double N-terminal extension or no N-terminal extension, respectively.362'366 The resistance conferred is referred to as narrow spectrum (Tn501 on pVSl from Pseudomonas aeruginosa,56 Tn21 on R100 from Shigella flexnerii), that is, lacking the lyase, and thus conferring protection is against Hg(II) salts but not against organomercurials; or broad spectrum (pI258 from Staphylococcus aureus, pDU1358 from Serratia marcescens,367 Bacillus sp., and Streptomyces lividans), that is, having in addition the lyase and thus conferring protection also against organomercurials. The enzyme has also been isolated from Thiobacillusferrooxidans, an organism used in leaching low grade ores,368 and is related to the Tn501- and Tn21-derived enzymes.369 This list is not intended to be complete, but rather gives those species from which the isolated mercuric reductase has been sequenced (fully or substantially, Section II.B.I).56-75'77 Far broader comparisons based on restriction analysis or immunological cross-reactivity are available.370'372 Regulation of the transcription of the merA gene has been examined.373 B. KINETICS 1. Reduction of the Enzyme by NADPH Reduction of mercuric ion reductase (Tn501) by excess NADPH proceeds in three spectrally distinguishable stages: E ±5 E-NADPH ±* EH2-NADP+ ±5 EH2-NADPH the rate of the final stage being limited by the dissociation of NADP + . 163 These stages are similar to those observed in the reduction of glutathione reductase by NADPH (Section IV.C.2).35 Observing the spectral changes in a rapid-scan stopped-flow spectrophotometer at 5°C and pH 7.3, the formation of E-NADPH is virtually complete within the dead time of the instrument. The spectrum of this intermediate has a slightly diminished main flavin band and a very broad charge-transfer band centered at 600 nm. No intermediates have been observed in the passage of reducing equivalents via the FAD to the disulfide, and EH2-NADP+ is formed with an overall rate of 43 s"1 (kH/kD = 1.4). Its charge-transfer band is centered at 580 nm (e = 3.3 mM^ 1 cm^ 1 ). The rate constant for the displacement of NADP + by NADPH is 8 s" 1 (kH/kD = 1.1), and the charge-transfer band at 530 nm has an e = 5.0 mA/~' cm~ l . 163 - 374 Reduction with limiting NADPH indicates that the first two steps require one equivalent of NADPH while the third step uses a second equivalent of NADPH. It was suggested that EH2*NADPH might be an intermediate in the overall reaction since its rate of formation is faster than the rate of overall catalysis, — 0.7 s"1.163 This is confirmed by showing that the spectrum of EH2*NADPH predominates during the quasisteady-state phase of the overall reaction. Thus, the rate of catalysis is limited by the rate of reoxidation of EH2-NADPH by Hg(H). When Hg(H) is in excess over NADPH, the spectrum of EH2-NADPH is still formed initially. As the NADPH is exhausted, the spectrum shifts to that of EH2-NADP+ rather than that of Eox, indicating that this species is not able to reduce Hg(II).375

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Chemistry and Biochemistry of Flavoenzymes

2. Steady-State Kinetics Two assays, one based on volatization of Hg(0) and the other based on NADPH oxidation have been employed in the measurement of mercuric reductase activity. The former relies on counting the remaining 203Hg(II). In this assay in the presence of 1 mM 2-mercaptoethanol, the E. coll (R831 plasmid) enzyme showed a pH optimum of 7.5 and a K M for Hg(II) of 13 (xM at 37°C. A similar KM of 16 fiM was determined in the assay in which NADPH oxidation was measured.-155 A turnover number of 12.4 s"1 and a KM for Hg(II) of 12 |xM have been determined for the Tn501 -encoded P. aeruginosa enzyme under the same conditions with the NADPH assay.16'376 Activation of this enzyme (see Section C.2, below) by preincubation with NADPH resulted in the following at 25°C, pH 7.3, 1 mM cysteine: KM[NADPH1 = 0.4 |xM, KM[Hg(II)] = 3.2 |xM and turnover number = 13 s"1.355 Earlier work had compared EDTA and 2-mercaptoethanol as external ligands for the E. coli enzyme. In these studies the external thiol concentration was varied at constant Hg(II) concentration and the Hg(II) concentration was varied at a single concentration of external thiol. The data were analyzed according to a hysteretic model in which the transition from active to less active enzyme was brought about by the thiol ligand.377 A simpler model has been proposed for the Pseudomonas enzyme where it was shown that a competition exists between the external thiol and thiols of the enzyme.378 Thus, at concentrations of 2-mercaptoethanol higher than 1 mM, there was inhibition, while at lower concentrations there was substrate inhibition by Hg(II) in anaerobic assays varying both the external thiol and the Hg(II) concentrations.378 This competition then, dictated an optimal concentration of external thiol, and this was the concentration (1 mM) used in the earlier studies.16-355 A 1:1 covalent complex between Hg(II) and NADPH was reversed by added thiois and thus does not interfere with the assay. This complex bound tightly to mercuric reductase but did not reduce it.379 Mercuric reductase (P. aeruginosa) catalyzed the 2,4,6-trinitrobenzenesulfonate-dependent oxidation of NADPH. The reaction was inhibited by excess NADPH and markedly stimulated by NADP + (cf. Section IV.E).380 C. ACTIVE SITE THIOLS 1. The Active Site DisuJfide — Cys-135, Cys-140 The mercury resistance transposon Tn501 was originally isolated from Pseudomonas aeruginosa PAT.381 A number of site directed mutants of this enzyme (cloned into E. coli) have been prepared and their properties studied.180-382-383 A mutation resulting in replacement of the interchange Cys-135 by Ser, and thus having an intact electron transfer thiol, exhibits a pronounced charge-transfer band in the oxidized state. Cys-140 in this mutant enzyme has a pKa of 5.2, 1.5 pH units higher than the analogous glutathione reductase EHR discussed above (Section IV.D). The pKa of Cys-140 is 6.3 in the mutated enzyme that has Ala at position 135, as would be expected in the less polar environment. Following the loss of the charge-transfer absorbance associated with titration of Ala-135/Cys-140 enzyme with Hg(II) indicates that the Hg(II) is bound much less tightly than to the wild-type enzyme. The polarity also influences the position and shape of the charge-transfer band: Cys-135/Cys140 EH2, shoulder at 530 nm; carboxymethyl-Cys-135/Cys-140, peak at 560 nm; Ser-135/ Cys-140, concave extension of the main flavin band; Ala-135/Cys-140, peak at 560 nm 16,48,180,383 Alkylatlon of Cys-58 in glutathione reductase (analogous to Cys-135 in mercuric reductase) does not cause any perturbation of the charge transfer band.33 The mutation which results in replacement of the electron transfer Cys-140 by Ser is fully reduced by two electrons without any red charge-transfer intermediate. Neither of the mutant enzymes is active in Hg(II) reduction.180 Thus, the assignments of distinct nascent thiol functions made for lipoamide dehydrogenase and glutathione reductase holds in mercuric reductase (Sections III.D and IV.B). The Ala-135/Cys-140 mutant enzyme binds Hg(II) much less tightly than does the wild type enzyme but does catalyze about 30 turnovers at a rate of 0.03% that of wild type enzyme in an assay measuring Hg(0) volatilization.383

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FIGURE 30. Minimal catalytic mechanism for wild-type mercuric reductase utilizing both the active site dithiol and the C-terminal dithiol. (From Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C T., Biochemistry, 28, 1194, 1989. With permission.)

The redox potential of the FAD has been determined in mercuric reductase altered in one or the other of the active site cysteine residues. This potential is linearly related to the log of the turnover number in the transhydrogenase and oxidase activities.383 Two of the mutants, Ser-135/Cys-140 and Ala-135/Cys-140, have detectable mercuric reductase activity. They are able to turnover about 30 times at rates as high as 0.06% that of wild type enzyme.383 The juxtaposition of the active site thiols to the flavin ring in mercuric reductase may be different from the "perpendicular" arrangement in glutathione reductase, indeed it may be somewhat like that thought to obtain in thioredoxin reductase (Section V.D). This relative positioning has been probed by replacing the natural FAD with 6-thiocyanato-FAD in mercuric reductase and in glutathione reductase.271'384 The 6-thiocyanato-FAD is readily converted to 6-mercapto-FAD by reaction with thiols.385 The spectrum of the 6-thiocyanatoFAD bound to mercuric reductase is typical of this flavin in an apolar environment.384 Reduction of the active center disulfide by NADH leads to the conversion of the initially formed charge-transfer complex spectrum to that of 6-mercapto-FAD indicating that one (or both) of the nascent thiols can attack the thiocyanato group to displace cyanide. That the interchange thiol, Cys-135, is the reactive thiol is demonstrated by the conversion of 6thiocyanato-FAD to the mercaptide in Cys-135/Ala-140 but not in Ala-135/CyslO mercuric reductase substituted with 6-thiocyanato-FAD.384 Similar experiments with glutathione reductase show that neither nascent thiol can react with the 6-position of the flavin as would be predicted from the structure.270-271 2. The Auxiliary Thiols — Cys-558, Cys-559 In contrast to lipoamide dehydrogenase and glutathione reductase which cycle between Eox and EH2, mercuric reductase cycles between EH2 and EH4, as shown in the scheme of Figure 30. Moreover, the enzyme must be activated by the reduction of Cys-558/Cys-559

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 31. Reactions of NADPH with the activated (C-terminal dithiol) and nonactivated (C-terminal disulfide) forms of mercuric reductase. (From Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D,, Ballou, D. P., and Walsh, C. T., Biochemistry, 28, 1194, 1989. With permission.)

disulfide as indicated in the scheme of Figure 31,386 The need for activation by NADPH or cysteine had been shown earlier.376 In referring to EH2 and EH4 of mercuric reductase, it is assumed that Cys-558 and Cys-559 are present as thiols, i.e., the activated enzyme. Formation of EH2 has previously been interpreted to require only two electrons for reduction of the active site disulfide, the excess electron uptake observed being ascribed to poor anaerobic conditions.16 In anaerobic titrations with NADPH or dithionite, four electrons are required for the equilibrium conversion Eox to EH2 (Figures 32 and 33).386 The first additions of reductant lead to long wavelength absorbance typical of the charge-transfer complex, but the absorbance fades. Kinetic studies at 4°C and at 25°C show that reduction of the active site occurs rapidly, as previously reported;16 this is followed by a slower reduction of another redox group via reaction with the active site. Thiol titrations of denatured Eox and EH2 show that an additional disulfide is the group in communication with the active site. [l4C]-Iodoacetamide labeling experiments demonstrate that Cys-558 and Cys-559, near the C-terminus, are involved in this disulfide. The acetylcholine receptor contains a disulfide formed between adjacent cysteine residues, and the structure of a ds-peptide L-cysteinyl-L-cysteine has been determined.387*388 Both the active center dithiol and the C-terminal dithiol can be rapidly reoxidized by stoichiometric 4,4'-dithiodipyridine, a characteristic reaction of juxtaposed thiols. Air reoxidation of the C-terminal dithiol pair has a half-time of about 40 h.386 The double mutation of Cys-558 and Cys-559 to Ala also results in inactive enzyme consistent with the proposed function of these residues.382 The single mutants, Cys-558 to Ala and Cys-559 to Ala have been constructed. Cys-558 to Ala has a very low turnover number — 2 to 3% of wild type — and an elevated KM for Hg(II) indicating that Cys-558 is essential for normal catalysis. Cys-559 to Ala has a turnover number twofold higher than

FIGURE 32. Dithionite titration of mercuric reductase in the presence of methyIviologen. Spectra were recorded when no further absorbance changes occurred at 456 nm after, (-) 0 equiv, (—) 0.92 equiv, (-•-) 1.84 equiv, and (-••-) 3.31 equiv of dithionite. The inset shows the A530 (O) (corrected for absorbance of reduced methylviologen) and the flavin fluorescence at 520 nm, Xex = 456 (•), as a function of the equivalents of dithionite/FAD. Equilibrium absorbance due to methylviologen radical was seen only after addition of 1.65 equiv of dithionite. (From Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C. T, Biochemistry, 28, 1194, 1989. With permission.)

FIGURE 33. NADPH titration of mercuric reductase. Spectra were recorded when no further absorbance changes occurred after, (-) 0 equiv, (—) \ .07 equiv, (-•-) 2.13 equiv, (-••-) 2.98 equiv, and (-•••-) 15. \4 equiv of NADPH. The insets show (A) the absorbance at 620 (O), 580 (•), and 456 nm (A) and (B) the fluorescence emission of flavin at 520 nm, Xex = 456 nm (O), and of NADPH at 460 nm, Xex = 340 nm (•), as a function of equivalents of NADPH/FAD. (From Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C. T., Biochemistry, 28, 1194, 1989. With permission.)

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that of wild type but a markedly higher KM for Hg(II) indicating that Cys-559, while not essential for catalysis, assists the enzyme in its competition with the external thiol (usually 2-mercatoethanol, Section VI.B.2). 389 The fluorescence, but not the absorbance, of the enzyme-bound FAD was found to be highly dependent on the redox state of the C-terminal thiols (Figure 33).386 This is not surprising given that in the homologous glutathione reductase, the active site formed primarily from one monomer utilizes a base, His-467' near the C-terminus of the other monomer. A similar folding of the polypeptide chain of mercuric reductase could bring the C-terminal dithiol pair into juxtaposition with active center disulfide. Thus, Eox with Cys-558 and Cys559 as thiols exhibits less than 50% of the fluorescence of Eox where the residues are present as a disulfide, indicating that the thiols remain intimately associated with the active site. Initial velocity measurements show that the auxiliary disulfide must be reduced before catalytic Hg(II) reduction can occur,386 consistent with the report of activation by NADPH or cysteine.376 Direct evidence for the formation of the active site of mercuric reductase from the redoxactive disulfide of one polypeptide chain and the C-terminal dithiol pair of the second chain has emerged from elegant genetic recombination experiments.390 Cells harboring plasmids in which the MerA gene has been engineered for either of the changes, Ala-135Ala-140Cys558Cys-559 (AACC) or Cys-135Cys-140Ala-558Ala-559 (CCAA), are not resistant to Hg(H). However, cells in which AACC and CCAA are coexpressed are resistant. Characterization of the purified coexpressed enzyme mixture showed that it had 24% the activity of wildtype enzyme. Moreover, treatment of an equimolar mixture of AACC and CCAA with 1 M guanidinium chloride or 1.5 M sodium perchlorate resulted in activity 21% that of wildtype enzyme (after removal of the denaturants). The statistically expected activity level would be 25% in these mixtures of 50% homodimers (inactive) and 50% heterodimers since the heterodimers would contain one active site with four Cys residues and the other with all Ala residues.390 Given that the C-terminal dithiol is required for catalytic reduction of Hg(II), its exact role is a question of interest. The multiple liganding of Hg(II) prior to reduction (as shown in Figure 30) is suggested by the thermodynamics. Whereas the redox potential for free Hg(II) is ~ + 850 mV, the redox potential for Hg(Cys)2 is - 304 mV (calculated from the Ka of 1039);391-392 but the redox potential for Hg(II) bis-chelated by dithiols, as would be the case in the active center of mercuric reductase, is -438 mV (calculated from the Ka of 1043 5).393 Thus, Hg(II) bis-chelated at the active center would be difficult to reduce. Multiple liganding should make this reduction more facile given that each of the Hg-S bonds in the polycoordinate complex should be more ionic in character resulting in more positive charge being localized on the Hg(II), and hence, a higher redox potential. Such polycoordinate Hg(II) complexes have been shown to occur, using 13C chemical shift data, when the ligand to Hg(II) ratio is greater than two at neutral pH.364 Given the thermodynamics of Hg(II) bisliganding, it is difficult to envisage the transfer of Hg(II) from one dithiol to another without a transient tris-ligand. The rate of reduction of Hg(II) by l,5-dihydro-3,(3-sulfopropyl)lumiflavin at pH 4.7 decreases as the ligand to Hg(II) ratio rises indicating that trisligation decreases the redox potential of the Hg(II).394 It could be argued that this system is not a good model for reduction in the enzyme where juxtaposition of reactants is constrained. Figure 34 shows three possible mechanisms for the reduction of Hg(II) by enzymebound FADH2, and while multiple liganding is pictured in each case it is not essential. Mechanism A is an addition-elimination pathway. Mechanism B pictures formation of a flavin C(4a) thiol adduct concomitant with an outer-sphere reduction of Hg(II). This mechanism is thought to be utilized by the other enzymes of this family (Section VI. D). Mechanism C involves successive one-electron reduction steps.

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 34. Three possible mechanisms for reduction of multiliganded Hg(II) by enzyme-bound FADH2. (From Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C. T., Biochemistry, 28, H94, 1989. With permission.)

D. ELECTRON TRANSFER VIA A THIOL TO C(4a)-FAD ADDUCT AND DIFFERENTIAL REACTIVITY OF THE SUBUNITS Mercuric reductase forms a thiol to C(4a)-FAD adduct at low pH, 21 ' 22 analogous to that observed with lipoamide dehydrogenase EHR and thioredoxin reductase substituted with 1deaza-FAD.20-23 It was shown that at pH 5.5, the rapidly formed E-NADPH complex decays to a mixture of species with k - 145 s" 1 and a deuterium isotope effect of 2.4. The increase in absorbance at 395 nm associated with this decay suggested that one of the species was the thiol to C(4a)-FAD adduct,21 The demonstration of the involvement of one or both of Cys-558 and Cys-559 in catalysis led to the creation of a mutant enzyme in which both these residues and Cys-135 have been changed to Ala; thus, only the flavin interacting thiol remained; this enzyme was referred to as ACAA.22 Like SCCC (Ser-135, Cys-140, Section VI.C.I) this enzyme was red and the pKa of the remaining thiol could be determined by monitoring the charge transfer absorbance as a function of pH; it was 6.7 as compared with 5.2 for SCCC, the difference presumably being due to decreased polarity. The same pKa was determined by monitoring the fluorescence since ACAA, like E. coli lipoamide dehydrogenase EH2, had high fluorescence at low pH.167 Values of the pKa of the electron transfer thiol in species lacking the interchange thiol are compared in Table 11. As expected, ACAA was devoid of mercuric reductase activity. Its tight binding of pyridine nucleotides indicated that it had folded normally.

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TABLE 11 Values of the pKa of the Electron Transfer Thiol in Species Lacking the Interchange Thiol Enzyme

pK.

Ref.

Glutathione reductase EHR Mercuric reductase Serl35 Mercuric reductase Ala 135 Mercuric reductase ACAA

3.7 5.2 6.3 6.7

166 180 383 22

FIGURE 35. (A) Reduction of the active site disulfide of disulfide oxidoreductases and (B) Reactions of ACAA at pH 5 with pyridine nucleotides. (From Miller, S. M., Massey, V., Ballou, D. R, Williams, C. H., Jr., Distefano, M. D., Moore, M. J., and Walsh, C. T, Biochemistry, 29, 2831, 1990. With permission.)

Two intermediates thought to be common to catalysis by enzymes of this family, were directly observed in ACAA at low pH.22 The scheme of Figure 35 shows the pyridine nucleotide half reaction in wild type enzyme in line A. Line B shows the common blocked step both in reduction of ACAA by NADPH (left to right) and in reaction of ACAA with NADP+ (right to left). Reduction of ACAA by NADPH led to the formation of E-FADH -. ACAA was converted quantitatively to the adduct of thiol to C(4a)-FAD by the addition of NADP + . As shown in Figure 36, the adduct was characterized by \max = 382 nm and e = 7.5 mM~lcm~\ Analysis of the kinetics of this reaction observed in the rapid reaction spectrophotometer showed that adduct formation followed very fast complex formation between NADP+ and ACAA and allowed the calculation of the rate of adduct break down (a catalytic step) to be 0.9 s"1 at 5°C. Reduction of ACAA by NADPH is biphasic with rate constants of 260 and 32s" 1 . Thus, the rates of formation of E-FADH ~~ and the breakdown of the C(4a)-adduct — two catalytic steps (Figure 35) — were fast relative to the rate of overall catalysis, estimated to be 0.02 s"1 at 5°C.22 These results confirmed the postulated series of events in the reductive half reaction.21 Addition of the nonreducible NADP + analogue, aminopyridme adenine dinucleotide phosphate to ACAA at low pH led to conversion to the C(4a)-adduct but in only half the molecules. This way only one of many manifestations of extreme differential reactivity of the two monomeric units observed in mercuric reductase and its mutated forms.22 The kinetic analysis of adduct formation is summarized in the Scheme of Figure 37 with deprotonated species in the top row.22 The formation of the C(4a)-adduct in mercuric reductase and in thioredoxin reductase substituted with 1-deaza-FAD is promoted by low pH

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 36. Formation of a flavin C(4a)-cysteinyl adduct at pH 4.9 induced by binding of NADP + . Mercuric reductase ACAA mutant enzyme was titrated with NADP + . The inset shows a plot of A446 vs. the concentration of NADP + . (From Miller, S. M., Massey, V., Ballou, D. P., Williams, C. H., Jr., Distefano, M. D., Moore, M. J., and Walsh, C. T., Biochemistry, 29, 2831, 1990. With permission.)

FIGURE 37. Equilibrium fonnation of the C(4a)-thiol adduct in the ACAA mutant. (From Millers, S. M., Massey, V., Ballou, D. P., Williams, C. H., Jr., Distefano, M. D., Moore, M. J., and Walsh, C. T., Biochemistry, 29, 2831, 1990. With permission.)

with apparent pKa values of 6.7 and 7.4, respectively, ascribed to the flavin interacting thiol.20*22 The need for a proton in adduct formation derives from the extremely high pKa of the N(5) of reduced flavin. There is apparently no ready acid catalyst in the vicinity of N(5) except the attacking thiol itself. The value of the pKa of the C(4a)-adduct was assumed and this set the equilibrium constant for the formation of the unprotonated C(4a)-adduct (6.3

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X 10~ 4 ), since all other values were determined. The illustrated value for the acid dissociation constant of the C(4a)-adduct (10~ 10 M) is considered an upper limit, being higher than those estimated for the proton on N(5) in reduced flavins,395 and results in an upper limit of 6 x 10~4 for the equilibrium constant for the formation of the unprotonated C(4a)adduct. The pH dependence of C(4a) adduct formation in lipoamide dehydrogenase EHR is more complex being tightest atpH 8.8 and 5.6 and weakest around neutrality.47 The additional pKa values associated with adduct formation in lipoamide dehydrogenase EHR may reflect NAD + binding as well as adduct formation per se.

VII. OTHER PYRIDINE NUCLEOTIDE-DISULFIDE OXIDOREDUCTASES A. TRYPANOTHIONE REDUCTASE Eukaryotic, unicellular, parasitic protozoa appear to be more sensitive to oxidative stress than are their hosts (see also Section IV.E). This opportunity for chemotherapeutic intervention has generated much interest in these organisms. The Trypanosomatidae, Trypanosoma brucei, T. cruzi, Leishmania mexicana, and Crithidia fasciculata contain a novel enzyme capable of catalyzing the reduction of glutathione disulfide by NADPH but only in the presence of a dialyzable, heat-stable cofactor.396 This cofactor was identified as Nl ,N*bis(glutathionyl)-spermidine and dubbed trypanothione.397 The spermidine residue bridges the carboxylates of the glycine residues replacing two negative charges with one positive charge. Thus, reduction of oxidized trypanothione yields a single molecule, trypanothione, in contrast to the reduction of glutathione disulf ide which yields two molecules of glutathione; the entropy of the two reactions then are different. The properties of trypanothione reductase are so similar to those of glutathione reductase as to suggest that it can be thought of simply as a glutathione reductase with altered substrate specificity. The change in substrate specificity is virtually absolute however for the C. fasciculata and T. cruzi enzymes (Table 12).398'399 The sequence of the T. congolense enzyme has 40% identity with that of human glutathione reductase which is a higher level of identity than is observed between distinct enzymes in this family but indicates a long time of divergence (Figures 6 and 7). Trypanothione reductase from C. fasciculata has been purified and thoroughly characterized.398 As with the other members of this family, it was inactivated by iodoacetamide only when two-electron reduced. In a test of substrate specificity, it was shown that glutathione disulfide provided only minimal protection against alkylation of NADPH-reduced enzyme (0.04 vs. 0.05 min ~*). The titration of unmodified enzyme with dithionite proceeded in two, two-electron stages, and the charge-transfer complex (EH2) formed in the first stage had spectral properties almost identical with those of glutathione reductase (Table 12). The FAD was reduced in the second stage.398 The detailed substrate specificity of the C. fasciculata enzyme has been tested with a number of synthetic substrates including A^l-monoglutathionylspermidine disulfide (two spermidines) which is a good substrate.400 Trypanothione reductase from 7. cruzi has been purified and crystallized.399 Its properties were again virtually identical to those of glutathione reductase and trypanothione reductase from C. fasciculata. The enzyme may be identical to a NADPH-cytochrome c reductase characterized earlier.401-402 Sequence analysis of a peptide isolated from a digest of the alkylated, two-electron reduced enzyme showed that the thiol homologous with the interchange thiol of glutathione reductase had been modified.399 The enzyme had 50% activity in the 2 M ammonium sulfate used to stabilize the crystals. This indicates that, as with glutathione reductase, it will be possible to compare the structures of the native enzyme and enzyme reacted with its substrates in the crystal. This in turn holds out great hope for rational drug design for Chagas' disease caused by T. cruzi.399 The drug nifurtimox, used in the

190

Chemistry and Biochemistry of Flavoenzymes TABLE 12 Properties of Trypanothione Reductase — Comparison with Glutathione Reductasea

Property Mr(kDa) Cof actor Intracelluiar concentration Substrates at the pyridine nucleotide site K M NADPH Activity ratio NADH/NADPH Substrates at the disulfide site K M glutathione disulfide K M trypanothione disulfide K M monoglutathionylspermidine disulfide Specific activity TN (min- 1 ) Glutathione Trypanothione Activity ratio (TSST/GSSG) pH optimum [buffer] TN/K M (M-'s-' x 10- 6) Inhibition by nifurtimox Eox, Xmax (nm) e at X^ (mM" 1 cm' 1 ) Charge transfer in EH2 (530 nm) €(mM~l cm- 1 ) e (mA/- 1 cm-'XEHz-NADPH) Highly reactive cysteine residues (native) Oxidized species Reduced species Total reactive cysteine residues (denatured) Oxidized species Reduced species Total cysteine residues (sequence) a b

T. cruzi trypanothione reductase

Human glutathione reductaseb

C. fasciculate trypanothione reductase

50.0

52.5

53.8

FAD

FAD

1.25 \LM

0.1 fxA/

FAD —

5 V.M

8.5 \LM 0.05

7 |JLM 0.003

0.03 no substrate 45 M,A/ 275 \LM 284 U/mg no substrate 14200

65 \LM

—

no substrate no substrate 240 U/mg

53 jJiA/ 149 \iM 576 U/mg

12600

3.1

9.6

— 7.5

0.0008

31000 10,000

7.3

7.8

40mA/ 5.26 negligible

40mA/ 3.23

100mA/ 9.75

yes 463

— 464

461 11.3

11.3

11.3

yes — 4.9

yes 3.6 4.5

yes

0.0 1.0

0.3 1.3 — —

— — —

10

3.63

— — — 2.2 3.9 12

Adapted from References 398 and 399. Cf. Table5.

treatment of Chagas' disease, inhibited trypanothione reductase (50% at 200 jjiAf probably a higher concentration that can be practically achieved physiologically) but inhibited human glutathione reductase (75% at 50 \^M) confirming the previously held notion that this drug acts by inhibiting the antioxidant system of the host.399-403-404 Three naphthoquinones and five nitrofurans have been synthesized and compared with nifurtimox as substrates and inhibitors of C. fasciculata trypanothione reductase. Aerobically the active compounds turnover. Anaerobically they combine with the reduced enzyme irreversibly. They also inhibit the normal trypanothione reductase activity. The ability of the compounds to act as substrates correlated with their inhibition of T. cruzi trypomastigotes infectivity.405 Several of these compounds have now been tested with the enzyme of interest for Chagas' disease, T. cruzi trypanothione reductase.406 A nitrofuran derivative, 2-(5-nitro2-furanylmethylidene)-A^,A^'-[l,4-piperazinediylbis(l,3-propanediyl)-bishydrazinecarboximidamide tetrahydrobromide], proved to be a more powerful inhibitor of trypanothione reductase (K; = 0.6 |xM) than of glutathione reductase (K^ = 45 (juAf). The dual effect of oxygen-dependent turnover and inhibition of trypanothione reductase by a naphthoquinone

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derivative, 2,3-bis[3-(2-arnidinohydrazono)-butyl]-l,4-naphthoquinonedrihydrochloride, previously observed with the C. fasciculata enzyme,405 was also seen with the enzyme from T. cmzi but was not observed with glutathione reductase.406 Site directed mutagenesis has been used to study the origin of the distinct substrate specificity of T. congolense trypanothione reductase. In glutathione reductase, Arg-37 and Ala-34 make contacts with the carboxylates of the glutathione glycine moities. Changing the analogous residues in trypanothione reductase, Trp-21 to Arg and Glu-18 to Ala, reduced the activity with oxidized trypanothione by ten- and twofold, respectively but did not increase the activity with glutathione disulfide.407 B. BIS-7-GLUTAMYLCYSTINE REDUCTASE "y-Glutamylcysteine appears to function in a manner similar to glutathione in halobacteria. It is formed by reduction of bis-'y-glutamylcystine and the reaction is catalyzed by bis-^glutamylcystine reductase.408 The enzyme isolated from Halobacterium halobium is similar to glutathione reductase in all properties tested. Thus, it is a NADPH-dependent dimer of apparent Mr 122,000 containing FAD. It is highly specific for bis-^-glutamylcystine, having less than 2% the activity with glutathione disulfide. It catalyzes the reduction of DTNB at 23% the rate with the natural substrate. The very preliminary characterization of the steadystate kinetics of bis-'y-glutamylcystine reductase indicated that it is much less active than glutathione reductase/08 The turnover number was 28 s~l per FAD at infinite substrate concentration and 1.7 mA/ NADPH in an assay measuring the reduction of DTNB, present at a concentration of 50 |xM. The apparent KM for fc/s-'y-glutamylcystine under these conditions was 0.81 mM. The apparent KM for NADPH was 0.29 mM measured at a substrate concentration of 2.0 mM in an assay measuring NADPH oxidation. The assay buffer for these studies contained 50 mM sodium phosphate, 3 M KCI, 1.3 M NaCl and 1 mM EDTA at 30°C. The pH optimum was 7.5 with 50 mM Tris added to buffer at high pH, in the assay measuring NADPH oxidation where the substrate and NADPH concentrations were 1.0 and 0.17 mM, respectively. Given the source of bis-'y-glutamylcystine reductase the effect of salts on activity was of interest. Whereas, glutathione reductase activity is highest at a concentration of NaCl of 0,16 M,193 the activity of bis-^-glutamylcystine reductase peaked at an ionic strength of 1.8 M and decreased only very slightly as the ionic strength was increased to 4.3 M in the assay measuring DTNB reduction.408 It was observed that salts inhibited the Cu(II)-catalyzed oxidation of glutathione and -y-glutamylcysteine. The half-times for 7-glutamylcysteine were 5 and 27 min at low and high ionic strength, respectively. The half-time for glutathione was 18 min at low ionic strength and this thiol decreased by only 10% at high ionic strength. The fact that 'y-glutamylcysteine was more stable at high ionic strength than was glutathione at low ionic strength was noted. C. CYSTINE REDUCTASE This enzyme has been studied only in crude extracts of yeast and pea seeds. It is specific for NADH.409 The apparent KM for L-cystine was 0.9 mM.410 Several a-substituted cystines were also good substrates having similar KM values and Vmax values.410 D. ASPARAGUSATE DEHYDROGENASE Asparagus mitochondria contain, in addition to lipoamide dehydrogenase, two related enzymes, asparagusate dehydrogenases I and II.411 Asparagusic acid is 4-carboxy-l,2-dithiolane. The three enzymes have been isolated and compared. The lipoamide dehydrogenase does not reduce asparagusic acid but both asparagusate dehydrogenases reduce lipoic acid with an apparent KM of 3 mM to be compared with an apparent KM for asparagusate of 20 mM. While the lipoamide dehydrogenase activity of asparagusic acid dehydrogenase was

192

Chemistry and Biochemistry of Flavoenzymes

stimulated by NAD + , the asparagusic acid dehydrogenase activity was not. Asparagusate dehydrogenase was inhibited by pCMB, Hg(II), and arsenite, but inhibition by the latter required reduction by NADH in the preincubation with arsenite prior to assay. All three enzymes were reported to contain only one FAD per mol of enzyme of Mr 112,000.4U E. SULFHYDRYL OXIDASE Sulfhydryl oxidase is a FAD-containing enzyme, isolated from the brine suspending the mycelia of Aspergillus niger, catalyzing the following reaction: 2GSH + O2 -> GSSG + H2O2 where GSH and GSSG represent glutathione and glutathione disulfide, respectively.412 Thus, the reaction is typical of that catalyzed by enzymes of the dehydrogenase/oxidase class of flavoproteins. Yet, there are compelling reasons to include sulfhydryl oxidase within the broader family of pyridine nucleotide-disulfide oxidoreductases. The enzyme is alkylated with loss of activity only when reduced. The spectrum following reaction has a single maximum at 391 nm — typical of a C(4a)-adduct.413 Sulfhydryl oxidase required only 15-fold purification to yield an apparently homogeneous enzyme — a single diffuse band on sodium dodecyl sulfate-polyacrylamide electrophoresis.412 Very careful amino acid analysis clearly distinguishes this enzyme from glucose oxidase, another dehydrogenase/oxidase excreted into the medium of A. niger414 The low tryptophan content of sulfhydryl oxidase is a characteristic of the reductases while its high carbohydrate content is typical of the oxidases.6 Glutathione was the preferred substrate having an apparent KM of 0.3 mM and a turnover number of 131 s~'. Cysteine, dithiothreitol, and 2-mercaptoethanol gave similar turnover numbers but apparent KM values 2 to 3 orders of magnitude higher. The enzyme also enhanced the rate of reoxidation of reduced ribonuclease. The acid pH optimum, 5.5, of this enzyme is a curiosity since the substrate must be deprotonated for attack on a putative active site disulfide. The FAD of sulfhydryl oxidase was fully reduced by excess glutathione; the only trace of long wavelength absorbance was that of blue semiquinone. The flavin is bleached only about 27% by suifite (KD = 3 mM) whereas full reduction represents 68% bleaching. This is suggestive of half-sites reactivity in the dimeric enzyme. Oxidases are typically bleached the same extent as in full reduction and reductases are not bleached at all by suifite.415 The highly active sulfhydryl oxidase of rat seminal vesicle secretion, a monomeric FAD-containing enzyme of Mr 66,000, was not bleached by suifite.416 Thus, the properties of A. niger sulfhydryl oxidase suggest that while the reaction catalyzed is typical of enzymes of the dehydrogenase/oxidase family, an active site disulfide is present. The sequence around the cysteine residues forming this disulfide will be of interest. F. NADH PEROXIDASE Two flavoenzymes, NADH oxidase and NADH peroxidase from the cytochrome-free Streptococcus faecalis 10C1 catalyze the following reactions:417-418 2NADH + O2 + 2H+ -> 2NAD+ + 2H2O NADH + H202 + H + -» NAD + + 2H2O (Hydrogen peroxide is produced in Streptococcus faecium by a third flavoenzyme that oxidizes a-glycerolphosphate.)419>420 The basis for the inclusion of NADH oxidase and NADH peroxidase in this family of reductases is two-fold: first, very early work on the NADH

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FIGURE 38. Amino acid sequences of the N-terminus of NADH peroxidase and two peptides from NADH oxidase aligned with glutathione reductase. (From Poole, L. B. and Claiborne, A., J. Biol. Chem., 264, 12322, 1989 and Ahmed, S. A. and Claiborne, A., J, Biol. Chem., 264, 19856, 1989. With permission.)

peroxidase showed that it formed a band at long wavelength, very similar to the chargetransfer band of lipoamide dehydrogenase EH2-NAD(H)+ , when reduced with excess NADH but not NADPH.417'421-422 Moreover, more than one equivalent of dithionite was required for complete, two-phased reduction of the FAD.422 A later study showed that two electrons were taken up in each phase.423 NADH oxidase also appeared to take up more than two electrons in the reduction of its FAD.424 Second, it has recently been discovered that both enzymes are related to other members of this family.425-426 Examination of the amino acid sequence of the N-terminus of the NADH peroxidase reveals a typical nucleotide binding fold. If this sequence, with a one residue insertion, is aligned with the homologous sequence of E. coli glutathione reductase as shown in Figure 38, a single cysteine residue is seen at the position of the charge-transfer (proximal) cysteine residue of glutathione reductase. The interchange thiol is absent.425 A peptide from the NADH oxidase is identical in sequence with the putative active site region of the NADH peroxidase at 10 of 18 positions including the cysteine residue near the center of the sequence (Figure 38).426 The nature of the second redox active group of NADH peroxidase, postulated on the basis of early results,422 has recently been clarified.427'428 It is proposed that the putative active site cysteine residue is present as the sulfenic acid (-SOH) in oxidized enzyme and as the thiol in EH2.427 A cysteine sulfenic acid residue is involved in the mechanism based on earlier kinetic studies to be discussed later.429 Several lines of chemical evidence support this proposal.427 The EH2 form of the peroxidase did not react with iodoacetamide to form an EHR as did other enzymes in the family, and this was not surprising since the peroxidase lacks a thiol at the position of the interchange thiol. The charge-transfer of the peroxidase EH2 as well as enzymatic activity were abolished by treatment with phenylmercuric acetate in a reaction reminiscent of the loss of charge-transfer induced in lipoamide dehydrogenase EHR by aminopyridine adenine dinucleotide.46 The peroxidase was reactivated by dithiothreitol. Anaerobically denatured peroxidase at the oxidized level reacted with 5-thio-2nitrobenzoate to form a mixed disulfide from which the thionitrobenzoate could be quantitatively released by dithiothreitol. The oxidized peroxidase was inactivated by 10 mM H2O2 presumably due to the oxidation of the putative cysteine sulfenic acid to the sulfinic or sulfonic acid levels; as expected, this oxidation was not reversible with dithiothreitol. The FAD of peroxide-inactivated enzyme was reduced in a single phase by ~ one equivalent of dithionite in contrast to the native peroxidase which required two equivalents in a twostage reduction. Each of these results were consistent with the presence of a cysteine sulfenic acid residue acting as the second redox active group in the NADH peroxidase.427 Fast atom bombardment mass spectral analysis of a chymotryptic octapeptide containing the unmodified cysteinyl residue did not confirm the presence of the cysteine sulfenic acid form of the residue. The predicted molecular ion would have been a mass of 930. The major observed mass was 978 or three oxygen atoms too heavy. There was a minor peak at a mass of 914 equivalent to the thiol form. These data were interpreted as indicating primarily the cysteic acid form plus methionine sulfoxide, with a small amount of the products of disproportionation of the cysteine sulfenic acid form to the thiol and cysteine sulfinic acid forms.427

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FIGURE 39. Scheme for the activation and catalytic cycle for NADH peroxidase. (From Poole, L. B. and Claiborne, A., /. Biol. Chem., 264, 12330, 1989. With permission.)

The mechanism proposed for NADH peroxidase is shown in the scheme of Figure 39. It involves a priming reaction (in agreement with the earlier kinetic mechanism discussed below)429 and then cycling between EH2 and EH2-NADH levels.427 Thus, the reductant of the peroxide is the thiolate rather than FADH2 as would be the case with the dehydrogenase/ oxidase family of flavoenzymes. The resulting sulfenic acid is immediately reduced by the already bound NADH via the FAD reducing its exposure to further oxidation by excess peroxide. The presence of the NADH makes the thiolate a better nucleophile in the formation of the S-O bond. Chemical precedents for this reaction were cited.427 A very complete steady-state kinetic analysis, which included product and dead-end inhibition measurements of NADH peroxidase at 25°C and pH 7.5, indicated a ping-pong mechanism.429 The KM (H2O2) was 12 fiM and the turnover number (assuming subunit Mr of 46,000) was 11.3 s"1. The KM(NADH) was so low, 2 jitM, as to make initial rate measurements difficult, but the kinetic pattern using equipotential desamino-NADH (KM = 24 jxAf) was unchanged. The 4(S)hydrogen of NADH was transferred in catalysis as with the other members of this family. There was no primary tritium kinetic isotope effect. Primary deuterium kinetic isotope effects were observed on V but not on V/K. The enzyme catalyzed a NADH to thio-NAD+ and NADH to NAD + isotope exchange at about 1 to 2% the rate with hydrogen peroxide. The mechanism based on these studies, in contrast to the more recent one discussed above,427 proposed a nucleophilic attack of the thiolate on preformed C(4a)-flavin hydrogen peroxide.429 A preliminary report of X-ray crystallographic studies on NADH peroxidase shows that the enzyme is active in the crystalline state and that the crystals diffract to at least 0.25 nm.430 Turning to the structurally related NADH oxidase of Streptococcus faecalis 10C1, a cysteine sulfenic acid residue has again been proposed as the second redox active group.426-431 This was based on the isolation of a peptide homologous with the putative active-site peptide of the peroxidase and on the sensitivity of the enzyme to inactivation by peroxide. The behavior of the oxidase in reductive titrations was markedly different from the peroxidase. The dimeric enzyme accepted only six electrons from dithionite to give full reduction of the FAD and it was suggested that one of the two cysteine sulfenic acid residues was not reduced. This was supported by the fact that addition of the nonreducable pyridine nucleotide analogue,

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FIGURE 40. Scheme for catalysis by NADH oxidase. (Modified from Ahmed, S. A. and Claiborne, A., 7. Biol. Chem., 264, 19864, 1989. With permission.)

aminopyridine adenine dinucleotide to the dithionite-reduced enzyme resulted in reoxidation of half the FAD indicating that electrons had passed to the cysteine sulfenic acid residue. Moreover, titration of dithionite-reduced enzyme with NAD + gave reoxidation of half the flavin without the formation of NADH. Thus, the oxidized pyridine nucleotide acts as an effector for the transfer of electrons from sulfur to flavin as does NAD + in lipoamide dehydrogenase.136 Another distinctive feature of the NADH oxidase was the facile reduction of FAD. The FAD was reduced even by acetylpridine adenine dinucleotide which has a much higher redox potential than the NADH/NAD+ couple. The mechanism proposed for the oxidase reflects the dual nature of this unique flavoenzyme as both an oxidase and a dehydrogenase (Scheme of Figure 40). The four-electron reduction of oxygen is accomplished in two stages following the 4-electron reduction of the enzyme by two equivalents of NADH: Eox —> EH2 —> EH4 —> EH2 —> Eox. FADH2 is the reductant of molecular oxygen, while as with the NADH peroxidase, thiolate is the reductant of the resultant peroxide.426*431

ACKNOWLEDGMENTS The author is grateful to his many colleagues who read the first draft and made helpful comments and suggestions: Jan Andreesen, Alan Berry, John Blanchard, Hiram Gilbert, John Guest, Nancy Hopkins, Luise Krauth-Siegel, Brett Lennon, Vincent Massey, Rowena Matthews, Susan Miller, Scott Mulrooney, Mulchand Patel, Richard Perham, Lester Reed, Lena Sahlman, Heiner Schirmer, Simon Silver, and John Sokatch. Enlightening discussion with Rowena Matthews was appreciated. John Blanchard and Jan Andreesen made unpublished data available and the latter helped design Table 8. The production of this chapter was made immeasurably easier by the dedicated efforts of Donna Veine. The work from this laboratory has been supported by the Department of Veterans Affairs and by the National Institute of General Medical Sciences.

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169. Wilkinson, K. D. and Williams, C. H., Jr., NADH inhibition and NAD activation of Escherichia coli lipoamide dehydrogenase catalyzing the NADH-lipoamide reaction, /. Biol Chem., 256, 2307, 1981. 170. Sahlman, L. and Williams, C. H., Jr., Lipoamide dehydrogenase from Escherichia coli — steady-state kinetics of the physiological reaction, J. Biol. Chem., 264, 8039, 1989. 1 7 1 . Allison, N., Williams, C. H., Jr., and Guest, J. R., Overexpression and mutagenesis of the lipoamide dehydrogenase of Escherichia coli, Biochem. J., 256, 741, 1988. 172. Koike, M., Shah, P. C., and Reed, L. J., a-Keto acid dehydrogenation complexes. III. Purification and properties of dihydrolipoic dehydrogenase of Escherichia coli, J. Biol. Chem., 235, 1939, 1960. 173. Williams, C. H., Jr., Studies on lipoyl dehydrogenase from Escherichia coli, J. Biol, Chem., 240, 4793, 1965. 174. Thorpe, C. and Williams, C. 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200. Arscott, L. D., Drake, D. M., and Williams, C. H., Jr., Inactivation-reactivation of two-electron reduced Escherichia coli glutathione reductase involving a dimer-monomer equilibrium, Biochemistry, 28, 3591, 1989. 201. Boggaram, V., Brobjer, T., Larson, K., and Mannervik, B., Purification of glutathione reductase from porcine erythrocytes by the use of affinity chromatography on 2',5'-ADP-sepharose 4B and crystallization of the enzyme, Anal Biochem., 98, 335, 1979. 202. Carlberg, I. and Mannervik, B M Purification by affinity chromatography of yeast glutathione reductase, the enzyme responsible for the NADPH-dependent reduction of the mixed disulfide of coenzyme A and glutathione, Biochem. Biophys. Acta, 484, 268, 1977. 203. Carlberg, I. and Mannervik, B., Purification and characterization of glutathione reductase from calf liver. An improved procedure for affinity chromatography on 2',5'-ADP-sepharose 4B, Anal. Biochem., 116, 531, 1981. 204. Worthington, D. J. and Rosemeyer, M. A., Glutathione reductase from human erythrocytes. Catalytic properties and aggregation, Eur. J. Biochem., 67, 231, 1976. 205. Mata, A. M., Pinto, M. C., and Lopez-Barea, J., Purification by affinity chromatography of glutathione reductase (EC1.6.4.2) from Escherichia coli and characterization of such enzyme, Z. Naturforsch., 39c, 908, 1984. 206. Gorin, G., Esfandi, A., and Guthrie, G. B., Jr., Glutathione: its reaction with NADP and its oxidationreduction potential, Arch. Biochem. Biophys., 168, 450, 1975. 207. Jocelyn, P. C., The standard redox potential of cysteine-cystine from the thiol-disulfide exchange reaction with glutathione and lipoic acid, Eur. J. Biochem., 2, 327, 1967. 208. Kolthoff, I. M., Stricks, W., and Kapoor, R. CM Equilibrium constants of exchange reactions of cystine with glutathione and with thioglycolic acid both in the oxidized and reduced state, J, Am. Chem. Soc., 77, 4733, 1955. 209. Gorin, G. and Doughty, G., Equilibrium constants for the reaction of glutathione with cystine and their relative oxidation-reduction potentials, Arch. Biochem. Biophys., 126, 547, 1968. 210. Engel, P. C. and Dalziel, K., The equilibrium constants of the glutamate dehydrogenase systems, Biochem. J., 105, 691, 1967. 211. Mapson, L. W. and Isherwood, F. A., Glutathione reductase from germinated peas, Biochem. J., 86, 173, 1963. 212. Epp, O., Ladenstein, R., and Wendel, A., The refined structure of the selenoenzyme glutathione peroxidase at 0.2-nm resolution, Eur. J. Biochem., 133, 51, 1983. 213. Graminski, G. F., Kubo, Y., and Armstrong, R. N., Spectroscopic and kinetic evidence for the thiolate anion of glutathione at the active site of glutathione S-transferase, Biochemistry, 28, 3562, 1989. 214. Bulleid, N. J. and Freedman, R. B., Defective co-translational formation of disulphide bonds in protein disulphide-isomerase-deficient microsomes, Nature, 335, 649, 1988. 215. Parkkonen, TM Kivirikko, K. L, and Pihlajaniemi, T., Molecular cloning of a multi-functional chicken protein acting as the prolyl 4-hydroxylase p-subunit, protein disulphide-isomerase and a cellular thyroidhormone-binding protein. Comparison of cDNA-deduced amino acid sequences with those in other species, Biochem. J., 256, 1005, 1988. 216. Carlberg, I., Depierre, J. W., and Mannervik, B M Effect of inducers of drug-metabolizing enzymes on glutathione reductase and glutathione peroxidase in rat liver, Biochim. Biophys. Acta, 677, 140, 1981. 217. Tuggle, C. K. and Fuchs, J. A., Glutathione reductase is not required for maintenance of reduced glutathione in Escherichia coli K-12, J. Bacterial, 162, 448, 1985. 218. Jocelyn, P. C. and Kamminga, A., The non-protein thiol of rat liver mitochondria, Biochim. Biophys. Acta, 343, 356, 1974. 219. Meredith, M. J. and Reed, D. J., Status of the mitochondria! pool of glutathione in the isolated hepatocyte, J. Biol. Chem., 257, 3747, 1982. 220. Griffith, O. W. and Meister, A., Origin and turnover of mitochondrial glutathione, Proc. Natl. Acad. ScL U.S.A., 82,4668, 1985. 221. Taniguchi, M., Hara, T., and Honda, H., Similarities between rat liver mitochondrial and cytosolic glutathione reductases and their apoenzyme accumulation in riboflavin deficiency, Biochem. Int., 13, 447, 1986. 222. Drumm-Herrel, H., Gerhaufler, U., and Mohr, H., Differential regulation by phytochrome of the appearance of plastidic and cytoplasmatic isoforms of glutathione reductase in mustard (Sinapis alba L.) cotyledons, Planta, 178, 103, 1989. 223. Ondarza, R. N., Abney, R., and Lopez-Colome, A. M., Characterization of a NADPH-dependent coenzyme A-SS-glutathione reductase from yeast, Biochim. Biophys. Acta, 191, 239, 1969. 224. Loewen, P. C., Identification of a coenzyme A-glutathione disulfide (DSI), a modified coenzyme A disulfide (DSII), and a NADPH-dependent coenzyme A-glutathione disulfide reductase in E. coli, Can. J. Biochem., 55, 1019, 1977.

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361. Brown, N. L., Bacterial resistance to mercury- reduciio ad absurdum?, Trends Biochem. Sci., 10, 400, 1985. 362. Silver, S. and Misra, T. K., Plasmid-mediated heavy metal resistances, Ann. Rev. Microbiol., 42, 717, 1988. 363. Walsh, C. T., Distefano, M. D., Moore, M. J., Shewchuk, L. M., and Verdine, G. L., Molecular basis of bacterial resistance to organomercurial and inorganic mercuric salts, FASEB J., 2, 124, 1988. 364. Cheesman, B. V., Arnold, A. P., and Rabenstein, D. L., Nuclear magnetic resonance studies of the solution chemistry of metal complexes. XXV. Hg(thiol)3 complexes and Hg(II)-thiol ligand exchange kinetics, J. Am. Chem. Soc., 110, 6364, 1988. 365. Rabenstein, D. L. and Fairhurst, M. T., Nuclear magnetic resonance studies of the solution chemistry of metal complexes. XI. The binding of methylmercury by sulfhydryl-containing amino acids and by glutathione, J. Am. Chem. Soc., 97, 2086, 1975. 366. Wang, Y., Mahler, L, Levinson, H. S., and Halvorson, H. O., Cloning and expression in Escherichia coli of chromosomal mercury resistance genes from a Bacillus sp, /. Bacteriol., 169, 4848, 1987. 367. Griffin, H. G., Fosterm, T. J., Silver, S., and Misra, T. K., Cloning and DNA sequence of the mercuricand organomercurial-resistance determinants of plasmid pDU1358, f roc. Natl. Acad. Sci. U.S.A., 84, 3112, 1987. 368. Booth, J. E. and Williams, J. W., The isolation of a mercuric ion-reducing flavoprotein from Thiobacillus ferrooxidans, J, Gen. Microbiol, 130, 725, 1984. 369. Inoue, C., Sugawara, K., Shiratori, T., Kusano, T., and Kitagawa, Y., Nucleotide sequence of the Thiobacillus ferrooxidans chromosomal gene encoding mercuric reductase, Gene, 84, 47, 1989. 370. Bogdanova, E. S., Mindlin, S. Z., Kalyaeva, E. S., and Nikiforov, V. G., The diversity of mercury reductases among mercury-resistant bacteria, FEES Lett., 234, 280, 1988. 371. Jobling, M. G., Peters, S. E., and Ritchie, D. A., Restriction pattern and polypeptide homology among plasmid-borne mercury resistance determinants, Plasmid, 20, 106, 1988. 372. Bogdanova, E. S. and Mindlin, S. Z., Two structural types of mercury reductases and possible ways of their evolution, FEBS Lett., 247, 333, 1989. 373. Veldhuisen, G., Van Dijk, M., Meijer, J., Enger-Vaulk, B. E., and Pouwels, P. H., Transient expressions in mammalian cells of the bacterial reporter gene encoding mercuric reductase: effects of various regulatory elements, Gene, 71, 381, 1988. 374. Sahlman, L., Lindskog, S., Lambier, A.-M., and Dunford, B., The reaction between NADPH and mercuric reductase, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 508, 1984. 375. Sands trom, A. and Lindskog, S., Rapid-scan stopped-flow studies of the flavoenzyme mercuric reductase during catalytic turnover, Eur. J. Biochem., 173, 411, 1988. 376. Sandstrom, A. and Lindskog, S., Activation of mercuric reductase by the substrate NADPH, Eur. J. Biochem., 164,243, 1987. 377. Rinderle, S. J., Booth, J. E., and Williams, J. W., Mercuric reductase from R-plasmid NR1: characterization and mechanistic study, Biochemistry, 22, 869, 1983. 378. Miller, S. M., Ballou, D. R, Massey, V., Williams, C. H., Jr., and Walsh, C. T., Two-electron reduced mercuric reductase binds Hg(II) to the active site dithiol but does not catalyze Hg(II) reduction, J, Biol, Chem., 261, 8081, 1986. 379. Marshall, J. L., Booth, J. E., and Williams, J. W., Characterization of the covalent mercury (II)-NADPH complex, J. Biol. Chem., 259, 3033, 1984. 380. Carlberg, I., Sahlman, L., and Mannervik, B., The effect of 2,4,6-trinitrobenzene-sulfonate on mercuric reductase, glutathione reductase and lipoamide dehydrogenase, FEBS Lett., 180, 102, 1985. 381. Stanisich, V. A., Bennett, P. M., and Richmond, M. H., Characterization of a translocation unit encoding resistance to mercuric ions that occurs on a nonconjugative plasmid in Pseudomonas aeruginosa, J. Bacteriol., 129, 1227, 1977. 382. Moore, M. J. and Walsh, C. T., Mutagenesis of the N- and C-terminal cysteine pairs of Tn501 mercuric reductase: consequences for bacterial detoxification of mercurials, Biochemistry, 28, 1183, 1989. 383. Distefano, M. D., Au, K. G., and Walsh, C. T., Mutagenesis of the redox-active disulfide in mercuric ion reductase: catalysis by mutant enzymes restricted to flavin redox chemistry, Biochemistry, 28, 1168, 1989. 384. Massey, V., Miller, S. M., Ballou, D. P., Williams, C. H., Jr., Moore, M., Distefano, M., and Walsh, C. T., Studies on the active site of mercuric reductase employing site-directed mutants and the thiol-reactive flavin, 6-SCN-FAD, in Flavins and Flavoproteins, McCormick, D. B. and Edmondson, D. E.,Eds., Walter de Gruyter, Berlin, 41, 1987. 385. Massey, V., Ghisla, S., and Yagi, K., 6-Thiocyanatoflavins and 6-mercaptoflavins as active-site probes of flavoproteins, Biochemistry, 25, 8103, 1986. 386. Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C. T., Evidence for the participation of Cys558 and Cys559 at the active site of mercuric reductase, Biochemistry, 28, 1194, 1989.

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387. Kao, P. N. and Karlin, A., Acetycholine receptor binding site contains a disulfide cross-link between adjacent half-cystinyl residues, J. Biol Chem., 261, 8085, 1986. 388. Capasso, S., Mattia, C., and Mazzarella, L., Structure of a c/s-peptide unit: molecular conformation of the cyclic disulphide L-oysteinyl-L-cysteine, Acta Cryst., 33, 2080, 1977. 389. Miller, S. M., Moore, M. J., Massey, V., and Walsh, C. T., Unpublished results. 390. Distefano, M. D., Moore, M. J., and Walsh, C. T., Active site of mercuric reductase resides at the subunit interface and requires Cysn5 and Cys!40 from one subunit and Cys558 and Cys559 from the adjacent subunit: evidence from in vivo and in vitro heterodimer formation, Biochemistry, 29, 2703, 1990. 391. Stankovich, M. T. and Bard, A. J., The electrochemistry of proteins and related substances. I. Cystine and cysteine at the mercury electrode, /. Electroanal. Chem., 75, 487, 1977. 392. Stricks, W. and Kolthoff, I. M., Reactions between mercuric mercury and cysteine and glutathione. Apparent dissociation constants, heats and entropies of formation of various forms of mercuric mercaptocysteine and -glutathione, J. Am. Chem. Soc., 75, 5673, 1953. 393. Casas, J. S. and Jones, M. M., Mercury (II) complexes with sulfhydryl containing chelating agents: stability constant inconsistencies and their resolution, /. Inorg. NucL Chem., 42, 99, 1980. 394. Gopinath, E., Kaaret, T. W., and Bruice, T. C., Mechanism of mercury(II) reductase and influence of ligation on the reduction of mercury(II) by a water soluble 1,5-dihydroflavin, Proc. Nat. Acad. Sci. U.S.A., 86, 3041, 1989. 395. Venkataram, U. V. and Bruice, T. C., On the mechanism of flavin-catalyzed dehydrogenation a,p to an acyl function. The mechanism of 1,5-dihydroflavin reduction of maleimides, /. Am. Chem. Soc., 106, 5703, 1984. 396. Fair lamb, A. H. and Cerami, A., Identification of a novel, thiol-containing co-factor essential for glutathione reductase enzyme activity in trypanosomatids, Mol. Biochem. Parasitology, 14, 187, 1985. 397. Fairlamb, A. H., Blackburn, P., Ulrich, P., Chait, B. T., and Cerami, A., Trypanothione: a novel bis(glutathionyl)spermidine cofactor for glutathione reductase in trypanosomatids, Science, 227, 1485, 1985. 398. Shames, S. L., Fairlamb, A. H., Cerami, A., and Walsh, C. T., Purification and characterization of trypanothione reductase from Crithidia fasciculata, a newly discovered member of the family of disulfidecontaining flavoprotein .reductases, Biochemistry, 25, 3519, 1986. 399. Krauth-Siegel, L., Enclers, B., Henderson, G. B., Fairlamb, A. H., and Schirmer, R. H., Trypanothione reductase from Trypanosoma cruzi. Purification and characterization of the crystalline enzyme, Eur. J. Biochem., 164, 123, 1987. 400. Henderson, G. B., Fairlamb, A. H., Ulrich, P., and Cerami, A., Substrate specificity of the flavoprotein trypanothione disulfide reductase from Crithidia fasciculata, Biochemistry, 26, 3023, 1987. 401. Kuwahara, T., White, R. A., Jr., and Agosin, M., A cytosolic FAD-containing enzyme catalyzing cytochrome c reduction in Trypanosoma cruzi. I. Purification and some properties, Arch. Biochem. Biophys., 239, 18, 1985. 402. Kuwahara, T., White, R. A., Jr., and Agosin, M., A cytosolic flavin-containing enzyme catalyzing reduction of cytochrome c in Trypanosoma cruzi: kinetic studies with cytochrome c as substrate, Arch. Biochem. Biophys., 241, 45, 1985. 403. Schirmer, R. H., Lederbogen, F., Krauth-Siegel, R. L., Eisenbrand, G., Schultz, G., and Jung, A., Flavoenzymes as drug targets, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 847, 1984. 404. Docampo, R. and Stoppani, A. O. M., Generation of superoxide anion and hydrogen peroxide induced by nifurtimox in Trypanosoma cruzi, Arch. Biochem. Biophys., 197, 317, 1979. 405. Henderson, G. B., Ulrich, P., Fairlamb, A. H., Rosenberg, I., Pereira, M., Sela, M., and Cerami, A., "Subversive" substrates for the enzyme trypanothione disulfide reductase: alternative approach to chemotherapy of Chagas disease, Proc. Natl. Acad. Sci. U.S.A., 85, 5374, 1988. 406. Jockers-Scherubl, M. C., Schirmer, R. H., and Krauth-Siegel, R. L., Trypanothione reductase from Trypanosoma cruzi. Catalytic properties of the enzyme and inhibition studies with trypanocidal compounds, Eur. J. Biochem., 180, 267, 1989. 407. Sullivan, F. X., Krauth-Siegel, R. L., Pai, E. F., and Walsh, C. T., Molecular approaches in analysis of the substrate specificity of trypanothione reductase, a flavoprotein from trypanosomatid parasites, in Protein and Pharmaceutical Engineering, UCLA Symp. Mol. Cell. Biol., Craik, S., Fletterick, R., Matthews, C. R., and Wells, J., Eds., A. R. Liss, Inc., New York, 1990, 119. 408. Sundquist, A. R. and Fahey, R. C., The function of ^-glutamylcysteine and bis-^-glutamylcystine reductase in Halobacterium halobium, J. Biol. Chem., 264, 719, 1989. 409. Romano, A. H. and Nickerson, W. J., Cystine reductase of pea seeds and yeasts, J. Biol. Chem., 208, 409, 1954. 410. Carroll, J. E., Kosicki, G. W., and Thibert, R. J., a-Substituted cystines as possible substrates for cystine reductase and L-arnino acid oxidase, Biochim. Biophys. Acta, 198, 601, 1970. 411. Yanagawa, H. and Egsimi, F., Asparagusate dehydrogenases and lipoyl dehydrogenase from asparagus mitochondria Physical, chemical, and enzymatic properties, J. Biol. Chem., 251, 3637, 1976.

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412. de la Motte, R. S. and Wagner, F. W., Aspergiilus niger sulfhydryl oxidase, Biochemistry, 26, 7363, 1987. 413. de la Motte, R. S. and Wagner, F. W., Aspergiilus niger sulfhydryl oxidase: a flavoprotein possessing a catalytically essential half-cystine, Fed. Proc., 46, 3529, 1987. 414. Pazur, J. H., Kleppe, K., and Cepure, A., A glycoprotein structure for glucose oxidase from Aspergiilus niger, Arch. Biochem. Biophys., I l l , 351, 1965. 415. Massey, V., Miiller, F., Feldberg, R., Schuman, M., Sullivan, P. A., Howell, L. G., Mayhew, S. G., Matthews, R. G., and Foust, G. P., The reactivity of flavoproteins with sulfite: Possible relevance to the problem of oxygen reactivity, J. BioL Chem., 244, 3999, 1969. 416. Ostrowski, M. C. and Kistler, W. S., Properties of a flavoprotein sulfhydryl oxidase from rat seminal vesicle secretion, Biochemistry, 19, 2639, 1980. 417. Dolin, M. I., The Streptococcus faecalis oxidases for reduced diphosphopyridine nucleotide. III. Isolation and properties of a flavin peroxidase for reduced diphophopyridine nucleotide, J. BioL Chem., 225, 557, 1957. 418. Hoskins, D. D., Whiteley, H. R., and Mackler, B., The reduced diphosphopyridine nucleotide oxidase of Streptococcus faecalis: purification and properties, J. BioL Chem., 237, 2647, 1962. 419. Jacobs, N. J. and Vandemark, P. J., The purification and properties of the oc-glycerophosphate-oxidizing enzyme of Streptococcus faecalis 10C1, Arch. Biochem. Biophys., 88, 250, 1960. 420. Claiborne, A., Studies on the structure and mechanism of Streptococcus faecium L-a-glycerophosphate oxidase, J. BioL Chem., 261, 14398, 1986. 421. Dolin, M. I., The Streptococcus faecalis oxidases for reduced diphosphopyridine nucleotide. IV. Properties of the enzyme-substrate complex formed between reduced diphosphopyridine nucleotide peroxidase and pyridine nucleotides, /. BioL Chem., 235, 544, 1960. 422. Dolin, M. I., Reduced diphosphopyridine nucleotide peroxidase. Intermediates formed on reduction of the enzyme with dithionite or reduced diphosphopyridine nucleotide, J. BioL Chem., 250, 310, 1975. 423. Poole, L. B. and Claiborne, A., Interactions of pyridine nucleotides with redox forms of the flavincontaining NADH peroxidase from Streptococcus faecalis, J. BioL Chem., 261, 14525, 1986. 424. Schmidt, H. L., Stocklem, W., Danzer, J., Kirch, P., and Limbach, B., Isolation and properties of an H2O-forming NADH oxidase from Streptococcus faecalis, Eur. J. Biochem., 156, 149, 1986. 425. Poole, L. B. and Claiborne, A., The non-flavin redox center of the streptococcal NADH peroxidase. I. Thiol reactivity and redox behavior in the presence of urea, /. BioL Chem., 264, 12322, 1989. 426. Ahmed, S. A. and Claiborne, A., The streptococcal flavoprotein NADH oxidase. I. Evidence linking NADH oxidase and NADH peroxidase cysteinyl redox centers, J. BioL Chem., 264, 19856, 1989. 427. Poole, L. B. and Claiborne, A., The non-flavin redox center of the streptococcal NADH peroxidase. II. Evidence for a stabilized cysteine-sulfenic acid, J. BioL Chem., 264, 12330, 1989. 428. Poole, L. B. and Claiborne, A., Evidence for single active-site cysteinyl residue in the streptococcal NADH peroxidase, Biochem. Biophys. Res. Commun., 153, 261, 1988. 429. Stoll, V. S. and Blanchard, J. S., Kinetic mechanism and nucleotide specificity of NADH peroxidase, Arch. Biochem. Biophys., 260, 752, 1988. 430. Schiering, N., Stoll, V. S., Blanchard, J. S., and Pai, E. F., Crystallization and preliminary X-ray diffraction study of the flavoprotein NADH proxidase from Streptococcus faecalis 10C1, /. BioL Chem., 264, 21144, 1989. 431. Ahmed, S. A. and Claiborne, A M The streptococcal flavoprotein NADH oxidase. II. Interactions of pyridine nucleotides with reduced and oxidized enzyme forms, J. BioL Chem., 264, 19864, 1989.

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Chapter 6

REFINED THREE-DIMENSIONAL STRUCTURE OF GLUTATHIONE REDUCTASE P. Andrew Karplus and George E. Schulz

TABLE OF CONTENTS I.

Introduction

214

II.

Materials and Methods

214

III.

Results and Discussion A. The Polypeptide Structure B. The Dimer Interface C. The FAD-Binding Site D. The Active Center

215 215 218 220 222

IV.

Conclusions

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References

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I. INTRODUCTION Glutathione reductase (EC. 1.6.4.2) catalyzes the reduction of glutathione disulfide (GSSG) at the expense of NADPH: GSSG + NADPH + H + ^± 2 GSH + NADP + The enzyme maintains a high ratio of [GSH]/[GSSG] in the cell and is as ubiquitous as its substrates.1 The enzyme is a member of a family of flavin-containing pyridine nucleotide:disulfide oxidoreductases.1-2 which also includes lipoamide dehydrogenase, thioredoxin reductase, and mercuric reductase. Structural similarities with the flavoenzymes respiratory NADH-oxidoreductase of Escherichia colP and /?-hydroxybenzoate hydroxylase4 have been observed. Glutathione reductase from human erythrocytes5-6 is a dimer of two identical subunits.7 Each subunit contains 478 residues and one FAD molecule with a total Mr of 52,400.8 The three-dimensional structure of the holoenzyme has been determined at a resolution of 2 A, using the multiple isomorphous replacement technique.9 Apart from the N-terminal 17 residues, which are invisible in the crystal structure and therefore presumably flexible, the whole polypeptide chain could be assigned to the electron density map. In the crystal, the substrate binding sites of glutathione reductase are not blocked and the catalytic activity is conserved.10~12 Additionally, the crystals are well ordered and allowed data collection beyond a 2 A resolution, rendering this system well suited for a detailed analysis of enzyme-substrate interactions and catalysis. Today, data have been measured and the enzyme structure has been crystallographically refined at a resolution of 1.54 A. 13

II. MATERIALS AND METHODS Glutathione reductase was prepared from outdated human-blood conserves following an established protocol.6 Crystals corresponding to form-B9 were grown from 1.2 M ammonium sulfate in 0.1 M potassium phosphate buffer at pH 7.0 and 4°C. For storage purposes and data collection, the ammonium sulfate concentration was increased to 2.0 M. Crystal formB belongs to space group B2 with unit cell parameters a = 119.8 A, b = 84.5 A, c — 63.2 A, y = 58.7 A. Out to a resolution of 1.54 A, the data were collected from about 20 crystals with sizes of approximately 1.3 mm X 0.8 mm x 0.6 mm using a 4-circle diffractometer. On the average, reflections were measured after the crystals had suffered from radiation damage equivalent to an intensity drop of 14%. The final data set consisted of 78,959 independent reflections, 1.2% of which were zero. The crystallographic refinement of glutathione reductase has been carried out using a restrained least-squares program package.14 For calculating the geometric term we used an amino acid residue dictionary containing standard bond lengths and angles.14 The standard geometry of FAD was composed of several pieces: The adenosine moiety was taken from the crystal structure of AMP;15 the isoalloxazine moiety was taken from the "ideal" lumiflavin geometry;16 the ribityl moiety was built up using C-C and C-O bond lengths of 1.53 A and 1.43 A, respectively, together with regular tetrahedral angles; the pyrophosphate moiety was derived by averaging results from four crystal structures.15'1719 The refinement started with a model which had been built at 2 A resolution assigning initial temperature factors of 12 A2 for each atom. When the parameter shifts became small enough, the refinement was stopped. The R-factor for all data from 10 to 1.54 A was 18.6%. A vivid impression of model quality can be obtained from the final 2F0-FC map. Figure 1 shows 2 cuts through this map, one in the plane of the isoalloxazine moiety and another in the plane of the adenine moiety of the prosthetic group FAD. In isoalloxazine, all (non-

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FIGURE 1. Quality of the final 2F0-FC electron density map.13 (a), Electron density in the best plane of the isoalloxazine moiety of FAD with contour levels at 0.3, 0.6, . . . , 2.7 e/A3. Stippled contour lines go downward forming holes in the ring centers. The final model is drawn out using no dots, small dots and large dots at carbon, nitrogen and oxygen positions, respectively, (b), Electron density in the best plane of the adenine moiety of FAD, applying the same symbols as in (a). (From Karplus, P. A. and Schulz, G. E., J. Mol BioL, 195, 701, 1987. With permission.)

hydrogen) atoms are clearly visible, although not separated by density incisions. All three ring centers are hollow, with the pteridine center decreasing to the lowest contour level. In adenine, the six-membered ring has a deep hole in its center, whereas the five-membered ring has not. Also, a distinction of atom type is noticeable as the oxygen atoms show, on the average, higher density than the nitrogen atoms, which in turn show higher density than the carbon atoms. In general, the final F0-FC map showed positive peaks at theoretical hydrogen positions. As these peaks were below the noise level, they were not included in the model. For the analysis of hydrogen bonds, we generated hydrogen atoms from known amino acid geometry.20

III. RESULTS AND DISCUSSION We are here concerned with the structure of the crystalline enzyme as observed at the rather high ionic strength, I = 6, of a 2 M ammonium sulfate solution. Furthermore, we are concerned with the enzyme in its oxidized state containing the prosthetic group FAD but no ligands.13 A. THE POLYPEPTIDE STRUCTURE There are 458 established trans peptide bonds in glutathione reductase. Their mean torsion angle is CD = 179.85° with a spread of 3.5°. The deviation from 180° has the same sense but is much smaller than the 1° deviation connected with a spread of 7° reported for a number of trypsin-related enzymes.21 There are two cis peptide bonds in glutathione reductase. These occur before Pro-375 and before Pro-468. Both proline residues have \angles near 150° and are in position i + 2 of reverse turns without hydrogen bonds. Pro-375 is in a tight reverse turn of the antiparallel sheet E in the protein interior. Pro-468 is next to His-467, which is directly involved in catalysis. The connecting cis peptide is stabilized by a strong hydrogen bond from the N(3) atom of the isoalloxazine of FAD to 467'-O. Since secondary structures are hydrogen-bonded blocks, their assignment depends critically on the definition of a hydrogen bond. In glutathione reductase we applied the distance

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criterion H . . . O < 2.5 A without consideration of the angle of approach, which is similar to a cutoff used earlier.22 Within the a-helix segments, we distinguished between core and noncore residues, which form two and one hydrogen bonds in a helix, respectively. The average conformational angles (4*,v|/) and hydrogen-bond parameters are similar for core ( — 63°, — 42°) and noncore ( — 65°, — 41°) residues, and do not deviate from the data of other proteins.22*23 The standard deviations, however, are much larger in the noncore parts (12°, 10°) than in the cores (4°,4°), showing that at both ends the a-helices taper out in more irregular conformations. An appreciable number of residues in glutathione reductase are involved in (3-pleated sheet formation. By definition, these sheets comprise all residues, regardless of conformation, which are interconnected by the appropriate pattern of hydrogen bonds. The conformational statistics and the inset in Figure 2 show that the average (cf>,v|>) angles of these sheets scatter around the "twist" position with a slight segregation of the parallel (A,B) and antiparallel (B,D,E) sheet positions. As shown in Figure 3, sheet E is of a rather special nature. The largest part of this antiparallel sheet is as planar as the originally suggested model.24 In addition, it contains a (3-bulge25 at Val-431. In addition to a-helices and ^-sheets, proteins contain 3]0-helices and reverse turns, with 310-helices being a subset of reverse turns. In glutathione reductase, we observe six of these helices, consisting of residues 170 to 174, 177 to 183, 299 to 304, 330 to 334, 405 to 409, and 470 to 476. Besides these, there are several 310-helices at the C-terminal ends of a-helices. As a general observation, we find that the 310-helix is a prominent conformation of the enzyme. The first 17 residues are not visible in the electron density map, indicating a high degree of mobility. Both the N- and C-terminal ends of the visible chain have higher than normal mobilities, as observed with many proteins.26 In general, there is a good correlation between B-factor and solvent accessibility of the chain. The overall average temperature factor of the refined model is 22.4 A 2 , while the average B-factor of the main chain is 18 A2. There are six segments (residues 58 to 70, 196 to 203, 338 to 349, 369 to 378, 433 to 446, and 465 to 472) where the B-factors drop below 10 A2. These segments contain the two cis proline residues (375, 468), many residues involved in the dimer interface as well as crucial residues at the catalytic center (Cys-58, Cys-63, Lys-66, Tyr-197, Glu-201, Val-370, His467' and Glu-472').* The catalytically competent isoalloxazine ring has an average B-factor of 8.7 A2, whereas the ribityl, phosphate, ribose and adenine moieties have average Bfactors of 10.4 A 2 , 12.6 A 2 , 12.5 A2, and 16.2A2, respectively. Again, the most important part is most thoroughly fixed. All solvent molecules are numbered according to their maximal electron densities in the final 2F0-FC map. The lowest numbers correspond to the highest densities. A lower cutoff of 0.6 e/A3 was applied, which restricted the total number of solvent molecules modelled as water to 523. The largest B-factor is 49 A2. Since the analyzed crystals contained 42% solvent based on a protein density of 1.35 g/cm3, the located solvent represents 21% of the total solvent in the unit cell. Recent results27 indicate that solvent molecules with occupancies above 0.8 and Bfactors below 25 A2 can be considered integral parts of a protein. These limits correspond roughly with the electron density cutoff of 1.4 e/A3. With this cutoff, we find 118 integral water molecules per subunit. Thus, the holoenzyme consists of about 97% polypeptide, 2% solvent and 1% prosthetic group. Since glutathione reductase crystals were analyzed in 2 M ammonium sulfate with 0.1 M phosphate buffer, one has to expect tightly bound cations and anions. Among the 118 integral water molecules, however, there was none that could be sensibly fitted as an ammonium ion into its surrounding hydrogen-bond pattern, showing that a molar ratio of about 1:14, ammonium ions cannot effectively compete with water for protein atoms. Anion *

Primed residue numbers refer to the other subunit.

FIGURE 2. Strand arrangements in the p-pleated sheets of glutathione reductase.13 The lengths of all hydrogen bonds are given with the H ... O distances (in A). The numbers of the first and the last residue of each strand are inserted. The average (,i|j)-angles for the internal residues of the sheets A to E are plotted in the inset. The ideal positions for parallel (p), antiparallel (a) and twisted (t) sheets are given. (From Karplus, P. A. and Schulz, G. E., J. MoL Biol., 195, 701, 1987. With permission.)

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. Stereo plot of the backbone of sheet E of the interface domain (residues 366 to 437).13 The major part of this antiparallel sheet is planar (see also Figure 4b). The upper part of the sheet is twisted as usual. The predominant planarity of sheet E is also expressed in the average (4>,i|/)-angles (Figure 2, inset), which place E nearest to the position of the theoretical planar antiparallel (3-sheet. Chain breaks are marked by dots, (From Karplus, P. A. and Schulz, G. E., J. MoL Biol, 195, 701, 1987. With permission.)

binding to glutathione reductase has been observed in the initial 3 A resolution map at the binding site of the 2'-phosphate of NADP.28 This anion was originally modeled as a sulfate, but the B-factor for the sulfur atom refined to an unreasonably high value in comparison with the B-factors of the oxygen atoms. Changing to a phosphate ion equalized the B-factors, suggesting that a phosphate ion is bound despite the fact that sulfate predominates by a factor of 20. Surprisingly, neither the phosphate nor the interacting side chains have turned out to be particularly well-ordered. They have B-factors near 35 A2. The assigned solvent molecules can be subdivided into clusters, which are defined as sets of solvent molecules that form at least one contact closer than 3.5 A with another member of the group. The largest cluster fills the cavity at the molecular dyad of glutathione reductase and comprises 2 x 5 1 = 102 molecules. There are three further large clusters, one in the crevice where GSSG binds (42 water molecules), another in the region of the bound adenosine moiety of FAD (30 water molecules), and a third at the NADPH binding site (25 water molecules). Based on the unrefined coordinates of bound GSSG and NADPH, these substrates replace approximately 15 water molecules each. In both cases, the majority of displaced water molecules is only weakly bound. Presumably, the concave nature of the substrate binding site facilitates the formation of larger clusters at these parts of the protein surface. B. THE DIMER INTERFACE Glutathione reductase forms a homo-dimer in which both subunits participate in both

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FIGURE 4. The dimer interface of glutathione reductase.13 (a), CA backbone of the complete dimer viewed from along the dimer axis; (b) CA backbone including residues 53 to 104 and 367 to 478 from each monomer. The right-hand monomer is drawn with thick lines and the "X" marks the center of the large solvent-filled activity. Viewing direction is from the left-hand side of (a), (c), Schematic drawing of the upper and lower interface regions. Viewing direction is from the right-hand side of (b). The placement of positive and negative charged atoms that form H-bonds upon dimerization is indicated. (From Karplus, P. A. and Schulz, G. E., J. Mol. Biol., 195, 701, 1987. With permission.)

catalytic centers. In particular, the substrate GSSG is bound between subunits and it is processed by side-chains extending from both subunits,10 e.g., Cys-58 and His-467'. As a consequence, the interface has to be considered an integral part of the enzyme; it is not conceivable that isolated subunits could show enzymatic activity. Two stereo views of this interface are depicted in Figure 4a and 4b. They show that the interface area can be subdivided into an upper and a lower part. These parts are separated by a cavity with channel extensions to the solvent, as sketched in Figure 4c. The upper part is mainly formed by the so-called interface domains of the enzyme, whereas the lower part involves the long helical extensions

220

Chemistry and Biochemistry of Flavoenzymes

FIGURE AC.

of the FAD-binding domains (residues 70 to 110). The protein-free cavity at the molecular dyad separating these areas can be approximated by an ellipsoid of approximate volume 1500 A3. Since this cavity is rather close to the glutathione binding site, it is tempting to speculate that effector binding in this cavity could play a role in enzyme regulation. The total interface area was calculated as 3400 A2, which amounts to 15% of the total surface of the monomer. Adding up the changes in accessible surface areas for the appropriate residues leads to estimates of 2100 A2 and 1200 A2, as areas of the upper and lower interfaces, respectively. As both interface parts are of comparable size they should, as a first estimate, have comparable binding energies.29 However, the nature of the association in the two regions is quite different, such that the lower part of the interface probably contributes much less than the upper part. The upper interface moiety is one of the least mobile regions of the whole protein, whereas the lower part of the interface is at the other end of the mobility scale. Here, residues 88 to 93 reach the highest B-factors of the whole model. The main-chain atoms of Cys-90, which forms an intersubunit disulfide bridge, have an average B-value as high as 50 A2. Also, there are some indications that alternative conformations do exist for Met-79 and Phe78 as well as for the contacting pair His-75 . . . His-82'. The sequence comparison between the enzyme from man and E. coli30 shows a very strong conservation at the upper interface. In this region, 92% of all residues are identical, as compared with 52% overall identity. In contrast, the lower interface has only 37% identical residues, which is less than average. Also, the intersubunit disulfide bridge Cys-90:Cys-90' is missing in the E. coli enzyme. In addition, the homologous sequence of the lipoamide dehydrogenase from E. coli has been related to the glutathione reductase.31 Here, we find 25% and 29% identical residues at the lower and upper interfaces, respectively. These values approach the general level of homology between lipoamide dehydrogenase and glutathione reductases, and therefore demonstrate that during protein differentiation the whole interface has not been particularly conserved, although the general geometry of the dimer is presumably identical.32 C. THE FAD-BINDING SITE As discussed above, the environment of the prosthetic group FAD is particularly rigid

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and therefore particularly well defined. The conformational angles of FAD, together with atom names, are given in Figure 5. The conformation of the ribityl moiety is fully extended, having all torsion angles within 5° of the trans position. An approximately extended conformation is also assumed by the torsion angles at the C-O bonds to the pyrophosphate, 6 and 9 A . The pyrophosphate itself is less extended. The angles v|/, , if/A and A are all closer to gauche than to trans, causing the chain to bend. Adenine assumes the favorable anti conformation, and the ribose pucker is close to the 2' endo conformation. The isoalloxazine ring appears to be quite planar, despite the fact that planarity restraints were applied only to the pteridine and to the benzene rings, allowing butterfly bending around the N(5) to N(10) virtual axis during refinement. The best planes of the two rings deviate from one another by 3.3%. This can be divided into a 2.6° bending toward the redox active disulfide around the N(5) to N(10) virtual axis, and 1.9° twisting around the long dimension of the isoalloxazine ring system. Deviation from planarity of this magnitude has been also observed in the crystal structure of 10-methyl isoalloxazine.16 The refined bond lengths of the flavin, on the other hand, show some small but possibly important deviations from standard geometry. There are five bonds, all near the redox active part of the flavin, which differ by more than one root mean square deviation (= 0.025 A) from their standard values. These differences include a lengthening of the double bonds to N(l) and N(5) and are summarized in Figure 5. Although the significance of this deviation is questionable, it may signal an influence of the polypeptide on to the electronic structure of the flavin. Figure 6 gives a stereo view of the complete FAD binding site. The geometries of all hydrogen bonds involved in FAD binding are listed in Table 1. As can be seen in Figure 6, the binding of FAD to the apoenzyme involves many water molecules, eight of which form direct hydrogen bonds to FAD atoms. In the following, some major aspects of the binding will be discussed. The binding of the pyrophosphate group has attracted particular interest because of the lack of compensating positively charged side chains nearby. For charge compensation of the phosphates, it has been suggested33 that the electron densities of Sol-2 and Sol-4 could represent cations. The refined structure showed, however, that Sol-2 and Sol-4 are indeed unprotonated water molecules, because their occupancies refined to 100% and their B-factors to around 10 A 2 , as is appropriate for the environment. Furthermore, each site has in its environment two hydrogen-bond donors and two hydrogen-bond acceptors in a tetrahedral arrangement. The dipole nature of the a-helix beginning at residue 29 may provide partial compensation of the negative charge. It has been suggested that the electrostatic potential of an a-helix arises mainly from the four peptide amide groups of the first a-helix turn.34 In glutathione reductase, we observe that OF1 interacts either directly or through Sol-4 with three of the first four amides of the helix residues beginning at position 29, and thus, very likely experiences significant charge compensation. OF2 is involved in a geometrically similar hydrogen-bonded network with Sol-2, but without a helix. Through its hydrogen bond to Sol-10, OA1 makes an indirect contact to the charged side chain of Arg-291. However, a complete charge compensation by Arg-291 is not possible, because Sol-10 also binds to the carboxyl of Asp-331. The adenine moiety is rather well connected to the protein through hydrogen bonds between N(1A) and N(6,aA), and the backbone at residue 130. The ribose forms very tight hydrogen bonds to the carboxyl group of Glu-50, which is also hydrogen bonded to 28-N and 52-N. Ribose binding to a carboxyl is known from almost all nucleotide-binding proteins.35 Among the free hydroxyl groups of the ribityl, only the middle one is very strongly bound; namely, to the carboxyl group of Asp-331, which is inaccessible to solvent. In contrast, O2' and O4' are involved in intra-FAD hydrogen bonds.

222

Chemistry and Biochemistry of Flavoenzymes

FIGURE 5. Nomenclature33 and observed conformation of FAD.13 The refined values for the torsion angles are given. Also given are the observed bond lengths of the flavin that deviate more than 0.02 A from the standard values.16 The deviation is indicated in parentheses. (From Karplus, P. A. and Schulz, G. E., /. Mol. Biol, 195, 701, 1987. With permission.)

The isoalloxazine is tightly bound with its N(3) atom to His-467'-O of the other subunit. The good hydrogen bond donor property of N(3) has been suggested for other flavoenzymes on the basis of Raman spectroscopic data.36 O(2a) forms a short hydrogen bond with Sol15, and another with 339-N at the start of an a-helix, the dipole moment of which could stabilize a partial negative charge at O(2a). In the environment of N(5) and O(4a) there is a net of hydrogen-bonded atoms (Figure 6). Here one finds a tightly held solvent molecule, Sol-18, which has to leave on binding of nicotinamide. The hydroxyl of Tyr-197 contacts N(10) (O . . . N = 3.15 A, H . . . H - 2.52 A, O-H . . . N = 121°) without forming a clear hydrogen bond; Tyr-197 has been considered as a lid for protecting the reduced enzyme against oxidation.10 D. THE ACTIVE CENTER The active center and the movement of reduction equivalents from NADPH to GSSG

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FIGURE 6. The FAD binding site.13 The segments of polypeptide included in the plot are 26 to 33; 49 to 67; 128 to 130; 155 to 158; 197 to 201; 291 to 298; 329 to 332; 337 to 342; 368 to 370; and 456' to 468'. For each segment, the CA atoms of the first and last residues are labeled, and the initial N and the final C atoms are marked with a dot. Among the 36 solvent molecules ( + ) in the plot, there are 12 (identified by number) involved in either primary or secondary H-bond interactions with FAD (see Table 1). (From Karplus, P. A. and Schulz, G. E., J. Mol. BioL, 195, 701, 1987. With permission.)

has been derived from lower resolution models.10 At 1.54 A resolution further details of interest were added.13 A stereo view of the active center, and a list of some important distances are given in Figure 7 and Table 2, respectively. The side chain of Lys-66 is rather rigid: the temperature factors of CA, CB, CG, CD, CE, and NZ are 9, 12, 14, 13, 13, and 10 A2, respectively. The decrease of mobility towards the end is easily correlated with the formation of an internal salt-bridge to Glu-201. As compared to the data at lower resolution, this side chain has now moved by about 1 A such that the distances from Lys-66-CE and Lys-66-NZ to N5 of the flavin are 4.0 and 3.0 A, respectively. The strength of the interaction between Lys-66-NZ and N(5) is not clear, because the orientation of the lysine ammonium rotor cannot be established unambiguously due to the presence of four possible hydrogen bond acceptors (Table 2). At a distance of 4 A, Lys-66-CE does not sterically hinder the protonation of N(5), as has been suggested earlier.10 Since the distance between N(5) and the proximal sulfur Cys-63-SG is 4.1 A, it becomes conceivable that the flavin does become protonated at N(5) upon reduction by NADPH, and that the proton proceeds to the redox-active disulfide. If such a proton transfer were to take place, it would reduce appreciably the suggested local proton gradient building up during catalysis.37 At lower resolution, the direction of the hydrogen bond from 339-N to either N(l) or O(2a) could not be clarified. In contrast, there is now a clear asymmetry; the distances to O(2ct) and N(l) are 3.1 A and 3.5 A, respectively (Table 1). That N(l) interacts only weakly with the protein is consistent with results from the analysis of FAD analogues in glutathione reductase.38 One may expect that an intermediate negative charge at O(2a) would be stabilized by this hydrogen bond and also by the dipole moment of helix 339 to 352. The importance

224

Chemistry and Biochemistry of Flavoenzymes TABLE 1 Hydrogen Bonding to and around FADa Primary interactions

(A)

D-H . . . A (degree)

N(1A) . . . 130-N N(3A) . . . 51-N N(3A) . . . 51-OG N(7A) . . . Sol-326 N(6«A) . . . 130-0d N(6aA) . . . 129-NDld N(6aA) . . . Sol-326d O2'A . . . 50-OE2 02'A . . . Sol-364 03'A . . . 50-OE1 03'A . . . Sol-54

2.90 3.13 3.33 3.14 3.08 3.39 3.18 2.65 2.51 2.69 2.78

176 138 170 — 150 166 174 177 — 175 —

OA1 . . . 57-OG1 OA1 . . . Sol- 10

2.85 3.10

172 —

OA2 . . . 57-N OA2 . . . 04 OA2 . . . Sol-359 OF1 . . . Sol-4

3.16 2.96 2.88 2.66

140 161 — —

OF1 . . . 31-N OF2 . . . 331-N OF2 . . . SoI-2

2.74 2.97 2.55

165 151 —

04' ... 02' O4' . . . Sol-359 O3' . . . 331-OD2 N(l) . . . 339-Ne 0(2a) . . . 339-N O(2a) . . . Sol- 15

2.65 3.17 2.76 3.49 3.10 2.96

143 — 179 159 147 —

N(3) . . . 467'-O 0(4a) . . . 66-NZf N(5) . . . 66-NZf

2.74 2.78 3.01

159 149 165

Atomsb

a

b 0 d e f

D... A

Secondary D...A Atom

(A)

H-bondsc D-H . . . A (degree)

52-N

2.54

155

28-N 53-O 53-N

2.71 2.89 3.24

152 — 139

331-N 291-NH1

2.76 3.09

— 132

155-O 29-N 32-N

2.88 2.88 2.96

— 131 154

329-O 157-N 332-N

2.80 2.86 3.11

— 140 163

Sol-5

2.76

—

467'-N Sol-3 Sol-30

3.00 2.60 2.82

178 — —

Nomenclature for H-bond parameters: D . . . A = donor-acceptor distance, D-H . . . A = angle of H-bond at H. Nomenclature for FAD atoms is given in Figure 5. FAD atoms listed first, starting at adenine and ending at flavin. Additional strong H-bonds for atoms directly H-bonded to FAD. These 3 H-bonds cannot occur in optimal geometry at the same time. Does not qualify at H-bond, but may be an important interaction. These 2 H-bonds cannot occur in optimal geometry at the same time. Secondary interactions of Lys-66-NZ are given in Table 2.

From Karplus, P. A. and Schulz, G. E., J. MoL BioL, 195, 701, 1987. With permission.

of this helix for the reaction is emphasized by the presence of a nonhomologous a-helix at the same place relative to the isoalloxazine in the flavoenzyme p-hydroxybenzoate hydroxylase.4 During refinement, the S-S bond of the redox-active disulfide remained unrestrained in

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FIGURE 7. The active site of glutathione reductase.13 The geometry is shown for several residues that probably play a role in catalysis.10 Only the riboflavin moiety of FAD is shown. Also included are 4 solvent molecules (Sol-17, Sol-18, Sol-70, and Sol-89) found in the active site region. A number of distances associated with this plot are given in Table 2. (FromKarplus, P. A. and Schulz, G. E., /. MoL BioL, 195, 701,1987. With permission.)

TABLE 2 Hydrogen Bonds and Distances at the Active Center D. . . A

(A)

H. . . A

Atom 2

(A)

D-H . . . A (degree)

66-NZ3 66-NZa 66-NZa 66-NZa 197-OH 467'-NDl 467'-NE2 Sol-18 Sol-70 SoI-89

201-OE1 FAD-0(4a) Sol-18 FAD-N(5) Sol- 17 472'-OE2 Sol-70 201-OE1 58-SG 201-OE2

2.70 2.78 2.96 3.01 2.61 2.77 2.87 2.70 3.36 2.74

1.69 1.86 1.98 2.02 1.63 1.80 2.09 # * *

179 149 161 165 164 164 134 * * *

467'-NE2b 63-SG 63-SG 63-SG 63-SG 63-SG

58-SG 467'-NE2 FAD-N(5) FAD-C(4a) FAD-C(lOa) FAD-N(l)

3.50 4.20 4.14 3.48 3.42 3.52

2.86 — — — —

122 — — — — —

Atom 1

—

a

All 4 H-bonds cannot be present at the same time. Remaining entries are not H-bonded interactions. * Optimal geometry gives an H . . . A distance of 1A less than D . . . A and D-H ... A of 180°.

b

From Karplus, P. A. and Schulz, G. E., /. MoL Biol., 195, 701, 1987. With permission.

order to detect any indication of unusual geometry. However, at 2.06 A, the resulting S-S bond length was normal. The side-chain torsion angles starting from Cys-58 are \i = + 178°, \2 = + 80°,ox3 - 133°, X2 = + H8°, xl = -30°, and the 58-CA to 63-CA distance is only 4.6 A. This conformation is rather unique,39'40 and may be an indication of the special role of this bridge. Also, the main chain around this bridge from 57-O to 64-N seems to be in a strained conformation; four of the six ()-angles are at the borderline of the allowed regions of the Ramachandran plot, and the torsion angles, o>, of all of the peptide bonds are negative, with an average value of w = — 175°. Such a mechanical strain in the S-S loop could help in opening the bridge.

226

Chemistry and Biochemistry of Flavoenzymes

The relative positions of the catalytically important His-467' and the redox-active disulfide are identical to earlier reports.9 The geometry of the contact between His-467'-NE2 and Cys-58-SG does not seem ideal for a direct proton transfer during enzyme reduction. As an addition to the model, now Sol-70 is observed within 0.5 A of the position to be occupied by the proximal sulfur of bound oxidized glutathione. This solvent molecule is hydrogen bonded to His-467 '-NE2 (Table 2), and it is in an ideal position to protonate Cys58 on enzyme reduction. The refinement also confirmed the strong hydrogen bond from His-467 '-ND1 to Glu472'-OE2, which fixes the imidazole. The assigned orientation of the imidazole is well supported by the final 2F0-FC map, in which the ring nitrogen atoms have higher electron density. Allowing for the reported 1 A movement of Cys-58-SG on enzyme reduction,10 it is now observed that the atoms, 472'-OE2, 467'-ND1, 467'-NE2, and 58-SG, are all placed on a straight line at hydrogen-bond distances. This is reminiscent of the "charge relay system" of the serine proteases,41 with the replacement of the serine hydroxyl by a thiol. This analogy is in turn supported by the observation that 58-SG of the reduced enzyme is a strong nucleophile.1

IV. CONCLUSIONS The crystal structure of human glutathione reductase has been refined at 1.54 A resolution. Based on 77,690 independent reflections of better than 10 A resolution, a final Rfactor of 18.6% was obtained with a model obeying standard geometry within 0.025 A in bond lengths and 2.4° in bond angles. Apart from 461 amino acid residues and the prosthetic group FAD, the model contains 524 solvent molecules, about 118 of which can be considered an integral part of the enzyme. The extended FAD molecule shows a mobility gradient between the very rigid flavin ( = 8.7 A2) and the more mobile adenine ( = 16.2 A2). The entire active center is particularly well ordered, with temperature factors around 10 A2. The dimer interface consists of a rigid contact area, which is well conserved in the Escherichia coli enzyme, and a flexible area that is not. The refined structure shows clearly that there are no buried cations compensating the charge of the pyrophosphate moiety of FAD. The flavin deviates slightly from standard geometry, which is possibly caused by the polypeptide environment. Atom N(5) of the flavin can accommodate a proton, and it is conceivable that this proton proceeds to the redox-active disulfide. Cys-58 of this disulfide may be activated in a manner similar to the "charge relay system" of serine proteases.

REFERENCES 1. WUIiams, C. H., Jr., Ravin-containing dehydrogenases, Enzymes, 13, 89, 1976. 2. Brown, N. L., Ford, S. J., Pridmore, R. D., and Fritzinger, D. C., Nucleotide sequence of a gene from the Pseudomonas transposon Tn 501 encoding mercuric reductase, Biochemistry, 22, 4089, 1983, 3. Young, I. G., Rogers, B. L., Campbell, H. D., Jaworowski, A., and Shaw, D. C., Nucleotide sequence coding for the respiratory NADH dehydrogenase of Escherichia coli, Eur. J. Biochem., 116, 165, 1981. 4. Wierenga, R. K., Drenth, J., and Schulz, G. E., Comparison of the three-dimensional protein and nucleotide structure of the FAD-binding domain of /7-hydroxybenzoate hydroxylase with the FAD- as well as NADPH-binding domains of glutathione reductase, /. Mol Biol., 167, 725, 1983. 5. Worthington, D. J. and Rosemeyer, M. A., Glutathione reductase from human erythrocytes, Eur. J. Biochem., 67, 231, 1876. 6. Krohne-Ehrich, G., Schirmer, R. H., and Untucht-Grau, R., Glutathione reductase from human erythrocytes. Isolation of the enzyme and sequence analysis of the redox-active peptide, Eur. J. Biochem., 80, 65, 1977. 7. Schulz, G. E., Zappe, H., Worthington, D. J., and Rosemeyer, M. A., Crystals of human erythrocyte glutathione reductase, FEBS Lett., 54, 86, 1975.

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8. Krauth-Siegel, R. L., Blatterspiel, R., Saleh, M., Schiltz, E., Schirmer, R. H., and Untucht-Orau, R., Glutathione reductase from human erythrocytes. The sequences of the NADPH domain and of the interface domain, Eur, J. Biochem., 121, 259, 1982. 9. Thieme, R., Pai, E. F., Schirmer, R. H., and Schulz, G. E., The three-dimensional structure of glutathione reductase at 2 A resolution, J. Mol BioL, 152, 763, 1981, 10. Pai, E. F. and Schulz, G, E., The catalytic mechanism of glutathione reductase as derived from X-ray diffraction analyses of reaction intermediates, J. Biol. Chem., 258, 1752, 1983. 11. Pai, E. F., Karplus, P. A., and Schulz, G. E., Crystallographic analysis of the binding of NADPH, NADPH fragments, and NADPH analogues to glutathione reductase, Biochemistry, 27, 4465, 1988. 12. Karplus, P. A., Pai, E. F., and Schulz, G. E,, A crystallographic study of the glutathione binding site of glutathione reductase at 0.3 nm resolution, Eur. J. Biochem., 178, 693, 1989. 13. Karplus, P. A. and Schulz, G. E., The refined structure of glutathione reductase at 1.54 A resolution, J, Mol. Biol., 195, 701, 1987. 14. Tronrud, D. E., TenEyck, L. F., and Matthews, B. W., An efficient general-purpose least-squares refinement program for macromolecular strucutres, Acta Crystallogr. Sect. A., 43, 489, 1987. 15. Kraut, J. and Jensen, L. H., Refinement of the crystal structure of adenosine-5'-phosphate, Acta Crystallogr., 16, 79, 1963. 16. Wang, M. and Fritchie, C. J., Jr., Geometry of the unperturbed flavin nucleus. The crystal structure of 10-methylisoalloxazine, Acta Crystallogr. Sect. B, 29, 2040, 1973. 17. Kennard, O., Issacs, N. W., Mother well, W. D. S., Coppola, J. C., Wampler, D. L., Larson, A. C., and Watson, D. G., The crystal and molecular structure of adenosine triphosphate, Proc. Roy. Soc. Ser A, 325, 401, 1971. 18. Parthasarathy, R. and Friday, S. M., Conformational variability of NAD in the free and bound states; A nicotinamide sandwich in NAD + crystals, Science, 226, 969, 1984. 19. Reddy, B. S., Saenger, W., Muhlegger, K., and Weinmann, G., Crystal and molecular structure of the lithium salt of nicotinamide adenine dinucleotide dihydrate (NAD + , DPN + , cozymase, codehydrase I), J. Amer. Chem. Soc., 103, 907, 1981. 20. Momany, F. A., McGuire, R. F., Burgess, A. W., and Scheraga, H. A., Energy parameters in polypeptides. VII. Geometric parameters, partial atomic charges, nonbonded interactions, hydrogen bond interactions, and intrinsic torsional potentials for the naturally occurring amino acids, /. Phys. Chem., 79, 2361, 1975. 21. Marquart, M., Walter, J., Deisenhofer, J., Bode, WM and Huber, R., The geometry of the reactive site and of the peptide groups in trypsin, trypsinogen and its complexes with inhibitors, Acta Crystallogr. Sect. B, 39, 480, 1983. 22. Baker, E. N. and Hubbard, R. E., Hydrogen bonding in globular proteins, Prog. Biophys. Mol. BioL, 44, 97, 1984. 23. Finzel, B. C., Weber, P. C., Hardman, K. D., and Salemme, F. R., Structure of ferricytochrome c' from Rhodospirillum molishianum at 1.67 A resolution, J. Mol. Biol., 186, 627, 1985. 24. Pauling, L. and Corey, R. B., Configurations of polypeptide chains with favored orientations around single bonds: two new pleated sheets, Proc. Nat. Acad. Sci. U.S.A., 37, 729, 1951. 25. Richardson, J. S., Getzoff, E. D., and Richardson, D. C., The 3 bluge: a common small unit of nonrepetitive protein structure, Proc. Nat. Acad. Sci. U.S.A., 75, 2574, 1978. 26. Karplus, P. A. and Schulz, G. E., Prediction of chain flexibility in proteins: a tool for the selection of peptide antigens, Naturwissenschaften, 72, 212, 1985. 27. Blevins, R. A. and Rulinsky, A., Comparison of the independent solvent structures of dimeric a-chymotrypsin with themselves and with •y-chymotrypsin, J. Biol. Chem., 260, 8865, 1985. 28. Schulz, G. E., Schirmer, R. H., Sachsenheimer, W., and Pai, E. F., The structure of the flavoenzyme glutathione reductase, Nature (London), 273, 120, 1978. 29. Chothia, C. and Janin, J., Principles of protein-protein recognition, Nature (London), 256, 705, 1975. 30. Greer, S. and Per ham, R. N., Glutathione reductase from Eschericia coli: cloning and sequence analysis of the gene and relationship to other flavoprotein disulfide oxidoreductases, Biochemistry, 25, 2736, 1986. 31. Stephens, P. E., Lewis, H. M., Darlison, M. G., and Guest, J. R., Nucleotide sequence of the lipoamide dehydrogenase gene of Escherichia coli K12, Eur. J. Biochem., 135, 519, 1983. 32. Rice, D. W., Schulz, G. E., and Guest, J. R M Structural relationship between glutathione reductase and lipoamide dehydrogenase, /. MoL Biol., 174, 483, 1984. 33. Schulz, G. E., Schirmer, R. H., and Pai, E. F., The FAD-binding site of glutathione reductase, J. Mol. Biol.t 160, 287, 1982. 34. Abraham, R. J., Hudson, B, D., Thomas, W. A., and Krohn, A., Electrostatic potentials of the alpha helix dipole and of elastase, J. Mol. Graphics, 4, 28, 1986. 35. Wierenga, R, K., De Maeyer, M. C. H., and Hoi, W. G. J., Interactin of pyrophosphate moieties with a-helixes in dinucleotide binding, Biochemistry, 24, 1346, 1985.

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Chemistry and Biochemistry of Flavoenzymes

36. Schmidt, J., Coudron, P., Thompson, A. W., Walters, K. L., and McFarland, J. T., Hydrogen bonding between flavin and protein: a resonance Raman study, Biochemistry, 22, 76, 1983. 37. Schirmer, R. H., Schulz, G. E., and Untucht-Grau, R., Hypothesis: on the geometry of leukocyte NADPH-oxidase, a membrane flavo-enzyme. Inferences from the structure of glutathione reductase, FEES Lett., 154, 1, 1983. 38. Krauth-Siegel, R. L., Schirmer, R. H., and Ghisla, S., FAD analogues as prosthetic groups of human glutathione reductase. Properties of the modified enzyme species and comparisons with the active site structure, Eur, J. Biochem., 148, 335, 1985. 39. Thornton, J. M., Disulphide bridges in globular proteins, J. Mol. Biol, 151, 261, 1981. 40. Richardson, J. S., The anatomy and taxonomy of protein structure, Adv. Protein Chem., 34, 167, 1981. 41. Huber, R. and Bode, W., Structural basis of the activation and action of trypsin, Ace. Chem. Res., 11, 114, 1978.

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Chapter 7

STRUCTURE AND FUNCTION OF SUCCINATE DEHYDROGENASE AND FUMARATE REDUCTASE Brian A. C. Ackrell, Michael K. Johnson, Robert P. Gunsalus and Gary Cecchini

TABLE OF CONTENTS I.

Introduction

230

II.

Types of Succinate Dehydrogenase (SDH) and Fumarate Reductase Preparations (FRD) A. Membrane-Bound Forms 1. Preparation 2. Topography of Complexes in Native Membranes B. Purified Complex 1. Isolation 2. General Properties C. Soluble Preparation Lacking Anchor Polypeptides

230 230 230 231 231 231 232 238

III.

Catalytic Activities A. Assay Methods B. Different Activities of Soluble and Particulate Enzyme Forms

239 239 241

IV.

Properties of the Catalytic Domain of Succinate Dehydrogenase and Fumarate Reductase A. Flavin Moiety B. Substrate-Binding Site C. Iron-Sulfur (Fe-S) Clusters

243 243 246 249

Properties of the Anchor Peptides A. Isolation B. Physical Properties C. Heme Component

266 266 266 266

V.

".

VI.

Assembly of Complex A. Processing of Subunits B. Incorporation of Prosthetic Groups C. Assembly of Complex

270 270 271 272

VII.

Interaction with Quinones

273

VIII.

Inhibition by Thenoyltrifluoroacetone and Carboxanilides

278

IX.

Molecular Biology and Genetics of SDH and FRD A. Gene Organization and Structure B. Regulation of Gene Expression

279 279 281

Acknowledgments

284

References

284

230

Chemistry and Biochemistry of ¥ lav o enzymes

I. INTRODUCTION Succinate dehydrogenase and fumarate reductase are complex flavoproteins with strikingly similar catalytic and physical properties. Succinate dehydrogenase is present in aerobic organisms as a membrane-bound component of the respiratory chain. A tricarboxylic acid cycle enzyme, it catalyzes the oxidation of succinate to fumarate and transfers the electrons directly to the quinone pool for common transport to oxygen and energy transduction. Fumarate reductase occurs in anaerobic and some facultative organisms, and may be either membrane-bound or free in the cytoplasm. 1 The membrane-bound enzyme catalyzes the terminal step in anaerobic electron transport when fumarate is utilized as the respiratory chain oxidant. During anaerobic growth the synthesis of succinate dehydrogenase in faculative anaerobes is repressed and fumarate reductase is induced in the presence of fumarate.23 Oxidative phosphorylation is then coupled to a foreshortened respiratory chain where the membrane-bound dehydrogenases are directly linked with fumarate reductase. The enzyme readily oxidizes succinate and can replace this function in sdh mutants of Escherichia coli if amplified by plasmid vectors.4 It has been proposed, in fact, on the basis of nucleotide sequence comparison that the genes encoding the catalytic subunits of E. coli fumarate reductase and succinate dehydrogenase are of common ancestral origin.5 Both enzymes are present in the membrane as multisubunit complexes. A hydrophilic catalytic domain comprised of two polypeptides contains an 8a-Af(3)-histidyl-FAD moiety and three iron-sulfur clusters: Centers 1 [2Fe-2S], 2 [4Fe-4S], and 3 [3Fe-4S]. The question of number and types of cluster present has only recently been resolved after some 20 years of extended effort by several laboratories and much controversy.6 A fifth prosthetic group, cytochrome b, is usually associated with the one or two hydrophobic subunits that anchor the catalytic subunits to the membrane and confer reactivity with quinones.7"14 Whereas beef heart succinate-ubiquinone oxidoreductase, or Complex II of the mitochondrial respiratory chain, has been the most intensively studied, attention of late is turning more to bacterial complexes that can be genetically modified and so used to answer questions of structurefunction not easily approached in the mammalian system. The result has been exciting and unprecedented advances in our understanding of the organization and operation of these enzymes, with several widely held beliefs called into question. Importantly, the similarities between succinate dehydrogenase and fumarate reductase have been confirmed at the molecular level. It is both expedient and beneficial, therefore, to summarize the properties of these enzymes collectively rather than separately.

II. TYPES OF SUCCINATE DEHYDROGENASE AND FUMARATE REDUCTASE PREPARATIONS Numerous procedures adopted over the years for isolating both particulate and soluble preparations of cardiac succinate dehydrogenase have been critically reviewed elsewhere.15'16 They need, therefore, only brief comment here for updating purposes, and to emphasize the general effectiveness of similar strategies for isolating succinate dehydrogenase or fumarate reductase from other sources. For the purposes of this discussion, preparations are categorized into three major types according to their structural sophistication. A. MEMBRANE-BOUND FORMS 1. Preparation Preparations of eukaryotic Complex II in the native environment of the mitochondrial inner membrane are usually obtained by differential centrifugation following disruption of isolated mitochondria by sonication or alkaline swelling. The membrane vesicles (ETP) are isolated in the everted configuration with the electron transport chain otherwise intact,

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including the major complexes I (NADH-ubiquinone oxidoreductase), II, III (ubiquinolcytochrome c oxidoreductase), IV (reduced cytochrome c oxidase), and V (ATP synthase). The properties and interactions of each of these complexes have been recently summarized by Hatefi,17 along with the coupling between electron transport and vectorial translocation of protons across the membrane, the driving force for ATP synthesis. Another membrane preparation, Keilin-Hartree particles, which was once a commonly-used source of simpler succinate dehydrogenase preparations, is obtained by grinding heart tissue with sand. The preparation consisting of damaged mitochondria and everted vesicles of inner membrane18 is considered modified in that the succinate dehydrogenase has a lower activity than when present in ETP.15-16 As source material, it nevertheless yields intact preparations of isolated Complex II.14 Preparations of microbial membranes containing succinate dehydrogenase or fumarate reductase are obtained by disruption of cells by standard techniques such as sonication, pressure cell extrusion, and grinding with glass beads, or from protoplasts by hypotonic shock or sonication. Respiratory chains of bacteria show great diversity, often containing several branches and terminal oxidases for oxidizing a variety of cytochromes and/or quinones directly. The quinone may be the benzoquinone ubiquinone, as in most aerobic systems, or menaquinone, a naphthoquinone. Details of bacterial energy transduction may also be found in several excellent review articles.19"22 2. Topography of Complexes in Native Membranes That the catalytic subunits of eukaryotic succinate dehydrogenase are located at the matrix surface of the mitochondrial inner membrane was first demonstrated by Klingenberg,23 who showed that ferricyanide, a membrane impermeant, was reduced by succinate in a reaction that was antimycin-insensitive in ETP but not intact mitochondria. This orientation of the enzyme is also evident in the inhibition of succinoxidase activity in ETP by antibody directed to the holoenzyme and by the lipid-insoluble protein-modifying reagent diazobenzene sulfonate (DABS).24 Use of [35S]DABS established the transmembrane arrangement of cardiac Complex II, by labeling selectively the flavoprotein (Fp) subunit in ETP and one of the two enzyme anchor peptides, CII-3, when intact mitochondria were used. Little isotope was incorporated in either experiment by either the second anchor peptide CII-4 or the ironsulfur protein (Ip) subunit, which are thus masked. Photolabeling of Complex II incorporated in egg lecithin vesicles with a series of aryloazidophospholipid derivatives confirmed both CII-3 and CII-4, as well as the Ip subunit of the enzyme, to be intercalated into the interior of the lipid bilayer.25 A diagramatic representation of Complex II structure in the mitochondrial inner membrane is given in Figure 1, which also summarizes our current knowledge regarding the subunit location of the three Fe-S clusters in these enzymes. Centers 1 [2Fe2S] and 3 [3Fe-4S] have been shown by genetic manipulation of E. coli fumarate reductase and B. subtilis succinate dehydrogenase to be present in the Ip subunit of the enzyme.26-27 Center 2 [4Fe-4S] may require cysteine ligands from the Fp subunit and thus might bridge both subunits (see Section IV.C. for discussion). Probing with antibodies, limited proteolysis, radiolabeling, and electron micrographs of progressively dismantled complex suggest a similar trans-membrane orientation for bacterial succinate dehydrogenase and fumarate reductase complexes in cytoplasmic and chromatophore membranes.28"32 Hydropathy plots used to predict the hydrophilic and hydrophobic regions of proteins provide further evidence for anchor polypeptides having several membrane-spanning a-helices.5'10-33 ~35 B. PURIFIED COMPLEX 1. Isolation Since the anchor peptides associated with fumarate reductase and succinate dehydrogenase

232

Chemistry and Biochemistry of Flavoenzymes

FIGURE 1. Topographical model of native succinate dehydrogenase and fumarate reductase.

are membrane intrinsic proteins (see Figure 1), liberation of the respective complexes requires disruption of the membrane. Nonionic detergents (Triton X-100, Emasol, Lubrol, dodecyl maltoside, or p-octylglucopyranoside) are now extensively used for this purpose, since they are less debilitory to protein structure than ionic types.36 Separation of the complexes from other respiratory components in the mixture can then be achieved by selective precipitation, assorted chromatographies, and centrifugation techniques, usually in combination. Inevitably over the years, an exceedingly large number of procedural variations and adaptations of these basic steps have been introduced, details of which are to be found in the original references quoted in Table 1. Obtained either as translucent suspensions of mixed micelles or in particulate form if the detergent is removed, the preparations contain substantial quantities of membrane phospholipids.7'37 2. General Properties The isolation of Complex II from beef heart mitochondria by Ziegler and Doeg37 in 1962 was arguably the major advance in the field, since it directly led to the first preparation of pure soluble succinate dehydrogenase that could be meaningfully characterized with respect to subunit composition and prosthetic groups. The complex itself consists of four major polypeptides apparently in unit stoichiometry: the Fp (70 kDa) and Ip (27 kDa) subunits of the enzyme38 and the two hydrophobic polypeptides of 15 and 13.5 kDa, respectively. The latter were initially referred to as CII-3 and CII-439 to depict their origin, and then more fashionably as QPs, when a possible role in ubiquinone reduction was realized.9 Direct evidence that these polypeptides can bind ubiquinone in the absence of the catalytic subunits is, however, scant and inconclusive. Speculation that mammalian Complex II might contain an additional small polypeptide or isoform has arisen because CII-4 often appears on SDSpolyacrylamide gels as a doublet.25'40 The complex, as isolated, contains by analysis ~8 gatom each of none-heme iron and labile sulfide per mol of histidyl flavin37 that constitute the [2Fe-2S], [4Fe-4S], and [3Fe-4S] clusters.41 The preparation also contains stoichiometric amounts of a £-type cytochrome.37 Only 60 to 70% pure by analysis (5 to 6 rather than 8.3 nmol FAD/mg of protein), the preparation is a heterogenous population of particles, as demonstrated by two-dimensional

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gel electrophoresis.24 The predominant contaminating polypeptides are from Complex III,39 with which there appears to be a close structural relationship. Preparations containing Complex III with Complex II in 1:1 stoichiometry or with the anchor polypeptides but not catalytic subunits of succinate dehydrogenase (the bcl complex) are easily obtained. Coprecipitation of both complexes by antibodies directed to peptides of either one, as well as evidence of structural changes when the complexes are combined, lend credence to the presence of a supermacromolecular entity in the membrane.24-42 In this "solid state" model electrons derived from succinate would be passed from Complex II to III via bound quinone without having to enter the ubiquinone pool. The model has difficulty explaining, however, why electron transfer is promoted by additional quinone in the membrane, and inhibited by dilution of the complexes with extra membrane phospholipid.43 A previously unidentified impurity is ETF (electron-transferring flavoprotein)~dehydrogenase (see Thorpe, Chapter 18, Vol. II of this series), which we have found by Western blotting to underlie the 70 kDa band of succinate dehydrogenase on gels (unpublished data). More recent procedures for isolating cardiac Complex II use Keilin-Hartree particles14 and succinate-cytochrome c oxidoreductase (Complexes II to III)40 as source materials and Triton X-100 as membrane-disrupting agent rather than the bile salts used in the original method. Although less harsh and protracted, these newer procedures do not significantly increase the purity or activity of the preparation, and drastically decrease the level of endogenous protoheme. This and the fact that the heme in cardiac Complex II is not extensively reduced by succinate37 have led to considerable skepticism regarding its origin and function.14-40 However, with the possible exceptions of yeast Complex II44 and E. coli fumarate reductase, complexes from other sources are isolated with stoichiometric amounts of cytochrome b present, and in several of these the heme is of sufficiently positive potential to be reduced by succinate at catalytically viable rates (Table 1). Even in mammalian Complex II, the dithionite-reduced cytochrome can be reoxidized by added quinone or fumarate,8 an indication of direct interconnection with other redox centers of the complex. A need for protoheme has been demonstrated in the assembly of the succinate dehydrogenase complex in B. subtilis membranes.45 As is evident from the examples listed in Table 1, the majority of succinate dehydrogenase and fumarate reductase complexes isolated from other phylogenetic sources closely resemble the cardiac one. They contain the analogous four polypeptides, identical 8a-7V(3) histidyl flavin moiety and Fe-S complement, and the same type of cytochrome b. For certain of the complexes, where analyses for non-heme iron and labile sulfide have not been done, e.g., those of M. luteus46 plant mitochondria,47 Ascaris suum48 and B. subtilis,49 the presence of Centers 1 [2Fe-2S] and 3 [3Fe-4S] have none-the-less been demonstrated by EPR spectroscopy. Variations noted in peptide molecular weights on gels can in part be attributed to the different electrophoretic procedures utilized. A sufficient number of amino acid sequences are by now available to establish that the structures of the Fp and Ip subunits are extensively conserved. For example, the primary sequences of the Fp subunits of B. subtilis50 and E. coli5 succinate dehydrogenases are —40% homologous with those of the fumarate reductases of E. coli51 and Proteus vulgaris,35 which are themselves 85% conserved. This is especially evident in critical stretches of the sequences constituting the binding fold and attachment site for the covalently-bound FAD (see Section IV. A), and those residues forming the active sites of these enzymes (see Section IVB). In addition, a 594-bp Sacll restriction fragment encompassing the 3' end offrdA and 5' end offrdB of the E. coli operon has been shown to strongly hybridize with DNA from other gram-negative bacteria. The list of organisms tested included, Shigella sonnei, S. flexneri, Salmonella typhimurium, Citrobacter freundii, Enterobacter cloacae, Klebsiella pneumoniae, and Serratia marcescens.52 In this same work, part of the fumarate reductase operon of S. sonnei was cloned into plasmid vectors and shown by in vitro protein synthesis to

Rhodopseudomonas sphaeroides Rhodospirillum rubrum Mycobacterium phlei

Desulfobulbus elongatus

Micrococcus luteus

Paracoccus dentrificans

Bacillus subtilis

Procaryotic Complex II Escherichia coli

Paragonimus westermani

Ascaris suum

Plant cotyledon (soybean)

Neurospora crassa

Bovine heart Bovine heart

Eucaryotic Complex II Bovine heart

Species

Fe/S

68; 30* 60; 25b 62; 26b

68.5; 27.5* 22

14.2; 12.8 65.2; 28. 3k 22.8 65; 29* 13.4; 12.5 72; 30* 17; 15

64.3; 26.6k

67; 30s 15; 13 68; 26* 15; 13.5 69; 27* 14.5; 12

14

— 57.3

— — —

5.2 — 8.0 —

_

—

—

5.6

10

9.85

62—67

— — —

— — —

—

— —

—

560"

556C

_ present* 6.9*

557C

558f

— 4,000 —

2,135

11. l u — 30.01-2 19.0U

—

18,300

5,600

2,112

—

lOO.O1'1"

20.8d-f

785

2.4 d - f 556C 556C

219 2.0"-'

558C

—

—

+ 36

—

-34

—

— —

—

557C

3,020 9,000

11,000

TNa

562

17.5d-f 54.0d-e

50— 55d-e

Specific activity

560 562h

557. 5C

3.7

13—20*

8.9*

2.0*

—

3.07

5.3*

—

9.3

—

—

5.0 present

__ —

-185

1.2 1.2—2.5

4.8—5.0

_

35—40

6.0

Cytochrome b (nmol/mg) (EJmV] (a-band[X])

—

48 50—54

5.8 5.9—6.2

unspecified 76; 26* 15.8; 14.9 72; 26b

32—38

(ngatom/mg)

4.6—5.0

FAD (nmol/mg)

73; 24b 13,7; 12.5

Major polypeptides (kDa)

TABLE 1 Characteristics of Isolated Complex II and Fumarate Reductase

85 86 75

84

83

33, 50 59, 81, 159 82

5, 54, 65

64

63, 64

47

60

7—9, 37 39, 80 40 14

Ref.

0

n

m

1

k

j

1

h

g

f

e

d

c

b

a

45; 32« 30; 27

65.8; 27k 15.0; 13.1 65.9; 27. 2k 14.7; 13 72.8; 27k 29.7

Mol. substrate reduced per min. per mol. of enzyme. Weber-Osborn gel system. At 77 K. Succinate-quinone oxidoreductase. At 38°C. At 25°C. Laemmli gel system. At 20°C. Modified Laemmli system. Reducible or partly reducible by succinate. Predicted from sequence. Succinate-PMS oxidoreductase. At 30°C Unspecified temp. Reduced benzyl viologen-fumarate oxidoreductase.

Desulfovibrio multispirans

Wolinella succinogenes

Proteus vulgaris

Procaryotic Fumarate Reductase Escherichia coii

67.7

8.5 absent

10.7J (diheme)

—

— 40—57

absent

32—40

6.0

—

4—5

„

-200

-20

—

—

110.0°'e

90.0°'"

560C

—

—

52.0°-e

—

—

13,000

15,000

—

11,500

69

66—68, 103, 158

34,

35

10—12, 41 51, 55

236

Chemistry and Biochemistry of Flavoenzymes

produce an Ip subunit and two anchor polypeptides with similar Mr values to those of the E. coli complex. Also, the Fp subunit of P. vulgaris will associate with E. coli subunits to form a functional complex.53 Comparison of the sequences of the Ip subunits of succinate dehydrogenase and fumarate reductases from mammalian and bacterial sources shows that the cysteine residues acting as ligands for the iron-sulfur centers are conserved and arranged in three distinct clusters in the sequence.50'54"57 Based on this information, oligonucleotide probes corresponding to two widely separated regions of the sequence conserved in both cardiac and £. coli succinate dehydrogenases have been used as primers in the polymerase chain reaction (PCR), in order to amplify and compare the gene sequences in other eukaryotes.58 The reaction products, representing about two thirds of the nucleotides encoding the whole subunit, were greater than 90% homologous for human, rat, beef, and Drosophila melanogaster. In accord with taxonomic hierachies, the equivalent sequences of Arabidopsis theliana, Schizosaccharomyces pombe, and Saccharomyces cerevisiae were >60% homologous to the human sequence. As opposed to the conserved nature of the Fp and Ip subunits, anchor polypeptides in the various organisms show little sequence homology. Notable exceptions to this generalization are those associated with the fumarate reductases of the related organisms E. coli and P. vulgaris, which exhibit 60% homology.35 An interesting divergence is the substitution of a single, larger polypeptide for the usual two anchor peptides. In Complex II of the strict aerobe B. subtilis the two enzyme subunits are associated only with cytochrome b55$ having a predicted Mr of 22,800.33 Approximately half of the heme content of 1.3 to 2.0 mol/mol of FAD is found to be rapidly reduced by succinate. Whether the protein contains one or two hemes is presently not known.59 The importance of the heme for proper processing of this cytochrome in the membrane is implied by the lack of assembly of the complex and accumulation of enzyme subunits in the cytoplasm of a hemA mutant starved for 8-aminolevulinate.45 The eukaryotic Complex II of Neurospora crassa similarly has only one anchor peptide, cytochrome b559. That this complex could be isolated intact from mitochondria of N. crassa grown in the presence of chloramphenicol is direct evidence that the cytochrome is encoded by nuclear genes, and thus different from cytochromes b of Complex III, which are encoded by mtDNA and repressed under these conditions.60 Previous spectroscopic evidence had also indicated cytochrome b of cardiac Complex II to be a distinct type.61 Whereas in most aerobic organisms electrons are transferred from succinate dehydrogenase to ubiquinone (Em = +110mV) for entry into the respiratory chain there are interesting exceptions. For example, the acceptor for the M, luteus52 and B. subtilis enzymes is likely menaquinone (Em = — 74mV). As has been pointed out, it is presumably not coincidental that the redox potentials of the Centers 3 of these particular enzymes should be slightly more negative than those of other succinate dehydrogenases in order to effect electron transfer.46 A good example of adaptation to environmental pressure is the mitochondrial complex of the mature parasite A. suumf3 which functions as a terminal oxidase in the anaerobic environment of the intestine by reducing fumarate. The electron donor for this reaction is reduced rhodoquinone of the membrane, a ubiquinone analogue (2-amino-3-methoxy-6methyl-5-isoprenyl-l,4-benzoquinone) with an Em ( — 63mV) much lower than that of ubiquinone itself, and thus thermodynarmcally disposed to reduce fumarate (Em = + 30mV). The parasite complex functions much less efficiently (15-fold) as a succinate-ubiquinone oxidoreductase.64 Also, the acceptor for the succinate dehydrogenase complex of the extreme thermoacidophilic (pH 2.3; 80°C) archaebacterium Sulfolobus acidocaldarius has been shown to be caldariellaquinone (a 5-methylthiobenzo[b]thiophen-4,7-quinone), which is unique to sulfur-metabolizing thermophiles (R. Moll, personal communication643). Menaquinol serves as the electron donor for the bacterial fumarate reductase complexes.

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As evident from Table 1, the physical properties of the E. coli enzyme closely ressemble those of cardiac Complex II except for an apparent absence of cytochrome b. The complex can reduce ubiquinone with succinate as substrate under appropriate conditions, at rates only two- to fourfold slower than fumarate is reduced.12 The operon has been cloned,12-51-55 thereby allowing individual and combinations of the structural genes (frdA,frdB,frdC, andfrdD) to be put in multicopy vectors for studies of complex assembly and function. The complex can be amplified up to 20-fold in transformed cells,11'12 much higher than has thus far been attained with other cloned systems such as B. subtilis33*50 and E. coli65 succinate dehydrogenases. These attributes make it a very attractive bacterial model for the mammalian system. Also of interest is the fumarate reductase complex of W. succinogenes, now cloned in its entirety66 (A. Kroger, personal communication663), since it contains only one anchor polypeptide, cytochrome b560,67 and thus is structurally analogous to the succinate dehydrogenase complexes of N. crassa and B. subtilis (Table 1). Earlier assertions68 that this enzyme contained only enough labile sulfide to accommodate two Fe-S clusters have been revised.67 A very unusual type of fumarate reductase has recently been isolated from the anaerobe Desulfovibrio multispirans69 that seemingly combines the properties of a membrane-bound complex (Table 1) with those of the soluble-type fumarate reductases found in baker's70-71 and brewer's72'73 yeasts grown on glucose (anaerobically). This bacterial complex typically is membrane-bound and comprised of four subunits with reported Mr values of 46, 32, 30, and 27 kDa, respectively. Unless the polypeptides have been misidentified on gels, these Mr values are in themselves atypical (Table 1). The preparation contains the usual 8 to 9 gatom each of Fe and labile sulfide per mol of enzyme, with [2Fe-2S] and [3Fe-4S] clusters being thus far identified by EPR (see Section IV. C.), but the FAD component is noncovalently bound, as it is in the soluble fumarate reductase of yeast. The latter enzyme is a single polypeptide of 58.8 kDa lacking non-heme iron,70-71 for which the physiological electron donor is probably NADH2. The reaction catalyzed by both the yeast and D. multispirans enzymes is unidirectional, with fumarate being reduced, but succinate not oxidized. Significantly, this nonreversible type of reaction mechanism is also catalyzed by mutant forms of E. coli fumarate reductase lacking the histidine residue (His-44) required for covalent linkage to the FAD, which can then bind only noncovalently.74 The data collectively suggest that one purpose of the 8a-M3)-histidyl-FAD linkage in evolution is to change the transition state of the reaction to allow succinate oxidation, possibly by raising and, hence, 'finetuning' the redox potential of the flavin (see Section IV.A for discussion). The succinate dehydrogenase of Mycobacterium phlei would be an exception to this rule by apparently not containing covalently bound FAD,75 in itself an indication of the major effect of noncovalent binding in raising the redox potential of the cofactor for catalysis. It has been suggested that the earliest fumarate-reducing systems were soluble and functioned primarily to retain redox balance in the cell by recycling NAD + for simple fermentations.76 Soluble enzymes have been reported in yeast, as discussed above, Veillonella alcalescens,71 and Erwinia herbicola.™ As noted by Gest,76 it would have become energetically advantageous in the course of evolution to couple phosphorylation with electron transport to fumarate, by association of the enzyme with the membrane, as in most contemporary organisms. This would have required the addition of intermediate electron carriers of suitable redox potential, i.e., iron-sulfur clusters, menaquinone and cytochrome b (see Figure 2). In extending this thesis, we suggest that at this juncture the enzyme flavin was probably still noncovalently bound as in the D. multispirans complex and His-44 mutants of E. coli fumarate reductase. The former might thus be considered a missing link,79 since a major next step towards aerobic electron transport would have been covalent attachment of the flavin moiety to allow succinate oxidation (our results, see above and Section IV.A). This reaction is thermodynamically favored in most aerobic organisms by the synthesis of UQ in replace of MQ to raise the redox potential of the electron acceptor. An additional

238

Chemistry and Biochemistry of Flavoenzymes

FIGURE 2. Evolution of succinate dehydrogenase. (A) Primitive, soluble fumarate reductase with noncovalent FAD, incapable of succinate oxidation. (B) Enzyme associates with membrane and acquires Fe-S clusters and cytochrome b for interaction with menaquinone (MQ).76 (C) Ability of contemporary fumarate reductases to oxidize succinate is the result of covalent attachment of FAD.74 (D) Adaptation to succinoxidase activity fostered by the presence of ubiquinone (UQ) and more positive redox potentials of constituent Fe-S clusters (Table 4).

refinement, presumably related to changes in protein structure, is seen in the redox potentials of the non-heme iron clusters serving as electron mediators, which are more positive in succinate dehydrogenases than in fumarate reductases (see Section IV.C). C. SOLUBLE PREPARATION LACKING ANCHOR POLYPEPTIDES Most of the information available on the soluble form of these enzymes comes from studies of beef heart succinate dehydrogenase. Older procedures for obtaining this mitochondrial enzyme entailed alkaline extraction of acetone powders of membranes or of ETP or Keilin-Hartree particles with w-butanol present, as well as treatment of membranes with cyanide, are discussed in References 15,16. These types of preparations were impure and often deficient in Fe-S complement due to damage of the exposed Center 3 by oxygen. The cluster has to be protected by conducting extractions anaerobically with succinate present. Solubilized E. coli fumarate reductase12 has been shown to be similarly oxygen-sensitive and not fully protected by succinate because of incomplete reduction of Center 3. From a technical standpoint, it becomes impractical to attempt extensive purification once an enzyme has been solubilized. Soluble forms of the enzymes are therefore best obtained by resolution of the purified complexes to minimize the steps involved and chances of damage by oxygen. Cardiac succinate dehydrogenase in fully reconstitutively active, and hence unmodified form, has been obtained by treating Complex II with sodium perchlorate38 or, at higher pH, with

Volume HI

239

n-butanol. 87 Use of perchlorate has the advantage of not inactivating the anchor peptides CII-3 and CII-4, so that a preparation of the latter in purified form can be subsequently obtained from the enzyme-free residue using detergent.7-8 The strength of enzyme binding to the membrane varies among species, so that no one extraction method is universally proficient. Whereas perchlorate and freeze-thaw (plus high salt), respectively, will liberate soluble succinate dehydrogenase from chromatophores of Rhodospirillum rubrunf6 and Halobacterium halobium** the methods are ineffectual when applied to ETP and membranes of other bacterial species. The butanol method used for obtaining pristine succinate dehydrogenase from beef heart Complex II87 will yield inactive fumarate reductase from the E. coli complex, as will the alternative perchlorate method, unless exposure to the chaotrope is minimized and the extraction is conducted at neutral rather than alkaline pH. 12 With these modifications to the procedure, a preparation of soluble fumarate reductase has been obtained in which 50% of non-heme iron cluster Center 3 remains intact. As previously mentioned, part of the inactivation may relate to the much lower redox potential of the cluster (Em) = - 70mV) in the bacterial as compared to beef enzyme (Em = + 60 mV),89 such that the center cannot be fully reduced and is thus protected from traces of oxygen by succinate.12 In an alternative method, E. coli complex has been bound to a Sepharose CL-6B column and washed with ammonium acetate to dissociate and elute the soluble form of the enzyme.90 Assessment of the quality of this preparation is difficult since no precaution was taken to protect the liberated enzyme from oxygen, nor has the residual reconstitutive capacity or final stoichiometry of iron-sulfur clusters been determined.

III. CATALYTIC ACTIVITIES A. ASSAY METHODS Succinate oxidation by both succinate dehydrogenase and fumarate reductase is most conveniently assayed spectrophotometrically using 5-7V-methyl phenazonium sulfate (PMS) as primary electron acceptor. This is a multipurpose acceptor in that it interacts with both soluble and paniculate forms of the enzymes. Reduction of ubiquinone, or one of its synthetic analogues.91'92 requires the presence of the "Q-binding" anchor peptides7"9'12-13 and is thus observed only with paniculate preparations containing unmodified complex. In both assays, 2,6-dichlorophenol indophenol serves as the terminal electron acceptor for the reaction. Detailed protocols for the methods are given elsewhere.16 Other efficient electron acceptors used with the cardiac enzyme are Wursters blue (TMPD*; a semiquindimine radical of AWA^'-tetramethyl phenylenediamine)93 and ferricyanide. Cardiac succinate dehydrogenase has been shown to exhibit two types of ferricyanide reductase activities attributable to a high KM and low KM site, respectively. The former activity is less (—50%) than that observed with PMS and is catalyzed by soluble and paniculate preparations alike.94 The ability to rapidly reduce low concentrations of ferricyanide (KM 250 (xAf) at rates similar to that of PMS is an exclusive property of reconstitutively active, and thus intact, soluble preparations.95 The activity is a result of interaction with iron-sulfur cluster Center 3 [3Fe-4S], which becomes exposed to solvent when the enzyme is solubilized and thus accessible to ferricyanide. Ordinarily masked by the binding peptides in particulate preparations, the cluster is considered the physiological electron donor to quinone. Exposure of soluble preparations of the enzyme to oxygen in the absence of succinate results in the rapid disruption of the cluster (t1/2 — 50 min. at 20°C) and loss of its EPR signal.96"98 The fact that this occurs concomitantly with loss of the low KM ferricyanide reductase activity indicates the close interrelationship of these three properties (Figure 3). It is important to note that the modified enzyme lacking Center 3 still retains succinate-PMS reductase activity, but at a decreased level and with changes in the KM app. for this acceptor.

240

Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. Decay of reconstitutively active (two subunit) beef heart succinate dehydrogenase on contact with air (0°C). (O) reconstitution activity; (D) 'Mow K m " ferricyanide activity; (A) EPR signal of oxidized Center 3 [3Fe-4S]. (From Beinart, H. et al., Arc/i. Biochem. Biophys., 182, 95, 1977. With permission.)

An earlier report that reconstitutively active cardiac succinate dehydrogenase will also reduce horse heart cytochrome c95 has recently been concluded not to reflect direct interaction, but rather mediation by an as yet unidentified radical species." This does not reconcile, however, with our own unpublished observations that saturation, albeit complex, enzyme kinetics are obtained under aerobic or anaerobic conditions provided the reaction is conducted in low ionic strength buffers, i.e., there is a strong electrostatic component to the interaction. Further, the presence of reduced cytochrome or substitution of an aryloazido group at lysine13 adjacent to the heme cleft of the oxidized cytochrome both inhibit activity. That the enzyme's [3Fe-4S] cluster is the probable electron donor in the reaction is suggested by the concomitant loss of cytochrome c and low KM ferricyanide reductase activities on exposure of the enzyme to air. Unfortunately, there are many conflicting data in the literature particularly regarding the catalytic efficiencies and stabilities of preparations of bacterial succinate dehydrogenases and fumarate reductases. These discrepancies can generally be traced to inappropriate assay conditions, such as the use of inefficient electron acceptors, activity measurements made at a single acceptor concentration rather than at Vmax? and the use of less-than-fully active enzyme preparations. Paniculate preparations of succinate dehydrogenase and now fumarate reductase12 are recognized to be isolated in a partially deactivated state due to the presence of enzyme molecules containing stoichiometric amounts of inhibitory oxaloacetate in tightly bound form (see Section IV.B). Full activity can be restored by incubation of such samples with succinate or malonate at elevated temperatures, by reduction, or by treatment with anions at acid pH if the oxidized form of the enzyme is required. The methods orginially developed for activating cardiac succinate dehydrogenase16 work equally well with bacterial succinate dehydrogenases and fumarate reductases. Assays of fumarate reduction are not influenced by tightly-bound oxaloacetate, since the binding is considerably weakened on reduction of the enzyme by the electron donor for the reaction. The affinity of oxaloacetate for the reduced forms of both cardiac succinate dehydrogenase and E. coli fumarate reductase is tenfold less than for their oxidized forms.100102 The reductant most often employed in the fumarate reductase assay is dithionitereduced benzyl viologen, which has a very low redox potential (Em = — 359mv) and will

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interact with both soluble and paniculate enzyme forms. The use of reduced quinones or their analogues as electron donors is restricted to assays of paniculate preparations because of the requirement for the anchor polypeptides. The quinone must also be of sufficiently negative redox potential to reduce Center 3 of the enzyme, which is considered the immediate acceptor of the electrons. Reported Em values for Center 3 range among species from -70 to + 60mV.46102 In their oxidized forms, both enzymes bind oxaloacetate extremely tightly at the active site in 1:1 stoichiometry and with loss of activity.102'149 The possibility that this inhibitor might bind at a regulatory site has been discounted. Measured koff rates for dissociation of oxaloacetate, which is faster from fumarate reductase (0.06 min" 1 ) than cardiac succinate dehydrogenase (0.02 min~ *), are approximately 5 to 6 orders less than the turnover rates of these enzymes (104 min" 1 at 25°C). The slow equilibrium of these complexes, minutes as compared to seconds with other ligands, precludes meaningful measurement of succinate oxidation rates or studies with other inhibitors unless the oxaloacetate is first removed.101 The unusually tight binding of oxaloacetate to cardiac succinate dehydrogenase has been attributed to interaction with an active site sulfhydryl and stabilization of the weak thiohemiacetal bond by the protein environment.150 Studies of the inhibition and reactivation processes indicate that tight binding develops subsequent to the initial loose interaction and is accompanied by conformation changes in the protein, as evidenced by induced spectral changes,148 loss of endogenous FAD fluorescence,100 and the high energy of activation (20 to 30 kcal mol ~ ' ) needed for reactivation.149 Both the enol and keto tautomers of oxaloacetate have been shown to be strong, slowly dissociating inhibitors of the enzyme, with the former inhibiting approximately three times faster than the keto form. 151 Significantly, during malate oxidation, suicide deactivation of the enzyme can be delayed if the initial enol product is rapidly converted to the keto form under conditions of high ionic strength and a trapping system (transaminase). In Scheme 1 the data are rationalized in terms of two kinetically

SCHEME 1.

248

Chemistry and Biochemistry of Flavoenzymes

FIGURE 6. Comparison of amino acid residues at the putative substrate binding site of fumarate reductase and succinate dehydrogenase. The partial sequences are for the bacterial enzymes cited in the legend to Figure 5. The human succinate dehydrogenase flavoprotein sequence (Human SdhFp) was kindly provided by M. Malcovati (personal communication) and deduced from a partial DNA sequence obtained from a human placental tissue Xgtll library. Identical residues are indicated by bold type.

distinct enzyme-oxaloacetate complexes where M is L- or D-malate, E is succinate dehydrogenase, A is an electron acceptor, Oe and Ok are the enol and keto forms of oxaloacetate, kt is the equilibrium constant for oxaloacetate tautomerism, and E.O* is the dead end, slowly dissociating complex.151 Sulfate has also been shown to induce a slow pHdependent (pKa —7.2) transformation of an initial enzyme-SO^ complex. Coincidence of the pKa values for formation of this complex and maximum activity of the enzyme suggests that ionization of the same group of the enzyme is involved in both processes.152 Reactivation of both succinate dehydrogenase and fumarate reductase at elevated temperatures is facilitated by the presence of inorganic anions (NO 3 ~, ClO^, I~ > Br~ > Cl~) at acid pH,153 substrates, or competitive inhibitors, which prevent rebinding of dissociating oxaloacetate. Use of oxaloacetate-trapping agents such as semicarbazide or glutamate-oxaloacetate transaminase does not affect activation rates. On the other hand, metabolic reduction of the membrane quinone pool will accelerate activation of the deactivated membranebound forms of the enzymes (''rear-end" activation).101-154 Reductive-activation has been shown to involve reduction specifically of the FAD prosthetic group of the enzyme and thus presumably reflects localized conformation changes at the active site that result in loosening of oxaloacetate binding.100'102 The composition of the active site and identities of functional residues are not yet unambiguously defined. An unusually reactive thiol has been located by chemical modification studies on the flavoprotein subunits of these enzymes.29'102-155157 Protection by substrate and competitive inhibitors suggest the cysteine to be present at or close to the active site, where it has been proposed to contribute to the catalytic mechanism157 and be responsible for the tight binding of oxaloacetate, as discussed above. The residue was subsequently mapped in E. coli fumarate reductase to the middle of the subunit,156 which is now known to be the location of the only invariant cysteine in the flavoprotein sequences of several enzymes, including E. coli succinate dehydrogenase and fumarate reductase,5'51 and the fumarate reductases of P. vulgaris35 and W. succinogenes.*58 Some 200 residues downstream of the histidine forming covalent linkage to FAD, the thiol is in a highly conserved part of the sequence (Figure 6) midway between those regions thought to form the FAD binding fold. Moreover, the His-Pro-Thr triad of residues present at the start of the sequences shown in Figure 6 is identical to the active-site histidine sequences of the flavoenzymes glutathione and lipoamide reductases.5 The succinate dehydrogenase of B, subtilis does not have the cysteine residue conserved in the other bacterial enzymes,50 nor is it sensitive to thiol reagents. Substitution of cysteine for Ala-251 by site-directed mutagenesis, which is without major effect on catalytic activity, produces the sensitivity to W-ethyl maleimide (NEM) observed with the other enzymes, and the inhibition can be prevented by the presence of substrate or competitive inhibitors.159 Conversely, in our own preliminary studies, substitution of the cysteine (Cys-247) of E. coli

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fumarate reductase resulted in loss of NEM sensitivity. An important observation is that both this E. coll variant and the wild-type B. subtilis enzyme159 bind oxaloacetate extremely tightly. Hence, an active site thiol would appear not to be "essential" either for catalytic functioning of succinate dehydrogenases and fumarate reductases or the binding of oxaloacetate through thiohemiacetal linkage, as previously proposed. Sulfhydryl reagents presumably act by impeding access of substrate to the active site. Chemical modification by other "residue-specific" reagents has indicated the presence also in cardiac succinate dehydrogenase of essential arginine and histidine residues. It has been suggested that an arginine participates in substrate binding by forming an ion pair with a carboxylic group and confers high reactivity on the neighboring active site cysteine.160 A histidine has been speculated either to help bind substrate or perform proton donor/acceptor function in the catalytic mechanism.161 That these residues are probably those located in the short sequences given in Figure 6 is evidenced by the fact that substitution of His-232 or Arg-248 in E. coli fumarate reductase has a profound effect on catalytic proficiency (unpublished data). Absence of Arg-248 decreased both succinate oxidation and fumarate reduction to less than 1% of wild-type levels. On the other hand, in the absence of His-232, succinate oxidation was essentially lost but fumarate reductase activity retained. Chemical modification of the enzyme using the histidine reagent Rose Bengal had precisely the same effects as mutating His-232 to serine. Thus, whereas this particular histidine could indeed be involved in proton abstraction and donation by the wild-type enzyme, as proposed, the possibility arises that these processes do or can occur by different routes. Unfortunately, no complete eukaryotic Fp sequence has been reported. A major fragment of the gene for the human placental succinate dehydrogenase subunit has recently been cloned whose nucleotide sequence covers some 67% of the B. subtilis sdhA gene without gap insertion and with striking homology (87%) at the amino acid level.563 A part of the fragment encodes the active site sequence shown in Figure 6, which is seen to lack the cysteine also missing from B. subtilis. By analogy, the placental enzyme would be expected to be insensitive to sulfhydryl reagents and thus different to the beef heart enzyme. A provocative point at present, therefore, is whether the isoforms are species or tissue specific. Comparison of the full and partial sequences available for human liver57 and human f ibroblast Ip subunits,58 respectively, provides evidence of a minimum of two amino acid residue changes and, hence, additional support for tissue specific isozymes of Complex II in human mitochondria. C. IRON-SULFUR (Fe-S) CLUSTERS Over the past 5 years evidence has emerged for three distinct types of Fe-S cluster in the most intact preparations of beef heart succinate dehydrogenase and E. coli fumarate reductase: Center 1, a [2Fe-2S]2+-1 + cluster; Center 2, a [4Fe-4S]2 + - 1 + cluster; Center 3, a [3Fe-4S]1 + >0 cluster. The results have been summarized in recent review articles.6'41-162 Although EPR spectroscopy has assumed the leading role in characterizing the properties of the Fe-S clusters in these enzymes, it is noteworthy that the current picture only became apparent as a result of biophysical studies using additional techniques, notably low temperature magnetic circular dichroism (MCD) spectroscopy.163'164 Bearing in mind the ease of oxidative degradation of Center 3 in purified soluble preparations (see Figure 3), a cluster composition consisting of one of each type of center is consistent with the available spectroscopic and analytical information. Moreover, it provides a rational explanation of earlier Fe-S core extrusion and interprotein core transfer studies using the most intact preparations of beef heart succinate dehydrogenase. These studies were interpreted in terms of two [2Fe2S] and one [4Fe-4S] center,165 but [3Fe-4S] clusters have yet to be extruded intact from any protein and have been shown to extrude as [2Fe-2S] centers in the case of aconitase.166 Based on the available EPR data it seems probable that all fumarate reductases and

250

Chemistry and Biochemistry of Flavoenzymes

succinate dehydrogenases have a similar Fe-S cluster composition, irrespective of their plant, animal or bacterial origin. To recapitulate, the only apparent exceptions are the fumarate reductases from D. multispirans and baker's yeast. The baker's yeast fumarate reductase consists of a single subunit and is devoid of non-heme Fe.70 Preliminary EPR studies of D. multispirans fumarate reductase were interpreted in terms of only [3Fe-4S] and [4Fe-4S] clusters.69 However, subsequent EPR and MCD investigations of anaerobically prepared samples found no evidence for a [3Fe-4S]1 + -° cluster (unpublished work, Kowal, A. T., He, D. V., DerVartanian, J., LeGall, J., and Johnson, M. K.)- Moreover they show that the almost axial EPR signal with gB = 2.03 and gi = 1.93 originates from a [2Fe-2S]1 + cluster (Center 1) rather than a [4Fe-4S]'+ cluster as originally proposed, and that a hithertounreported rhombic resonance, g = 2.05, 1.95, 1.90, with relaxation properties indicative of a [4Fe-4S]1 + is apparent in dithionite-reduced samples. These studies are still in progress and further discussion of Fe-S centers in this enzyme must await completion of these investigations. The spectroscopic and redox properties and the evidence for the assignment of clustertype are described separately below for each center. This is followed by discussions of their spatial proximity as assessed by intercluster magnetic interactions, and the ongoing attempts to identify the subunit location and the specific residues involved in cluster ligation. The midpoint reduction potentials for centers 1, 2, and 3 have been determined for a wide range of succinate dehydrogenases and fumarate reductases by EPR-monitored mediator titrations, and the values (pH 7.0) are summarized in Table 4. The potentials are not significantly pHdependent, which is consistent with a pure electron transport role for these clusters. In common with all known [2Fe-2S] centers, the oxidized form of Center 1 is diamagnetic at low temperatures as a result of strong antiferromagnetic coupling between two high spin Fe(III) sites (S = 5/2) giving rise to a S = 0 ground state, and the reduced form exhibits an EPR-active S = 1/2 ground state as a result of antiferromagnetic coupling between high spin Fe(III) (S = 5/2) and Fe(II) (S = 2) sites. The g-values and relaxation properties of reduced Center 1 are typical of those observed for both synthetic and biological [2Fe-2S]1 + centers.174 The EPR relaxation properties of [2Fe-2S]1+ centers are known to vary as a result of differences in the magnitude of the antiferromagnetic coupling constant (e.g., J = 65 cm" 1 for beef heart Rieske protein, J = 90 cm" 1 for beef heart succinate dehydrogenase, J = 270 cm~' for adrenodoxin).175 However, in contrast to [4Fe-4S]l + clusters, EPR signals from S = 1/2 [2Fe-2S]1+ clusters are generally observable at 77 K. Additional evidence for the assignment of Center 1 as a [2Fe-2S]2 + ' 1+ center has come from CD,163 variabletemperature MCD,163 and linear electric field effect EPR studies176 of beef heart succinate dehydrogenase. The EPR characteristics of reduced Center 1 are remarkably similar in all succinate dehydrogenases and fumarate reductases (see Table 5). The spectra comprise an anisotropic resonance, gz = 2.030 ± 0.005, gy = 1.935 ± 0.005, and gz = 1.905 ± 0.015 that accounts for ~1 spin /FAD in dithionite reduced samples,162"164 and can be observed without significant broadening at temperatures up to 80 K. For example, Figure 7 shows EPR spectra for the Center 1 in the succinate- and dithionite-reduced E. coli fumarate reductase complex. Dithionite reduction results in loss of the FAD semiquinone radical signal centered at g = 2.004, minor changes in the g-value anisotropy, and a dramatic enhancement of the spin relaxation properties.191 In common with other fumarate reductases, but in contrast to succinate dehydrogenases, succinate does not effect complete reduction of Center 1 in E. coli fumarate reductase. This is illustrated by the relative intensity of the EPR signals shown in Figure 7, which quantify to 0.6 ± 0.1 and 1.05 ± 0.15 spins/molecule for the succinatereduced and dithionite-reduced samples, respectively. In accord with this, EPR redox titrations indicate a midpoint potential for Center 1 in the range - 10 mV to + 80 mV in succinate

b

a

MQ MQ

UQ UQ UQ UQ MQ MQ -59

-20 to -79

+ 70 + 80

_7

+ 10 + 50

0

Center 1

-320 which becomes fully reduced, for entry into the Q pool (Scheme 3). The different

SCHEME 3.

electron-transferring and thermodynamic characteristics of the QA and QB quinones reflect structural differences at the respective binding sites.286 In the linear-sequence model, Center 3 of succinate dehydrogenase or fumarate reductase would serve as the donor/acceptor of successive electrons in interactions with the immediate quinone, QA. As in photosynthetic reaction centers, the partner quinone, QB, is considered able to exchange with other quinones, whereas QA is nonexchangeable and functions as an obligatory n = 1 system, unless the complex be structurally modified (see discussion above). Mutations in the anchor polypeptides of the E. coli fumarate reductase complex that have already been reported to prevent menaquinol oxidation and, hence, growth, include an Arg substitution for His-82 in FrdC,114 truncations in FrdC or FrdD,271*272 and an as yet unidentified lesion in FrdD.115 We have isolated an additional eight mutants, seven of which reflect frameshifts and truncation of FrdC (5 mutants) or FrdD (2 mutants) and one where a G to A transition produced a Glu to Lys exchange at residue 29 of FrdC.293 The truncations are severe, representing deletions of the second and/or third transmembrane loops of the respective polypeptides (Figure 17). Glutamate-29 is positioned at the N-terminus of the first loop of FrdC and thus predicted to be in juxtaposition to His-82 at the C-terminus of the second transmembrane loop, on the cytoplasmic side of the membrane, close to the bound catalytic subunits. Table 7 summarizes the effects of these mutations on the assembly of complex in the membrane and on the ability to oxidize and reduce quinones. In all mutants containing truncated FrdC or FrdD polypeptides, membrane-bound fumarate reductase complex was depleted similarly to 12 to 20% of the level observed in wild type cells. That truncations of FrdD after the first and second transmembrane a-helices, respectively, affected complex assembly identically confirms the observations of Latour and Weiner271 that the C-terminal portion of this polypeptide is important for structural integrity. Similar arguments with respect to the C-terminal portion of FrdC are less persuasive, since all of the frdC frameshift mutations resulted in truncations close to the termination of the first helix. Significantly, mutant complexes lacking the second and third transmembrane ahelixes of FrdC (pDW2537 and pDW2506) or the third helix of FrdD (pDW2510) lost most of their ability to oxidize menaquinol or reduced benzyl viologen (see Section III.A, for discussion of assays) yet retained 57 to 100% of that for reducing DPB and PMS. Such clear-cut separation of oxidative and reductive activities is evidence for two quinone binding sites in the native complex, consistent with the reaction mechanism formulated in Schemes 2 and 3 and the properties proposed for reaction centers QA and QB. Just as significantly, therefore, incorporation of a severely truncated FrdD polypeptide, such as that produced by plasmid pDW2501, affects steps apparently central to the overall mechanism by drastically curtailing catalytic competence in both the oxidative and reductive directions. Such a major

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FIGURE 17. Schematic drawings of wild type and mutant FrdC and FrdD polypeptides of E. coli fumarate reductase. The probable membrane-spanning helical domains are indicated by the hatched lines and the nonwild type sequence resulting from a frameshift mutation is shown by the solid line. Plasmid pDWIOO encodes wild type FrdC and FrdD peptides, pDW2106 encodes the FrdCE29K substitution, pDW126 encodes the FrdCH82R substitution, pDW2537 and pDW2506 encode FrdC frameshift deletions, and pDW2501 and pDW2510 encode FrdD frameshift deletions. The mutations are described in detail in Reference 293.

TABLE 7 Turnover Numbers of Mutant Complexes*

Wild Type 2537 FrdC 2506 FrdC FrdC Glu29Lys FrdC His82Arg FrdD 2501 FrdD 25 10 a

PMS

DPB

DPB/PMS

BVred

BV/PMS

MQH2

15,000 13,500 10,800 15,800 6,400 4,800 14,500

9,200 8,600 5,200 1,600 1,400 1,700 7,500

0.61 0.64 0.48 0.10 0.28 0.35 0.52

59,200 11,900 12,800 60,600 48,600 7,600 3,500

3.9 0.9 1.2 3.8 6.9 1.6 0.2

52,600 2,700 4,000 0 8,000 0 0

Activities (38°C) are expressed as JJL mol of succinate oxidized or fumarate reduced per minute per mol covalent FAD in the membrane.

effect would be expected, for example, if the QA site were disrupted or its electron-exchanging capabilities with either QB or Center 3 of the enzyme. The data outlined above make it unlikely that catalytic and structional roles can be apportioned expressly between the two subunits. Nevertheless, specific loss of menaquinol oxidation due to deletion of the major part of FrdC or the terminal region of FrdD is clearly compatible with an affect on the structural integrity of QB in Scheme 3 and the ability to bind and/or deprotonate reduced quinones. Retention of DPB reductase activity by these particular mutants is not unreasonable provided DPB is still bound by the defective complexes, since an increase in the effective pKa would promote protonation of the reduced form following electron transfer. Alternatively, if the QB site were completely disrupted,

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Chemistry and Biochemistry of Flavoenzymes

QA might function as an n = 2 system, precedence for which is the apparently modified reaction mechanism conducted by reconstituted Complex II preparations.285 The/rJCE29K mutant is unable to oxidize menaquinol or reduce DPB. Unlike the//Y/C frameshift mutants, this modified complex assembles normally in the membrane and has wild type activities in the reduced benzyl viologen and PMS assays. Interestingly, an essential, protonatable glutamate residue is present at the QB site of plant and bacterial photoreaction centers, where it is known from the X-ray crystal structure to be in contact with the ring of the bound quinone.286 It may not be coincidental, therefore, that the mutation of this conserved glutamate residue294 and that on FrdC both result in loss of reactivity with quinones. Also, mutation of the photoreaction center glutamate is considered to prevent proton donation to the reduced quinone but not to interfere with electron transfer between the QA and QB sites. By analogy, therefore, the/rdCE29K mutation in the fumarate reductase complex could also affect protonation rather than electron transport. This would be in keeping withthe observed oxidation of reduced benzyl viologen and reduction of PMS in this mutant, if these dyes have access to the specifically bound quinones. The similar activity profiles exhibited by the/r^CE29K and/rJCH82R mutants suggest these residues to be essential constituents of the same quinone binding site, possibly QB.

VIII. INHIBITION BY THENOYLTRIFLUOROACETONE AND CARBOXANILIDES Thenoyltrifluoroacetone (TTFA) and carboxanilides are potent, lipophilic inhibitors of the succinoxidase, succinate-cytochrome c and succinate-CoQ reductase activities of membrane fractions of a select number of organisms. These specific inhibitors do not influence, however, the activities of the solubilized form of the enzyme.295-296 Carboxin (5,6-dihydro2-methyl-l, 4-oxathiin-3-carboxanilide) and its analogues were developed initially as systemic fungicides to control Basidiomycetes, primarily rusts and smuts, and the activity extends to certain other species. The list includes, inter alia, Ustilago species,108'297-298 on which most research has been conducted, Rhizoctonia solani,215 min) (fully reduced); (H) spectrum F to compare g values with the "fully reduced" sample G. Modulation 1 mT; microwave power 2 mW; temperature 8 K.

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homology is observed when the physicochemical data available are compared within this group of APS reductases (Table 1). The active center composition is quite conserved: 8 ± 1 iron atoms per flavin and per minimal molecular mass. Other parameters vary within a narrow range: molecular mass (100 to 180 kDa); pH for optimal activity (7.4 to 7.6), and kinetic data (KM for sulfite, 0.34 to 0.76 mA/ and for AMP, 0.16 to 0.31 mM, using the ferrycianide assay). The UV/visible spectra are also quite similar with maximum absorption bands around 375 and 394 nm. Upon addition of sulfite, the absorption spectra of all the reductases decreases between 500 and 350 with a concomitant increase in the 320 nm region attributed the formation of the sulfite-flavin adduct. The EPR data obtained in different states of the enzyme (native, AMP plus sulfite reacted and dithionite reduced) also indicates the high degree of homology (Figure 10). The unusual high midpoint redox potential determined for Center I in the D. gigas enzyme (0 mV, respectively). Center II was determined to be quite negative (< -400 mV) in the case of the above enzymes. It is evident that, in Desulfovibrio, APS reductase is a conserved enzyme that represents an optimized solution for this essential enzymatic reaction. This situation is not always true as, in the case of hydrogenase and nitrogenase systems, multiple enzymatic solutions are found.59-60 Thus, APS reductase appears to be the enzyme most characteristic of the SRBs and Odom has developed a technique for the rapid detection of SRBs in the environment based on polyclonal antibodies to APS reductase.63

VL THE MECHANISM OF APS REDUCTASE Michaels et al.38-39 published two articles in which they suggested possible mechanisms for APS reductase activity: (1) direct oxidation of AMP and sulfite to APS involving the formation of an intermediate, adenosine 5'-phosphosulfite, and (2) the oxidation to sulfate of enzyme-bound sulfite, followed by the sulfate transfer to a mononucleotide, forming APS. The second possibility was considered to be the most likely as the adenosine 5'phosphosulfite molecule was known to be unstable,62 and the interaction of the sulfite molecule with the reductase, even in the absence of the mononucleotide, had been observed. It was not explained how the FADH2 was reoxidized and the involvement of the non-heme iron in the mechanism of the reductase was not indicated. This involvement of non-heme iron only became apparent when some flavin metalloproteins were showed by EPR to have a '* 1.94" type signal characteristic of reduced iron-sulfur centers.30 Also the extensive study of the sequential addition of sulfite and AMP as well as the description of the sulfite-flavin adduct formation led to the description of a mechanism for APS reductase including the involvement of an iron-sulfur center.53 The major considerations were (1) the reversible reaction of sulfite with the flavin moiety leading to the formation of an adduct in the N(5) position; (2) transfer of the sulfur group to a mononucleotide acceptor (AMP) with the formation of FADH2; (3) fast intramolecular electron transfer step, from the FADH2 to another redox center (possibly an iron-sulfur center). No evidence for the formation of the semiquinone form of FAD was found.63 The proposed mechanism could in general account for the observed UV/visible spectroscopic changes but the involvement of the iron-sulfur centers was uncertain. EPR experiments were carried out at 17 K53-62 and the effect of the addition of pHMB to the enzyme was studied.49 These experiments showed that the addition of pHMB (an agent which is known to destroy some iron-sulfur clusters) inhibited the reductase activity using ferrycianide as electron acceptor but the activity was retained in the presence of pHMB when reduced cytochrome c or MV were used as electron donor for the reduction of APS. Moreover, EPR experiments showed the existence of an isotropic signal with g = 2, in the native reductase and a "1.94" type signal was detected upon addition of AMP plus sulfite or dithionite. Also the sequential addition of pHMB, sulfite AMP, and dithionite in different

350

Chemistry and Biochemistry of Flavoenzymes

FIGURE 10. EPR spectra of several APS reductases from Desulfovibrio strains. Left panel: reductases plus sulfite plus AMP. Right panel: reductases "partially reduced" (dithionite 15 sec.)

orders was studied by EPR and visible spectroscopy as previously indicated, and the results can be summarized as follows: 1. 2. 3. 4.

The addition of sulfite causes the appearance of a small "1.94" type signal, probably due to the existence of small quantities of AMP in the native enzyme. The addition of AMP alone does not induce the formation of this EPR signal. The order of the addition of AMP and sulfite is not important. The presence of pHMB prevents formation of the "g = 1.94" EPR signal only if it is added before the addition of both AMP and sulfite. If pHMB is added after AMP and sulfite, the intensity of the "1.94" type signal is almost the same as if no pHMB is added.

The studies of enzymatic activity49 with ferrycianide, cytochrome c and MV, coupled with the addition of pHMB, showed that there are several electronic pathways among the different electron carriers and the reductase. Only the ferrycianide reduction is inhibited by pHMB and appears to require non-heme iron. Moreover, cytochrome c reduction seems to

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FIGURE 10 (continued). APS reductases "fully reduced" (dithionite > 15 min). Modulation 1 ml; microwave power 2 mW; temperature 10 K.

be dependent on the existence of super-oxide anions, formed by the monoelectronic reduction of oxygen by a reduced flavin as postulated by Massey et al.42 In the MV assay, it was considered that the reduced MV gave its electrons directly to the sulfite-enzyme complex. Considering these facts, the authors suggested a reaction sequence for APS reductase (Figure 4) emphasizing the roles of electron carriers and adduct formation. The previously proposed mechanisms appear to be essentially correct in describing the reactions of the APS reductase from the SRBs; however, they should not be viewed as a detailed mechanism of action for the reductase. Our EPR and Mossbauer studies have been directed toward further defining the mechanism of the reductase and have conclusively established that the non-heme iron is organized as two [4Fe-4S] clusters; Center I with an E0 ~0 mV and Center II with an E^ -400 to -450 mV; however, their roles in intra- and intermolecular electron transfer processes have not been resolved. In the presence of AMP plus sulfite, only non-heme iron Center I is reduced but Center II is only observed after extended exposure to dithionite at pH 9.0; thus, the function of Center II is not understood

352

Chemistry and Biochemistry of Flavoenzymes

except in the negative sense, that it does not appear to serve as an electron acceptor for the oxidation of FADH2. As [4Fe-4S] clusters are one electron acceptors, one might expect that the oxidation of FADH2 occurs by a single electron process leading to the formation of a flavin-semiquinone; however, the formation of semiquinone form of FAD has not been detected by visible or EPR spectroscopy.62-63 In this regard, it should be noted that the occurrence of a red (anionic) flavin semiquinone has been observed upon the addition of AMP to the sulfite-reduced APS reductase of T. Thioparus. The third general problem for the mechanism involves the nature of the interaction between AMP (or APS) and Center I. Kinetic, EPR and Mossbauer observations all indicated that AMP interacts significantly with the non-heme iron Center I but the exact nature of this interaction remains obscure. In stopped-flow studies with nucleotide-free reductase, (R. Bramlett and H. D. Peck, Jr., unpublished observations) it was shown that the rate of formation of the N(5) adduct was increased 100-fold by the presence of low amounts of AMP. Further studies dealing with the characterization of these intermediate states are very important for an overall appreciation of the mechanism of this central enzyme in the utilization of sulfate in nature, and we hope that in the near future these questions will be answered. The problem is even broader as the above considerations apply only to APS reductases from the SRBs of the genus Desulfovibrio; other microorganisms such as the Thiobacilli and phototropic bacteria may contain significantly different reductases but this question can only be resolved by extensive enzymological studies and determination of their primary structures by recombinant DNA techniques.

ACKNOWLEDGMENTS This work was supported by Instituto Nacional de Investigate Cientifica (J.J.C.M.), Junta Nacional de Investigate Cientifica e Tecnologica (J.J.G.M.), CEE-BAP, Contracts No. 0259-P-TT (JJ.G.M.) and No. 0269 (G.F.), Contract DEA-509-79 ER 10499 from the Department of Energy (H.D.P.), Grant GM 32187 (B.H.H.) and GM 41482 (H.D.P. and J.L.) from the National Institutes of Health and Grant DMB 8413918 (J.L. and H.D.P.) from the National Science Foundation. We thank the staff from the Fermentation Plant of the University of Georgia for growing D. baculatus 57Fe-enriched cells. The authors also wish to thank Drs. B. H. Huynh and D. V. DerVartanian for many helpful discussions.

REFERENCES 1. Peck, H. D., Jr. and Lissolo, Thierry, Assimilatory and dissimilatory sulfate reduction, in The Nitrogen and Sulfur Cycles, Cole, J. and Ferguson, S., Eds., Symposium of the Society for General Microbiology, 42, 99, 1988. 2. Kredich, N. M M Biosynthesis of cysteine, in Escherichia coli and Salmonella typhimurium Cellular and Molecular Biology, Neidhardt, F. C., Ingraham, J. L., Low, K. B., Magasanik, B., Schaechter, M., and Umbarger, H. E., Eds., Vol. 2, American Society of Microbiology, Washington, D.C., 1987, 418. 3. Tsang, M. L.-S. and Schiff, J. A., Studies of sulfate utilization by algae 18. Identification of glutathione as a physiological carrier in assimilatory sulfate reduction by Chlorella, Plant Sci. Lett., 11, 177, 1978. 4. Wilson, L. G., Asahi, T., and Bandvorski, R. S., Yeast sulfate reducing system 1. Reduction of sulfate to sulfite, /. Biol. Chem,, 236, 1822, 1961. 5. Porque, P. G., Baldestein, A., and Reichard, P., The involvement of the thioredoxin system in the reduction of methionine sulfoxide and sulfate, J. Biol. Chem., 245, 2371, 1961. 6. Rassel, M., Model, P., and Holmgren, A., Thioredoxin or glutaredoxin in Escherichia coli is essential for sulfate reduction but not for deoxyribonucleotide synthesis, J. BacterioL, 172, 1923, 1990. 7. Abrams, W. R. and Schiff, J. A., Studies of sulfate utilization by algae. II. An enzyme-bound intermediate in the reduction of adenosine-5'-phosphosulfate (APS) by cell-free extracts of wild-type Chlorella and mutants blocked for sulfate reduction, Arch. MikrobioL, 94, 1, 1973.

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8. Schmidt, A., Sulfate reduction in a cell-free system of Chlorella. The ferredoxin dependent reduction of a protein-bound intermediate by a thiosulfonate reductase, Arch. Mikrobioi., 93, 29, 1973. 9. Koguchi, O. and Tamura, G., Reduction of 5-sulfoglutathione by cyanobacterial ferredoxin-sulfite reductase, Agric. BioL Chem., 53, 783, 1989. 10. Schmidt, A., Distribution of APS sulfotransferase activity among higher plants, Plant Sci. Lett., 5, 407, 1975. 1 1 . Saidha, T., Na, S.-Q., Li, J., and Schiff, J. A., A sulphate metabolizing centre in Euglena mitochondria, Biochem. J.t 253, 533, 1988. 12. Tsang, M. L.-S. and Shiff, J. A., Studies of sulfate utilization by algae 14, Distribution of adenosine3'-phosphate-5'-phosphosulfate (PAPS) and adenosine-5'-phosphosulfate (APS) sulfotransferases in assimilatory sulfate reducers, Plant Sci. Lett., 4, 301, 1975. 13. Schmidt, A., Assimilatory sulfate reduction via 3'-phosphoadenosine-5'-phosphosulfate (PAPS) and adenosine-5'-phosphosulfate (APS) in blue-green algae, FEMS Microbiol. Lett., I, 137, 1977. 14. Truper, H. G. and Fischer, V., Anaerobic oxidation of sulphur compounds as electron donors for bacterial photosynthesis, Philos. Trans. R. Soc. (London) Ser. B, B298, 99, 1982. 15. Schmidt, A., Adenosine 5'-phosphosulfate (APS) as sulfate donor for assimilatory sulfate reduction in Rhodospirillum rubrum, Arch, Microbiol., 112, 263, 1977. 16. Schmidt, A., The adenosine-5'-phosphosulfate sulfotransferase from spinach (Spinacea oleracea L.). Stabilization, partial purification and properties, Planta, 130, 257, 1976. 17. Brunold, C. and Schmidt, A., Regulation of adenosine-5'-phosphosulfate sulfotransferase activity by H2S in Lemna minor L., Planta, 133, 85, 1976. 18. Jenni, B. E., Brunold, C., Zryd, J.-P., and Lavanchy, P., Properties and regulation of adenosine 5'phosphosulfate transferase from suspension cultures of Nicotiana sylvestris, Planta, 150, 140, 1980. 19. Deuereux, R M Delaney, M., Widdel, F., and Stahl, D. H., Natural relationships among sulfate-reducing eubacteria, J. BacterioL, 171, 6689, 1989. 20. Stetter, K. O., Lauerer, G., Thornm, M., and Neuner, A., Isolation of extremely thermophilic sulfate reducers: evidence for a novel branch of archaebacteria, Science, 236, 822, 1987. 21. Odom, J. M. and Peck, H. D., Jr., Hydrogenase, electron transfer proteins and energy coupling in the sulfate reducing bacteria, Desulfovibrio, Ann. Rev. Microbiol., 38, 551, 1984. 22. Kremer, D. R., Veenhuis, M., Fauque, G., Peck, H. D., Jr., Le Gall, J., Lampreia, J., Moura, J. J. G., and Hansen, T. A., Immunocytochemical localization or APS reductase and bisulfite reductase in three Desulfovibrio strains, Arch. Microbiol., 150, 296, 1988. 23. Kobavashi, K., Seki, Y., and Ishimoto, M., Biochemical studies on sulfate reducing bacteria XIII. Sulfite reductase from Desulfovibrio vulgaris — mechanism of trithionate, thiosulfate and sulfide formation and enzymatic properties, J. Biochem. (Tokyo), 75, 519, 1974. 24. Lee, J.-P. and Peck, H. D., Jr., Purification of the enzyme reducing bisulfite to triphionate and its identification as desulfoviridin, Biochem. Biophys. Res. Commun., 45, 583, 1973. 25. Hatchikian, E. C. and Zeikus, J. G., Characterization of a new type of dissimilatory sulfite reductase present in Thermodesulfobacterium commune, J. BacterioL, 153, 1211, 1983. 26. Trudinger, P. A., Carbon-monoxide-reacting pigment from Desulfotomaculum nigrificans and its possible relevance to sulfite reduction, /. BacterioL, 104, 158, 1970. 27. Lee, J.-P., Yi, C. S., Le Gall, J., and Peck, H. D., Jr., Isolation of a new pigment from Desulfovibrio desulfuricans (Norway 4) and its role in sulfite reduction, J. BacterioL, 115, 453, 1973. 28. Lee, J.-P., Le Gall, J., and Peck, H. D., Jr., Isolation of assimilatory- and dissimilatory-type sulfite reductases from Desulfovibrio vulgaris, J. BacterioL, 115, 529, 1973. 29. Moura, I., Lino, A. R., Moura, J. J. G., Xavier, A. V., Fauque, G., Peck, H. D., Jr., and Le Gall, J., Low spin sulfite reductases: a new homologous group of non-heme iron-siroheme proteins in anaerobic bacteria, Biochem. Biophys. Res. Commun., 141, 1032, 1986. 30. Kramer, M. and Cypionka, H., Sulfate formation via ATP sulfurylase in thiosulfate- and sulfite- disproportionating bacteria, Arch. Microbiol., 151, 232, 1988. 31. Peck, H. D., Jr., APS as an intermediate in the oxidation of thiosulfate by Thiobacillus Thioparus, Proc. Natl. Acad. Sci. U.S.A., 46, 1053, 1960. 32. Bowen, T. J., Happold, F. O., and Taylor, B. F., Studies on adenosine 5'-phosphosulfate reductase from Thiobacillus denitrificans, Biochim. Biophys. Acta, 118, 566, 1966. 33. Sklodowska, A., Polythionales and adenosine-5'-phosphosulfate formation during thiosulfate oxidation by Thiobacillus neapolitanus, Can. J. Microbiol., 34, 1283, 1988. 34. Peck, H. D., Jr., Some evolutionary aspects of inorganic sulfur metabolism. Lecture series on theoretical and applied aspects of modern microbiology. University of Maryland Press, 1966. 35. Thiele, H. H., Sulfur metabolism in thiorhodaceae V. enzymes of sulfur metabolism in Thiocapsa floridans and Chromatium species, Antonievan Leeuwenhoeck, J. Microbiol. SeroL, 34, 350, 1968.

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36. Truper, H. J. and Peck, H. D., Jr., Formation of adenylyl sulfate in phototropic bacteria, Arch. Mikrobiol, 73, 125, 1970. 37. Bias, U. and Triiber, H. G., Species specific release of sulfate from adenylyl sulfate by ATP sulfurylase and ADP sulfurylase in the green sulfurbacteria Chlorobium limicola and Chlorobium vibrioforme, Arch. Microbiol., 147, 406, 1987. 38. Michaels, G. B., Davidson, J. T., and Peck, H. D., Jr., A flavin-sulfite adduct as an intermediate in the reaction catalyzed by adenylyl sulfate reductase from Desulfovibrio vulgaris, Biochem. Biophys. Res. Commun., 39, 321, 1970. 39. Michaels, G. B., Davidson, J. T., and Peck, H. D., Jr., Studies on the mechanism of adenylyl sulfate reductase from the sulfate reducing bacterium, Desulfovibrio vulgaris, in Flavins andFlavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 555. 40. Adachi, K. and Suzuki, I M A study on the reaction mechanism of adenosine 5'-phosphosulfate reductase from Thiobacillus Thioparus, an iron-sulfur flavoprotein, Can. J. Biochem., 55, 91, 1977. 41. Swoboda, B. E. P. and Massey, V., On the reaction of the glucose oxidase from Aspergillus niger with bisulfite, /. Biol. Chem., 241, 3409, 1966. 42. Massey, V., Palmer, G., and Ballou, D., On the reaction of reduced flavins and flavoproteins with molecular oxygen, in Flavins andFlavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 349. 43. Miiller, F. and Massey, V., Flavin-sulfite complexes and their structures, J. Biol. Chem., 244, 4007, 1969. 44. Massey, V., Miiller, F., Feldberg, R., Schuman, M., Sulivan, A., Ho well, L., Mayhew, S. G., Matthews, R. G., and Foust, G. P., The reactivity of flavoproteins with sulfite. Possible relevance to the problem of oxygen reactivity, J. Biol. Chem., 244, 3999, 1969. 45. Schwenn, J. D, and Biere, M., APS reductase activity in the chromatophores of Chromatium vinosum, FEMSLett., 6, 19, 1979. 46. Lampreia, J., Moura, I., Teixeira, M., Peck, H. D., Jr., Le Gall, J., Huynh, B. H., and Moura, J. J. G., The active centers of adenylyl sulfate reductase from Desulfovibrio gigas: characterization and spectroscopic studies, Eur. J. Biochem., 188, 653, 1990. 47. Stille, W, and Truper, H. G., Adenylyl sulfate reductase is some new sulfate redoing bacteria, Arch. Microbiol,, 137, 145, 1984. 48. Ishimoto, M. and Fujimoto, D., Biochemical studies on the sulfate reducing ' ,.eria x. Adenosine-5'phosphosulfate reductase, J. Biochem. (Tokyo), 50, 299, 1961. 49. Bramlett, R. N. and Peck, H. D., Jr., Some physical and kinetic properties of adenylyl sulfate reductase from Desulfovibrio vulgaris, J. Biol. Chem., 250, 2979, 1975. 50. Lyric, R. M. and Suzuki, I., Enzymes involved in the metabolism of thiosulfate by Thiobacillus Thioparus. II. Properties of adenosine-5'-phosphosulfate reductase, Can. J. Biochem., 48, 344, 1970. 51. Truper, H. G. and Rogers, L. A., Purification and properties of adenylyl sulfate reductase from the phototropic sulfur bacterium, Thiocapsa roseopersiciana, J. BacterioL, 108, 112, 1971. 52. Peck, H. D., Jr., Deacon, T. E., and Davidson, J. T., Studies on adenosine 5'-phosphosulfate reductase from Desulfovibrio desulfuricans and Thiobacillus Thioparus: I. The assay and purification, Biochim. Biophys. Acta, 96, 429, 53. Peck, H. D., Jr. and Bramlett, R. N., Flavoproteins in sulfur metabolism, in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Eds., Elsevier/North-Holland, Amsterdam, 1982, 851. 54. Skyring, G. W., A comparison of the electrophoretic properties of the ATP sulfurylases, APS reductases and sulfite reductases from dissimilatory sulfate reducing bacteria, Can. J. Microbiol., 19, 375, 1973. 55. Speich, N. and Truper, H. G., Adenylyl sulfate reductase in a dissimilatory sulfate-reducing Archaebacterium, J. Gen. Microbiol., 134, 1419, 1988. 56. Kirchhoff, J. and Truper, H. G., Adenylyl sulfate reductase of Chlorobium limicola, Arch. Microbiol., 100, 115, 1974. 57. Huynh, B. H., Moura, J. J. G., Moura, L, Kent, T. A., Le Gall, J., Xavier, A. V., and Miinck, E., Evidence for a three iron center in a ferredoxin from Desulfovibrio gigas, J. Biol. Chem., 255, 3242, 1980. 58. Cammack, R., Dickinson, D. R., and Johnson, C. E., Evidence from Mossbauer spectroscopy and magnetic resonance on the active center of the iron sulfur protein, in Iron-Sulfur Proteins, Vol. 3, Lovenberg, W., Ed., Academic Press, N.Y., 1977, 283. 59. Prickril, B. C., He, S.-H., Li, C., Menon, N., Choi, E.-S., Przybyla, A. E., DerVartanian, D. V., Peck, H. D., Jr., Fauque, G., Le Gall, J., Teixera, M., Mowia, L, Moura, J. J. G., Patil, D., and Huynh, B. H., Identification of three distinct classes of hydrogenase in the genus Desulfovibrio, Biochem. Biophys. Res. Commun., 149, 369, 1987. 60. Joergev, R. D. and Bishop, P. E., Bacterial alternative nitrogen fixation systems, Crit, Rev. Microbiol., 16, 1, 1988.

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61. Wilson, L. G. and Bandurski, R. S., Enzymatic reactions involving sulfate, sulfite selenate and molybdate, J. Biol CHem., 233, 975, 1958. 62. Peck, H. D., Jr., Bramlett, R., and DerVartanian, D. V., On the mechanism of adenylyl sulfate reductase from the sulfate reducing bacterium, Desulfovibrio vulgaris, Z. Naturforsch., 27, 1084, 1972. 63. Lampreia, J., Moura, I., Xavier, A. V., Le Gall, J., Peck, H. D., Jr., and Moura, J. J. G., unpublished results.

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Chapter 12

THE STEREOCHEMISTRY OF THE PROSTHETIC GROUPS OF FLAVOPROTEINS Emil F. Pai

TABLE OF CONTENTS I.

Introduction

358

II.

Biochemical Studies with 5-Deaza-Analogues A. 5-Deaza-5-carba-flavins B. 5-Deaza-5-carba-7,8-didemethyl-8-hydroxy-flavins C. 5-Deaza-5-carba-8-demethyl-8-hydroxy-flavins

359 359 359 360

III.

X-Ray Crystallographic Studies

363

IV.

Discussion

363

Acknowledgments

364

References

364

358

Chemistry and Biochemistry of Flavoenzymes

INTRODUCTION One of the characteristics of enzymic reactions is their specificity. Enzymes, in general, show extraordinary chiral recognition and use complex interactions to achieve close to perfect steric control of the reactions they catalyze. Sophisticated methods have been developed which allow to explore these fascinating examples of stereochemistry.] Since coenzyme molecules, like the nicotinamide and flavin nucleotides, are ubiquitous in all living systems and involved in many different reactions, their behavior is especially interesting in this respect. By far most of the reactions in which these coenzymes are involved are redox processes. Pure electron transfer, of course, does not lend itself to stereochemical analysis. As soon as hydrogen transfer occurs, however, deuterium or tritium label experiments are possible. For the nicotinamide nucleotide coenzymes, Westheimer and co-workers2-3 showed that there is direct hydrogen transfer between substrate and coenzyme. They also found that this transfer is reversibly stereospecific with respect to the redox site of the coenzyme. Pullman et al.4 established that this site was position C(4) of the pyridine ring. In the reduced form of this coenzyme this is a prochiral center according to the definition of Hanson.5 Cornforth and co-workers6 proved its absolute configuration by converting enzymically deuteriumlabeled NADH to succinic acid and comparing the ORD curve of the product with the corresponding curve of an authentic sample. When more data had been accumulated, generalizations about enzymic redox reactions involving nicotinamide nucleotides were drawn.7"10 A very comprehensive review on the subject, including many flavoproteins, has recently been published by You.11 Another class of coenzymes involved in redox conversions are the flavin nucleotides. They are "at the crossroads of biological redox chemistry."12-13 C(4a) and N(5) of the isoalloxazine ring system are the key loci of interaction between the flavin coenzymes and the enzyme's substrates. This has been shown applying enzymic14'18 as well as bioorganic model studies.19-20 For many flavoproteins the transfer of a hydride equivalent to N(5) has been proposed as part of their mechanism. As C(4) in the nicotinamide ring, position 5 in flavins is prochiral (Scheme 1).

SCHEME 1. Si-face and re-face of (A) flavin, (B) 5-deaza-5-carba-flavin, (C) 5-deaza-5-carba-7,8-didemethyl8-hydroxy-flavin, and (D) 5-deaza-5-carba-8-demethyl-8-hydroxy-flavin.

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Until recently, information on the stereochemical course of reactions catalyzed by flavoenzymes was restricted to substrates (including nicotinamide coenzymes). The obvious question, whether any particular enzyme always interacts with its substrate(s) via the same face of the flavin ring, was much more difficult to answer. When isotopic labels were introduced into the flavin ring, the labile nature of the bond formed between the nitrogen and hydrogen atoms led to instantaneous exchange with water molecules. This precluded experiments analogous to the classical ones performed on nicotinamide coenzymes.

II. BIOCHEMICAL STUDIES WITH 5-DEAZA-ANALOGUES A. 5-DEAZA-5-CARBA-FLAVINS In a first attempt to prevent loss of label to the solvent 5-deaza-5-carba-analogues of the flavin coenzymes were synthesized21"24 (see also Chapter 4 by Yoneda and Kokel, Vol. I of this series). Several research groups succeeded in demonstrating direct hydrogen transfer from substrates to these enzyme-bound coenzyme analogues22'25 28 (Scheme 2). When they,

SCHEME 2. Direct hydrogen transfer from substrates to position 5 of enzyme-bound coenzyme analogues.

however, tried to establish relative stereospecificities by labeling the flavin analogues while bound to a given enzyme, transferring them to another protein matrix, and analyzing them there, they realized that rapid exchange of label between reduced and oxidized 5-deazaflavins occurred.23 Mixtures of oxidized and reduced 5-deazaflavins undergo rapid two-electron disproportionation; the corresponding bimolecular rate constant at 0°C is 22 M~l s"1. This scrambling of label was too fast to allow accurate measurements. B. 5-DEAZA-5-CARBA-7,8-DIDEMETHYL-8-HYDROXY-FLAVINS In 1978 it was reported that methanogenic bacteria contained a 5-deaza-5-carba-riboflavm derivative.29 The authors termed the complete 5-deaza-5-carba-7,8-didemethyl~8-hydroxyisoalloxazine coenzyme F420, because of its strong absorption at this wavelength. The part equivalent to riboflavin was named F0; it is phosphorylated at the 5'-hydroxyl group. Linked to this phosphate group is a lactyl oligo-^-glutamyl tail with varying numbers of glutamates. Jacobson and Walsh30 described that the exchange of label between the reduced and oxidized forms of this flavin analogue is several orders of magnitude slower than has been found for 5-deaza-5-carba-flavin. This is probably due to the development of a negative charge in the ring upon reduction. Taking advantage of this effect, first relative31 and later absolute32 stereochemistries of cofactor F420-dependent enzymes were determined (Table 1). All enzymes investigated in these studies3134 used the same face of the tricyclic ring system to interact with the substrates tested. In order to establish the absolute stereochemistry, a selenium-containing hydrogenase35 was used to stereospecifically introduce a deuterium label at position C(5) of the modified isoalloxazine system. The resulting reduced riboflavin analogue was then chemically degraded to the [4-2HJ-3,4-dihydro-7-hydroxy-l-hydroxyethylquinoline (Scheme 3). By comparing the ORD spectrum of the product to the corre-

360

Chemistry and Biochemistry of Flavoenzymes TABLE 1 Stereospecifkity of Flavin-Substrate Interactions for 5-Deaza-5-Carba-7,8Didemethyl-8-Hydroxy-Flavin Dependent or Converting Enzymes Enzyme

Source

Hydrogenase Hydrogenase NADP-reductase Formate dehydrogenase NADH-oxidoreductase

Methanococcus vaniellu Methanobacterium thermoautotrophicum Methanococcus vaniellii Methanobacterium formicicum Beneckea harveyi

Substrate

Side

Hydrogen Hydrogen NADP Formate NADH

si si si si re

Ref. 32 33, 34 32 34 15, 32

SCHEME 3. Degradative method for determination of chirality of enzyme-labeled dihydro-5-deaza-5-carba-flavin analogue.

spending curves of authentic, chemically synthesized samples,36 Yamazaki and co-workers32 found that it was the si-side of the flavin system which was involved in the reactions. C. 5-DEAZA-5-CARBA-8-DEMETHYL-8-HYDROXY-FLAVINS A major drawback of the system based on coenzyme F420 and described above is the fact that the analogue is easily available only at the riboflavin and FMN level; both can be obtained through cleavage of the native cofactor.37 As neither the riboflavin nor the FMN form are converted to the FAD level by the established and most widely used enzymic procedures,3839 work with FAD dependent flavoenzymes would have required chemical adenylylation. Fortunately, it is possible to take advantage of the redox properties of coenzyme F420 and at the same time use the Brevibacter synthetase system. Slightly modifying the procedure of Ashton and Brown40 pure 5-deaza-5-carba-8-demethyl-8-hydroxy-riboflavin can be synthesized in good yield.39-41 In this analogue the methyl group in position 7 is not removed making the riboflavin as well as the FMN form rather good substrates for the enzymic

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FIGURE 1. Stereoscopic representation of the arrangement of NADPH and FAD at the active site of glutathione reductase from human erythrocytes.43

conversion to FAD if the pH is carefully controlled and kept at 5.9.39 The structural features are as close as possible to those of native riboflavin nucleotides, permitting easier reconstitution of apoproteins with the analogue. Two more experiments were crucial for the successful development of a method suitable for the determination of absolute stereospecificity of flavoenzyme prosthetic groups. The first was the finding by Ghisla et al. 42 that about 90% of tritium label is incorporated on one side of the ring system when medium chain acyl-CoA dehydrogenase reconstituted with 5-deaza-5-carba-FAD is treated with NaB3H4. This provided a very convenient way to obtain stereospecifically labeled flavin analogues. The second important result was the elucidation of the mode of binding to substrates and of the stereochemical framework for the reaction mechanism of the flavoenzyme glutathione reductase from human erythrocytes by crystallographic methods. Using differenceFourier techniques it could be shown that the nicotinamide ring of the substrate NADPH interacts with the re-side of the isoalloxazine ring43 (Figure 1). This protein was the first flavoenzyme for which the absolute stereochemistry of the complete catalytic reaction became known.43'45 The successful analysis established a point of reference for all other flavoenzymes (Table 3). (See also Chapter 6, this volume.) As a first step the relation between the unknown stereospecificity of medium chain acylCoA dehydrogenase and the known one of glutathione reductase had to be determined. As the method employed has proven to be applicable to most enzymes tested so far, the general procedure will be described.46'47 Medium chain acyl-CoA dehydrogenase reconstituted with 5-deaza-5-carba-FAD was incubated with NaB3H4. The labeled analogue was released by heat-denaturing the protein matrix. Precipitate was removed by centrifugation. Apoglutathione reductase was then added to the solution. After gel filtration, NAD + was used to reoxidize the modified isoalloxazine ring. The protein was separated from the smaller molecules by another gel filtration column or by HPLC. More than 90% of the radioactivity was recovered in the nucleotide fraction proving that NaB3H4 transfers the tritium label to the re-side of the flavin analogue bound to the medium chain acyl-CoA dehydrogenase. An alternative method to introduce the isotropic label uses tritiated glucose and glucose oxidase reconstituted with the appropriate analogue.46 Its advantage is that all its constituents are commercially available. Scheme 4 gives a "flowchart" for the procedure of determining the respective stereospecificity of the prosthetic groups of flavoenzymes. If medium chain acyl-CoA dehydrogenase (MCAD) or glucose oxidase (GO) are used to label the flavin analogue, radioactivity coeluting with the small-molecules-fractions indicates respecificity of the new enzyme. In contrast, if the majority of counts is found in the protein peak, then the tested flavoenzyme and the substrate used interact on the si-face of the enzyme's prosthetic

362

Chemistry and Biochemistry of Flavoenzymes

SCHEME 4.

Reaction scheme for determination of stereospecificity.

group. The prerequisites for successful analyses are reasonable stability of the apoform of the enzyme to be tested, and formation of a stable complex between apoenzyme and analogue. For the large majority of flavoenzymes analyzed so far this has been found to be the case.46'47 The results of these experiments are shown in Table 2.

363

Volume III TABLE 2 Stereospecificity of Flavin-Substrate Interactions for FMN or FAD Dependent Enzymes Determined by Biochemical Methods

Medium chain acyl-CoA dehydrogenase

Pig kidney

Glucose oxidase Mercuric ion reductase Thioredoxin reductase p-Hydroxybenzoate hydroxylase Melilotate hydroxylase Anthranilate hydroxylase MHPCa-oxygenase Cyclohexanone oxygenase D-Amino acid oxidase L-Lactate oxidase D-Lactate dehydrogenase

Aspergillus niger Pseudomonas aeruginosa Escherichia coli Pseudomonas fluorescens Pseudomonas sp. Trichosporum cutaneum Pseudomonas sp. MA-1 Acinetobacter NCIB Pig kidney Mycobacterium smegmatis Megasphera elsdenii

a

Substr.

Source

Enzyme

Acyl-CoA NaBH 4 Glucose NADP NADP NADP

NAD NADP

NAD NADP Pyruvate + NH4 Pyruvate Pyruvate

Side

Ref.

re re re re re re re re re re re si si

46 46 46 46 46 46 46 46 47 47 47 47 47

2-Methyl-3-hydroxypyridine-5-carboxylic acid.

TABLE 3 Stereospecificity of Flavin-Substrate Interactions for FMN or FAD Dependent Enzymes Determined by X-ray Crystallography Enzyme Glutathione reductase Flavocytochrome b2 Trimethylamine dehydrogenase Glycolate oxidase Medium chain acyl-CoA dehydrogenase Mercuric ion reductase

Source Human erythrocytes Baker's yeast Bacterium W3A1 Spinach Pig liver Bacillus sp. RC607

Substr. NADP Pyruvate Trimethylamine Glycolate Octanoyl-CoA NADP

Side

Ref.

re si si si re re

43 48 49 50 51 52

III. X-RAY CRYSTALLOGRAPHIC STUDIES As already mentioned above, the first information about the absolute Stereospecificity of the prosthetic group of a flavoenzyme came from X-ray crystallographic studies on flavoenzyme-substrate complexes. Apart from glutathione reductase43"45 the binding sites of substrates have been reported for flavocytochrome b2 from baker's yeast,48 a bacterial trimethylamine dehydrogenase,49 glycolate oxidase from spinach,50 and for medium chain acylCoA dehydrogenase from pig liver mitochondria51 as well as for mercuric ion reductase from Bacillus sp. RC60752 (Table 3). The very closely related medium chain acyl-CoA dehydrogenase from pig kidney and mercuric ion reductase from Pseudomonas aeruginosa had been tested with the labeling method.46 The results of the biochemical experiments agree with the findings of the crystallographic work; both enzymes interact via the re-face of the flavin ring with their respective substrates.

IV. DISCUSSION So far, there is only very limited information available on the absolute stereochemistry of the prosthetic groups of flavoenzymes; especially, when compared to the nicotinamide

364

Chemistry and Biochemistry of Flavoenzymes

nucleotides.11 On the other hand, the enzymes probed include FMN- and FAD-dependent ones, as well as those employing coenzyme F420. Their reaction mechanisms are thought to involve carbanion intermediates or hydride transfer. Although the number of enzymes in each class is small, there seem to be some remarkable distinctions. All the F420-dependent enzymes from methanogenic bacteria analyzed perform stereospecific transfer of hydrogen at the si-face of the prosthetic group. The flavoenzymes interacting with a-hydroxy acids also employ the si-side of the isoalloxazine ring. Those flavoenzymes reacting with a pyridine nucleotide all use the re-face of the flavin for this reaction. Together with sequence homologies and the results of structural analyses by X-ray crystallography the findings suggest that there are a few classes of flavin-binding domains. It is reasonable to assume that during evolution the catalytic set-up required for a given enzymic reaction has been preserved. The functional groups which determine specificity could have been adapted from one enzyme to another. The method described in Scheme 4 might also be applied to electron transfer proteins like flavodoxins, in order to test the accessibility of the flavin groups to solvent. Preliminary experiments with flavodoxin indicate a clear preference for removal of label from the siside of the flavin ring when the reduced analogue is reoxidized by ferricyanide.54 Studies are in progress with acetolactate synthase from Escherichia coli, an enzyme that does not catalyze a redox reaction, but still needs flavin for activity. When the native flavin is replaced by the 5-deaza-5-carba-8-demethyl-8-hydroxy-analogue, the three isoenzymes of acetolactate synthase differ greatly in their response (90 to 30% of original activity). It is also possible to test the accessibility of the flavins when bound to this protein.53 Hopefully, the methods developed for stereochemical analysis will turn out to be of help to the flavinologist in more than one way.

ACKNOWLEDGMENTS I am very grateful to D. J. Manstein for the most valuable part he played in our research into stereospecificity. I thank the many colleagues who contributed ideas or samples. I am indebted to K. C. Holmes for letting me pursue my *'hobby".

REFERENCES 1. Retey, J. and Robinson, J. A., Stereospecificity in Organic Chemistry and Enzymology, Verlag Chemie, Weinheim, Germany, 1982. 2. Fisher, H. F., Ofner, P., Conn, E. E., Vennesland, B., and Westheimer, F. H., Direct enzymic transfer of hydrogen atoms between substrates and DPN, Fed. Proc., II (Abstr.), 211, 1952. 3. Westheimer, F. H., Fisher, H. F., Conn, E. E., and Vennesland, B., The enzymatic transfer of hydrogen from alcohol to DPN, J. Am. Chem. Soc., 73, 2403, 1951. 4. Pullman, M. E., San Pietro, A., and Colowick, S. P., On the structure of reduced diphosphopyridine nucleotide, J. BioL Chem,, 206, 129, 1954. 5. Hanson, K. R., Applications of the sequence rule. I. Naming the paired ligands g.g at a tetrahedral atom X ggjJ . II. Naming the two faces of a trigonal atom Y^, J, Am. Chem. Soc., 88, 2731, 1966. 6. Cornforth, J. W., Ryback, G., Popjack, G., Donninger, C., and Schroepfer, G., Jr., Stereochemistry of enzymic hydrogen transfer to pyridine nucleotides, Biochem. Biophys. Res. Commun., 9, 371, 1962. 7. Colowick, S. P., van Eys, J., and Park, J. H., Dehydrogenation, in Comprehensive Biochemistry, Vol. 14, Florkin, M. and Stotz, E. H., Eds., Elsevier, Amsterdam, 1966, 1. 8. Bentley, R., Dehydrogenation and related processes, in Molecular Asymmetry in Biology, Vol. 2, Academic Press, New York, 1970, 1. 9. Vennesland, B., Stereospecificity in biology, Top. Curr. Chem., 48, 39, 1974.

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10. Alizade, M. A. and Brendel, K., Tentative classification of NAD(P)-linked dehydrogenases in regard to their stereochemistry of hydrogen transfer to the coenzyme, Naturwissenschaften, 62, 346, 1975. 11. You, K., Stereospecificity for nicotinamide nucleotides in enzymatic and chemical hydride transfer reactions, CRC Crit, Rev. Biochem., 17, 313, 1984. 12. Walsh, C., Flavin coenzymes: At the crossroads of biological redox chemistry, Ace. Chem. Res,, 13, 148, , 1980. 13. Bruice, T. C., Mechanisms of flavin catalysis, Ace. Chem. Res., 13, 256, 1980. 14. Jorns, M. S. and Hersh, L. B., Demonstration of enzymic hydrogen transfer from substrate to a flavin, J. Am. Chem. Soc., 96, 4012, 1974. 15. Fisher, J. and Walsh, C., Enzymatic reduction of 5-deazariboflavin from reduced nicotinamide adenine dinucleotide by direct hydrogen transfer, /. Am. Chem. Soc., 96, 4345, 1974. 16. Thorpe, C. and Williams, C. H., Jr., Spectral evidence for a flavin adduct in a monoalkylated derivative of pig heart lipoamide dehydrogenase, /. Biol. Chem., 251, 7726, 1976. 17. Hersh, L. B. and Walsh, C., Preparation, characterization, and coenzymic properties of 5-carba-5-deaza and 1-carba-l-deaza analogs of riboflavin, FMN, and FAD, Methods EnzymoL, 66, 277, 1981. 18. Pompon, D. and Lederer, F., Deazaflavins as cofactors for flavocytochrome b2 from baker's yeast, Eur. J. Biochem., 96, 571, 1979. 19. Briistlein, M. and Bruice, T, C., Demonstration of a direct hydrogen transfer between NADH and a deazaflavin, J. Am. Chem. Soc., 94, 6548, 1972. 20. Loechler, E. L. and Hollocher, T. C., Reduction of flavins by thiols. 1. Reaction mechanism from the kinetics of the attack and breakdown steps, J. Am. Chem. Soc., 102, 7312, 1980. 21. O'Brien, D., Weinstock, L., and Cheng, C., Synthesis of 10-deazariboflavin and related 2,4-dioxopyrimido[4,5-b]quinolines, J. Heterocyclic Chem., 7, 99, 1970. 22. Hersh, L. B. and Jorns, M. S., Use of 5-deaza-FAD to study hydrogen transfer in the D-amino acid oxidase reaction, J. Biol, Chem., 250, 8728, 1975. 23. Spencer, R., Fisher, J., and Walsh, C., Preparation, characterization, and chemical properties of the flavin coenzyme analogues 5-deazariboflavin, 5-deazariboflavin-5'-phosphate, and 5-deazariboflavin-5'diphosphate,5'^5'-adenosine ester, Biochemistry, 15, 1043, 1976. 24. Ashton, W. T., Brown, R. D., and Tolman, R. L., New routes to l-deoxy-(3,4-dihydro-7,8-dimethyl2,4-dioxopyrimido[4,5-b]-quinolin-10(2H)yl)-D-ribitols (5-deazariboflavins), /. Heterocyclic Chem., 15, 489, 1978. 25. Jorns, M. S. and Hersh, L. B., Af-methylglutamate synthetase. Substrate-flavin hydrogen transfer reactions probed with deazaflavin mononucleotide, J. Biol. Chem., 250, 3620, 1975. 26. Jorns, M. S. and Hersh, L. B., Nucleophilic addition reactions of free and enzyme-bound deazaflavin, /. Biol Chem., 251, 4872, 1976. 27. Hersh, L. B., Jorns, M. S., Peterson, J., and Currie, M., The formation of a semiquinone form of deaza-FAD bound to D-amino acid oxidase, /. Am. Chem. Soc., 98, 865, 1976. 28. Fisher, J., Spencer, R., and Walsh, C., Enzyme-catalyzed redox reactions with the flavin analogues 5deazariboflavin, 5-deazariboflavin-5'-phosphate, and 5-deazariboflavin-5'-diphosphate,5'—»5'-adenosine ester, Biochemistry, 15, 1054, 1976. 29. Eirich, L. D., Vogels, G. D., and Wolfe, R., Proposed structure for coenzyme F420 from Methanobacterium, Biochemistry, 17, 4583, 1978. 30. Jacobson, F. and Walsh, C., Properties of 7,8-didemethyl-8-hydroxy-5-deazaflavins relevant to redox coenzyme function in methanogen metabolism, Biochemistry, 23, 979, 1984. 31. Yamazaki, S., Tsai, L., Stadtman, T. C., Jacobson, F. S., and Walsh, C. T., Stereochemical studies of 8-hydroxy-5-deazaflavin-dependent NADP + reductase from Methanococcus vaniellii, J. Biol. Chem., 255, 9025, 1980. 32. Yamazaki, S., Tsai, L., Stadtman, T. C., Teshima, T., Nakaji, A., and Shiba, T., Stereochemical studies of a selenium-containing hydrogenase from Methanococcus vaniellii: Determination of the absolute configuration of C-5 chirally labeled dihydro-8-hydroxy-5-deazaflavin cofactor, Proc. Natl. Acad. Sci. U.S.A., 82, 1364, 1985. 33. Livingston, D. J., Fox, J. A., Orme-Johnson, W. H., and Walsh, C. T., 8-Hydroxy-5-deazaflavinreducing hydrogenase from Methanobacterium thermoautotrophicum. 2. Kinetic and hydrogen-transfer studies, Biochemistry, 26, 4228, 1987. 34. Schauer, N. L., Ferry, J. G., Honek, J. F., Orme-Johnson, W. H., and Walsh, C., Mechanistic studies of the coenzyme F420 reducing formate dehydrogenase from Methanobacterium formicicum, Biochemistry, 25, 7163, 1986. 35. Yamazaki, S., A selenium-containing hydrogenase from Methanococcus vaniellii. Identification of the selenium moiety as a selenocysteine residue, J. Biol. Chem., 257, 7926, 1982. 36. Teshima, T., Nakaji, A., Shiba, T., Tsai, L., and Yamazaki, S., Elucidation of Stereospecificity of a selenium-containing hydrogenase from Methanococcus vaniellii — Syntheses of (R)- and (S)-[4-2H!]-3,4dihydro-7-hydroxy-l-hydroxyethylquinolinone, Tetrahedron Lett., 26, 351, 1985.

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37. Eirich, L. D., Vogels, G. D., and Wolfe, R. S., Distribution of coenzyme F420 and properties of its hydrolytic fragments, J. Bacterial., 140, 20, 1979. 38. Walsh, C., Fisher, J., Spencer, R., Graham, R. W., Ashton, W. T., Brown, J. E., Brown, R. D., and Rogers, E. F., Chemical and enzymatic properties of riboflavin analogues, Biochemistry, 17, 1942, 1978. 39. Manstein, D. J. and Pai, E. F., Purification and characterization of FAD synthetase from Brevibacterium ammoniagenes, J. Biol. Chem., 261, 16169, 1986. 40. Ashton, W. T. and Brown, R. D., Synthesis of 8-demethyl-8-hydroxy-5-deazaflavins, J. Heterocyclic Chem., 17, 1709, 1980. 41. Ashton, W. T., Brown, R. D., Jacobson, F., and Walsh, C., Synthesis of 7,8-didernethyl-8-hydroxy-5deazariboflavin and confirmation of its identity with the deazaisoalloxazine chromophore of Methanobacterium redox coenzyme F420, J. Amer. Chem. Soc., 101, 4419, 1979. 42. Ghisla, S., Thorpe, C., and Massey, V., Mechanistic studies with general acyl-CoA dehydrogenase: Evidence for the transfer of the (i-hydrogen to the flavin N(5)-position as a hydride, Biochemistry, 23, 3154, 1984. 43. Pai, E. F. and Schulz, G. E., The catalytic mechanism of glutathione reductase as derived from X-ray diffraction analyses of reaction intermediates, /. Biol. Chem., 258, 1752, 1983. 44. Pai, E. F., Karplus, P. A., and Schulz, G. E., Crystallographic analysis of the binding of NADPH, NADPH fragments, and NADPH analogues to glutathione reductase, Biochemistry, 27, 4465, 1988. 45. Karplus, P. A., Pai, E. F., and Schulz, G. E., A crystallographic study of the glutathione binding site of glutathione reductase at 0.3 nm resolution, Eur. J. Biochem., 178, 693, 1989. 46. Manstein, D. J., Pai, E. F., Schopfer, L. M., and Massey, V., Absolute stereochemistry of flavins in enzyme-catalyzed reactions, Biochemistry, 25, 6807, 1986. 47. Manstein, D. J., Ghisla, S., Massey, V., and Pai, E. F., Stereochemistry and accessibility of prosthetic groups in flavoproteins, Biochemistry, 27, 2300, 1988. 48. Lederer, F. and Mathews, F. S., Mechanism of L-lactate dehydrogenation catalyzed by flavocytochrome b2 from baker's yeast, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 133. 49. Bellamy, H. D., Lim, L. W., Mathews, F. S., and Dunham, W. R., Studies of crystalline trimethylamine dehydrogenase in three oxidation states and in the presence of substrate and inhibitor, /. Biol. Chem., 264, 11887, 1989. 50. Lindqvist, Y. and Branden, C.-L, The active site of spinach glycolate oxidase, J. Biol. Chem., 264, 3624, 1989. 51. Kim, J.-J. P. and Wu, J., Three-dimensional structure of medium-chain acyl-CoA dehydrogenase, in this volume, 1991, chapter 8. 52 Schiering, N., Kabsch, W., Moore, M. J., Distefano, M. D., Walsh, C. T., and Pai, E. F., Structure of the detoxification catalyst mercuric ion reductase Bacillus sp. strain RC607, Nature, 352, 168, 1991. 53. Schloss, J. and Pai, E. F., Stereospecific reduction of acetolactate synthase, FASEB J., 4, A2283, 1990. 54. Manstein, D. J., unpublished results.

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STRUCTURE AND MECHANISM OF SPINACH GLYCOLATE OXIDASE Ylva Lindqvist

TABLE OF CONTENTS I.

Introduction A. General B. Substrates and Inhibitors C. Properties of the Coenzyme

368 368 368 369

II.

Description of the Structure A. Crystallization B. Reliability of the Model C. The Overall Structure of Glycolate Oxidase D. Secondary Structures E. Solvent Accessibility F. Flexibility G. Water Structure H. Subunit Packing I. FMN Binding J. The Substrate Pocket K. The Catalytic Mechanism

369 369 369 370 370 373 373 373 374 374 376 378

III.

Comparison with Other FMN-Binding Enzymes A. (3/a-Barrel Similarity Between Trimethylamine Dehydrogenase and Glycolate Oxidase B. Structural Comparison of Glycolate Oxidase and Flavocytochrome b2 1. Overall Similarities 2. Active-Site Geometry and Substrate Binding 3. Mechanistic Implications References

380 380 381 381 382 385 385

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I. INTRODUCTION A. GENERAL Glycolate oxidase (glycolate:oxygen oxidoreductase E.G. 1.1.3.1) is a peroxisomal flavoenzyme that catalyzes the oxidation of a-hydroxy acids to the corresponding a-ketoacids. Reoxidation of the reduced enzyme by molecular oxygen produces H2O2 and closes the catalytic cycle. In green plants, glycolate oxidase is one of the key enzymes in photorespiration where it oxidizes glycolate to glyoxylate. Net photosynthesis is drastically reduced due to this metabolic pathway initially catalyzed by the oxygenase reaction of ribulose-1,5-bisphosphate carboxylase/oxygenase.J Pure enzyme has been prepared from spinach,2 pea,3 pumpkin4 and cucumber5 cotelydons, Nopalea dejecta6 (a CAM plant), pig liver,7 rat liver,8 chicken liver,9 and human liver. 10 - 11 The vertebrate enzyme is believed to be involved in the metabolic production of oxalate by the oxidation of glycolate through glyoxylate. Inhibition of the enzyme might thus be of potential use12 in the treatment of those disease states (calcium oxalate kidney stones and primary hyperoxalurias13) in which the pathological consequences are due to the crystallization of calcium oxalate. Glycolate oxidase molecules from human liver,10 cucumber,5 and spinach2 are present in solution as tetramers and/or octamers of identical subunits of molecular weight around 43,000 Da. The enzyme is very basic, the isoelectric point determined for the pea enzyme is 9.6.3 The optimum pH value for the spinach glycolate oxidase is 8.3,14 and for the human liver enzyme, 8.8.11 The amino acid sequence of the spinach polypeptide chain has been determined from peptide sequencing15 and inferred from the DNA sequence of a cDNA clone.16 B. SUBSTRATES AND INHIBITORS The highest substrate specificity is found for glycolate followed by L-lactate, glyoxylate, and longer straight chain a-hydroxy acids up to a-hydroxy caproate.3-10'11'14'17"18 Aldehyde-bisulfite addition compounds, a-hydroxy sulfonates, are effective inhibitors in the enzymatic oxidation of glycolate.19 Except for the formaldehyde compound, they react with enzymatically produced glyoxylate to form glyoxylate-bisulfite which is the actual inhibitor.20 A thorough investigation on the inhibition of pig liver enzyme by mono- and dicarboxylic acids has been conducted.18 Straight chain monocarboxylic acids are noncompetitive inhibitors with glycolate and glyoxalate as the variable substrate and DCIP (dichlorophenol indophenol) as the electron acceptor. The inhibition constants become progressively lower as the alkyl chain becomes larger. Dicarboxylic acids are competitive inhibitors with respect to glycolate and noncompetitive with respect to glyoxalate. The affinity of the enzyme for oxalate is high compared to its affinity for monocarboxylic acids. The inhibition constants for the dicarboxylic acids increase with the number of carbon atoms. DL-p-phenyllactate and /V-octyloxamate were shown to be noncompetitive reversible inhibitors of glycolate oxidation.10 2-Hydroxy-3-butynoate has been shown to be an irreversible inhibitor of glycolate oxidase10'21 through covalent addition to FMN at position C(4a) and N(5).22 3-Butynoate, lacking the hydroxygroup, is a competitive inhibitor.21 An investigation of the inhibition of human liver glycolate oxidase by 70 compounds of three types, substituted glycolic, oxyacetic, and glyoxylic acid type, has been conducted23 as well as inhibition by a huge range of other compounds.24-25 Besides organic anions (e.g., heptanoate, oxalate), inorganic anions SO2", SO^", Cl~,

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PO;}^ and AsO^ can also bind to the enzyme as seen from pertubations of the flavin spectrum.18 It is suggested that they all have a common binding site. The enzyme undergoes bleaching on SO2" binding,7 due to covalent addition to N(5) of FMN.26 Phosphate and arsenate were shown to activate the enzyme.18 C. PROPERTIES OF THE COENZYME The cofactor, flavin mononucleotide, FMN, is not covalently bound to the protein and it can readily be reversibly dissociated from the protein by using high ionic strength media.2 Glycolate oxidase has an unusual absorption spectrum27 with a large hypsochromic shift from 370 nm in free FMN to 346 nm in FMN bound to glycolate oxidase and a low extinction coefficient at 448 nm which suggests that the flavin N(3)-H group of the FMN coenzyme is ionized at pH 8.3. The pKa for FMN bound to glycolate oxidase was determined to be 6.5, about 4 pH units lower than for the free flavin.7'28 In general, flavoprotein oxidases stabilize a negative charge at the N1-C2=O locus29 as can be demonstrated by (1) formation of a flavin N(5)-sulfite adduct, (2) stabilization of the anionic flavosemiquinone on enforced one-electron reduction, and (3) stabilization of the benzoquinoid form of 8-mercapto-flavin. Glycolate oxidase behaves accordingly.28 The redox potentials of glycolate oxidase are shifted markedly more positive of those of unbound FMN as compared to other flavin oxidases,30 the midpoint potential at pH 7.1 in 0.1 M phosphate is -0.06 V. The following presentation is essentially a review of References 31 to 36.

II. DESCRIPTION OF THE STRUCTURE A. CRYSTALLIZATION Two crystal forms have been obtained for glycolate oxidase. The crystals used in the structure determination, the holoenzyme, were obtained using tertiary butanol as precipitating agent. They are tetragonal with unit cell dimensions of a=b=148.1 A and c=135.1 A. The space group is 1422 with one subunit in the asymmetric unit. The other crystal form, of the apoenzyme, was obtained with ammonium sulfate as precipitating agent and has space group P422 with two subunits in the asymmetric unit, cell dimensions a=b=145.5 A, c=103.5 A. B. RELIABILITY OF THE MODEL Native data to 2 A resolution were collected at the synchrotron radiation source in Daresbury, U.K. The initial isomorphous phases from two heavy atom derivatives were improved using solvent flattening. From the resulting electron density map, a model was built and was refined using restrained parameter reciprocal space least squares. A final R-value of 18.9% was obtained in the range 5.5 to 2.0 A. This R-value is rather high partly due to the fact that all measured reflections larger than one sigma are included, i.e., the error in the measured data is rather high. This also implies that the maximum coordinate error of about 0.25 A calculated from a Luzatti plot37 gives a too high estimate of the error. A good estimate of the quality of the model can be obtained from the final electron density maps. Figure 1 shows the electron density map ([2|F0-Fc|]ac) of the coenzyme at a contour level of 0.44e/A.3 Figure 1 is representative of the quality of the electron density map, except for the C-terminal ten amino acids and a surface loop consisting of residues 189 to 197, where interpretable electron density is missing. A few solvent exposed side chains also have ill-defined electron density. The final model consists of residues 1 to 188 and 198 to 359, FMN and 298 water molecules, comprising a total of 3020 nonhydrogen atoms. The stereochemistry of this model is close to ideal geometry with an rms deviation of 0.015 A in bond lengths and 2.6° in bond angles.

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FIGURE 1. The electron density map ([2|F0|-|Fc|]ac) of the coenzyme FMN at a contour level of 0.44e/A3

C. THE OVERALL STRUCTURE OF GLYCOLATE OXIDASE The glycolate oxidase molecules in the crystals are octamers with a diameter of approximately 100 A and with strict 422 symmetry in the subunit arrangement. The 369 amino acid residues of the glycolate oxidase polypeptide chain are folded into the p/ot-barrel structural motif with eight parallel (i-strands and eight helices first described for triosephosphate isomerase.38 A schematic diagram of the subunit structure, as obtained from the high resolution structure determination, is shown in Figure 2A seen from a side view perpendicular to the p/a-barrel axis. Figure 2B is a stereo view of the Ca-atoms looking down the 3/a-barrel axis. The main features are the eight helices, strands, and the connecting loops of the barrel structure. The remaining residues have been assigned to mainly two regions. The first region comprises 70 residues at the N-terminal. The second region is part of a long peptide segment of 45 residues between strand four and helix four of the barrel. Figure 2A also illustrates that most of the structure not belonging to the barrel is located outside the carboxyl end of the (3-strands of the barrel and forms a "lid" on the barrel at that end, which partly shields the active site. Residues 189 to 197 and 359 to 369 are not visible in the electron density map and presumably have a flexible conformation. D. SECONDARY STRUCTURES Figure 3 shows the sequence of glycolate oxidase and the corresponding secondary structure elements. The most prominent features of the structure are the eight helices and strands in the £/ a-barrel but there are a number of additional secondary structures present. There are in total 14 a-helices ranging in length from 5 to 22 residues. As is common in soluble a/£-proteins the a-helices in glycolate oxidase are exposed to the solvent and are thus amphiphilic with the exception of helix F. This a-helix, which binds the phosphate group of FMN at the helix N-terminus, is almost buried and is part of an unusual helixturn-helix motif. The turn from helix F to helix 8 is extremely tight and this conformation puts severe restrictions on the residues involved, i.e., Gly-322 has to be conserved for steric reasons and Gly-319 for conformational reasons in this type of structure. Helix 6 in the barrel has a definite kink at residue 267. Instead of the regular hydrogen bonds, a water molecule is bound between O 267 and N 271.

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FIGURE 2A. Schematic diagram of the subunit structure of glycolate oxidase seen from a side perpendicular to the barrel axis. Cylinders represent a-helices, arrows (3-strands. FMN, bound at the carboxy end of the (3-sheet, is shown as a ball and stick model. The circle in broken line outside the N(5)-position of the flavin is the position of a peak in a difference Fourier map to low resolution corresponding to a substrate analogue, thioglycolate.

FIGURE 2B. barrel axis.

Stereo view of the Cct-atoms of glycolate oxidase seen looking down the

A stereo view of the barrel strands is shown in Figure 4. Built up from hydrophobic residues (with some exceptions at the ends), these strands form the core of the subunit against which the eight barrel helices pack. The angle between successive strands is around 20° except between strand 1 and 2 which is smaller —10°, and between 7 and 8, and 8 and 1 which are larger, —30°. This asymmetry is correlated with the packing of the isoalloxazine

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FIGURE 3. The amino acid sequence of glycolate oxidase and the corresponding secondary structure elements labelled as in Figure 2A. Cylinders represent a-helices, arrows, (3-strands.

FIGURE 4. Stereo representation of p-strands numbered 1 to 8 in the (3/a barrel. For clarity only Cp of the side chains are included. Hydrogen bonds are indicated by dashed lines.

ring of FMN against strand one and the following loop, and the binding of the phosphate group of FMN in the cleft where the loops from strand seven and strand eight turn away from the barrel. The twist of the strands with respect to the barrel axis is of the order of 35° as expected39 and the strands form an almost symmetrical circular barrel. Other elements of (5-structure present are a 3-hairpin loop (residues 62 to 69) covering the bottom of the barrel and two antiparallel pieces of strand comprising amino acids 45 to 47 and 354 to 356. In addition to a-helices and (3-sheets, proteins contain 310-helices and reverse turns,

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with 310-helices being a subset of reverse turns. There are five 310-helices which are three residues long in glycolate oxidase and furthermore, as is commonly observed, several of the a-helices have residues with 310-helical conformation at their ends. The longest 310helix, containing four residues, is such an extension of a-helix C. One of the three-residue 310-helices (254 to 256) is lefthanded. In the barrel there are two categories of loops; those going from the p-strands to the ahelices and those going from the a-helices to the strands. The number of residues in the loops of the second kind are few, between 3 and 6 (average 4) residues, securing the stability of the structure. Practically all of the active site residues are in the loops going from the p-strands to the a-helices. These loops are generally longer, between 4 and 13 (average 8) residues for seven of the loops and in one case, between p-strand four and a-helix four, there is a long excursion of 57 residues. It is this loop which contains the flexible residues 189 to 197. E. SOLVENT ACCESSIBILITY Solvent accessibility calculations40 show that the distribution of different amino acids between the inside and the surface of the protein is quite normal. Looking down the p/a-barrel axis, the buried hydrophobic residues form circular layers; one layer of residues from the P-strands pointing towards the barrel axis, the second layer pointing outwards from the p-strands toward the a-helices and pointing from the helices towards the strands. Due to the inclination of the P-strands, side chains from every second strand are at the same height along the barrel axis. The amino acid composition of the surface of the protein is approximately as expected41 with the exception of an unusual high amount of arginine residues which gives the enzyme very basic properties with a pl>9. Most of the basic residues that are not involved in intraor intersubunit salt bridges are lining the entrance to the active-site cleft or are involved in FMN-phosphate or substrate binding. F. FLEXIBILITY The average isotropic temperature factor B is 27.1 A2 for all protein atoms (26.5 A2 for the main-chain protein atoms, 27.8 A2 for the protein side-chain atoms and 45.2 A2 for the water molecules). Figure 5 shows the variation of B-value (average of main-chain atoms for each residue) along the chain. As can be seen in this figure, the most rigid parts of the structure are in the core of the molecule, consisting of the eight p-strands of the barrel, and a-helix C (residues 32 to 40) which makes lattice contacts with another subunit. The most flexible parts are the ends of the chain, the residues flanking the piece of sequence that is disordered in the crystal and some of the loops, especially those leading from the helices to the strands of the barrel. The average B-value for FMN is 25.2 A2 with the isoalloxazine ring being most rigid. G. WATER STRUCTURE Altogether 298 water molecules have been identified in the electron density maps of the crystals of glycolate oxidase. Of these, three are in the interior of the protein and can be considered an intrinsic part of the structure. One of these binds to one of the FMN phosphate oxygens and main chain O 306 and N 286 and one is bound in a pocket close to the C(4)O position of the isoalloxazine ring in FMN and will be discussed later. Involved in hydrogen bonds between symmetry related subunits are 14 water molecules also firmly bound as judged from their low average B-value (23.7 A2 compared to 45.2 A2 for all water molecules). In the inner solvation shell 164 more water molecules are making hydrogen bonds to the protein atoms at the surface. Hydrogen bonds to these water molecules are made from 68 other water molecules. Most of the water molecules (76%) are part of clusters,

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FIGURE 5. Temperature factor as function of residue number. The average temperature factor for the main chain atoms of each residue is plotted.

i.e., they are within 3.5 A to another water molecule. There are 57 such clusters with 2 to 18 members. H. SUBUNIT PACKING The glycolate oxidase molecules in the crystals are octamers with strict 422 symmetry in the subunit arrangement. They have the approximate shape of a cube with a side length of —100 A. In the middle of the molecule there is a hole about 20 to 25 A in diameter. This hole arises as a natural consequence of packing roughly spherical large objects around a fourfold symmetry axis. There are extensive and strong interactions between the subunits that are related by the fourfold axis. In total, 22 direct hydrogen bonds including 8 salt-bridges, plus 10 waterbridges contribute to the stability of this interaction in which an accessible surface of 3110 A 2 , 21% of the subunits total accessible surface, is buried. The twofold contacts between the two tetramers forming the octamer are much less extensive with only 372 A2 buried per subunit. I. FMN BINDING The FMN molecule is bound to the barrel structure at the carboxy end of the strands. The funnel-shaped binding site is formed by the loops (henceforth numbered as the preceding strand) that join the parallel strands with the subsequent helices. All residues that form hydrogen bonds to the coenzyme are in these loop regions or at the carboxy terminal of the p-strands in the barrel structure. Figure 6A is a schematic drawing that illustrates the position of FMN in relation to the strands of the barrel and also highlights residues involved in FMN binding, and Figure 6B is a stereo view of bound FMN in the same orientation. Within the accuracy of the electron density map the isoalloxazine ring is flat. The ribityl side chain is in a fairly extended conformation (see Figure 7). The phosphate group is located at the amino end of the short helix H immediately after strand eight with the helix dipol moment compensating part of the negative charge of the phosphate. Ionic bonds are made to Arg-309 at the beginning of this helix and Arg-289 in loop seven. The phosphate also makes hydrogen bonds to main-chain nitrogen atoms in these two loops and to bound water molecules in contact with bulk solvent.

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FIGURE 6A. Schematic picture of the FMN binding site in glycolate oxidase looking down the barrel axis. The p-strands of the p/a barrel are shown as arrows and residues involved in polar interactions with FMN are shown as wire models.

FIGURE 6B. Stereo diagram of FMN bound to glycolate oxidase in the same view as Figure 6A, only the Cot-tracing is shown for the p-strands.

FIGURE 7.

The conformation^ angles of FMN.

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FIGURE 8.

Hydrogen bond network around FMN in glycolate oxidase.

The ribityl side chain is buried inside the barrel. It interacts mainly with the ends of strand one and seven. One ribityl oxygen atom forms a hydrogen bond to the main chain oxygen of Pro-77 at the end of strand one and a second oxygen to the side chain of Asp285 in strand seven. The isoalloxazine ring is sandwiched between side chains from loop six against its siface, and the end of strand one covering the re-face. The methyl groups are shielded from bulk solvent by tyrosine side chains from helix B. In addition the polar atoms of the flavin ring participate in an extensive network of hydrogen bonds both directly to polar protein atoms and indirectly through water molecules. This effectively buries the ring system to such an extent that the only part that is exposed to the solvent is the si-face around the N(5) edge of the ring. Figure 8 shows the primary and secondary partners in this complex network and Table 1 lists the polar contacts with FMN that are shorter than 3.3 A. Atoms N(l) and O(2) of the flavin ring are both within hydrogen bonding distance to Lys-320 in strand five. The positive charge of this lysine residue causes the stabilization of the negative charge at the N1-C2=O locus.29 O(2) also makes an hydrogen bond to Thr155 in strand four. This threonine side chain also interacts with N(3). O(4) is hydrogen bonded to Tyr-129 in strand three. This interaction is responsible for lowering the pKa of N(3) as has been shown through site-directed mutagenesis of this tyrosine to a phenylalanine. 28 A considerable number of the amino acids surrounding the flavin ring system are aromatic and the approximate angles between the normals of their side-chain rings and that of the isoalloxazine ring are 45° (Tyr-25), 45° (Tyr-129), 120° (Tyr-24) and 100° (Trp-108), respectively. J. THE SUBSTRATE POCKET Outside the reactive N(5) position of the isoalloxazine ring there is a pocket which is accessible to the solvent via a channel through the "lid" part of the structure. Residues lining this pocket in the vicinity of the cofactor are Trp-108 from strand two, Tyr-129 in loop three, Tyr-24 in helix B, His-254 and Arg-257 from the very long loop six, and further

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TABLE 1 Polar Contacts of FMN in Glycolate Oxidase. (Contacts Shorter Than 3.3 A are Included) Distance FMN atoms

Protein atoms

(A)

N(i) 0(2) 0(2) N(3) N(3) 0(4) 0(4) N(5) 0(2*) 0(1*) 0(3*) 0(3*) 0(3*) 0(3*) OP(1) OP(1) OP(2) OP(2) OP(2) OP(3) OP(3) OP(3)

K230 Nz K230 Nz T155 Og Water T155 OG Y129 OH Water Water K230 Nz Carbonyl 77 O D 285 Odl D 285 Od2 K 230 Nz S 252 Og Amide 309 N R309 NH Amide 308 N R289 NH Water R289 NH R289 NH Amide 287 N

3.0 2.9 2.7 3.3 3.2 2.8 2.7 3.0 3.3 2.7 2.7 3.3 3.1 3.2 2.8 3.3 2.8 3.2 2.5 3.1 2.8 2.8

Note: Asterisks denote that these atoms are part of the ribityl side chain as opposed to the isoalloxazine ring system and the phosphate.

FIGURE 9.

Stereo view of the active site in glycolate oxidase.

out in the channel Arg-164 (see Figure 9). A 5 A difference Fourier electron density map calculated for the substrate analogue thioglycolate, showed a peak in this area. In the native structure there are water molecules filling the space of this pocket close to the coenzyme. Two of these have an internal arrangement such that it mimics the structure of the carboxyl group of a bound glycolate molecule (cf. Figures 9 and 10). Glycolate has been model built into the substrate pocket. In this model the carboxyl group of glycolate makes an ionic bond to Arg-257 and a hydrogen bond to Tyr-24 while the hydroxyl group hydrogen bonds to Tyr-129. Atom O(2) in glycolate then comes close to atom N(5) of FMN and to His~254 (Figure 10).

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FIGURE 10. Model of glycolate (thick lines) in the active site of glycolate oxidase.

K. THE CATALYTIC MECHANISM Very few solution studies have been made on the catalytic mechanism of plant glycolate oxidase. On the other hand, spectroscopic and kinetic studies on other FMN dependent oxidases42'46 as well as flavocytochrome b2,47 have provided the following general hypothesis on the mechanism for this class of enzymes. In the first step in the substrate oxidative half reaction, after substrate binding, a proton is abstracted from the substrate C(2) carbon atom, producing a carbanion. This reaction in glycolate oxidase is stereospecific, only re-hydrogen of glycolate is removed.27 The carbanion can subsequently attach to the N(5) atom of FMN, perhaps transiently forming a covalent adduct which transfers two electrons to the isoalloxazine ring. The reduced FMN is then reoxidized by oxygen in the coenzyme oxidative halfreaction. It is not known where oxygen attacks the flavin ring in the oxidases but for free flavin48 and for flavin oxygenases49 it is known that 4a-hydroperoxide is formed during the reaction. This causes a considerable bending of the flavin ring. The structure of glycolate oxidase is consistent with this general mechanism and residues that are likely to participate in the various steps of the reaction can be identified. In the substrate binding pocket and in the vicinity of bound FMN there are a large number of amino acids with charged and polar side chains as well as several firmly bound water molecules. These form an extended hydrogen-bond network to which substrate and polar atoms of FMN are bound (see Figure 8). The model of bound glycolate is compatible with a carbanion mechanism. The C(2) position of glycolate is close to N(5) of FMN and in addition close to N(3) of the side chain of His-254. This side chain is in position to abstract the re-proton from C(2) of the substrate. The subsequent electron transfer to the flavin ring from the carbanion of the substrate is facilitated by the interaction between the positively charged Lys-230 and N(l) as well as O(2) of the flavin ring which enhances the oxidative propensity of the N(5) position and stabilizes the reduced FMN anion. A similar interaction occurs in FAD enzymes50 where the N1-O2 atoms interact with the positive dipole of an a-helix instead of a positively charged side chain. The formation of a transient covalent intermediate in the electron transfer step can be neither proved nor falsified on structural grounds. In the second part of the reaction, where reduced FMN is reoxidized by oxygen to form hydrogen peroxide, the structural model of glycolate oxidase suggests the transient formation of a 4a-hydroperoxide. Two enantiomers of this intermediate are possible. The bending of the flavin ring could be stabilized if the O(4) oxygen turned towards the re-face where there is a small pocket, in the native structure occupied by a water molecule (Figure 9). In this position O(4) would make new hydrogen-bonds to Ser-106 and Gin-127 and main chain oxygen of Thr-18, compensating for those lost on bending. The hydroperoxide part would then be oriented on the si-face towards His-254. However, the structure of 4a,5-epoxy-ethano-3-methyl-4a,5-dihydrolumiflavin51 (Figure 11), an analogue of the flavin-4a-hydroperoxide intermediate, is of the other enantiomer,

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FIGURE 11. (A) Flavin 4a-hydroperoxide. Op: proximal oxygen; Od: distal oxygen. (B) 4a,5-Epoxy-ethano-3-methyl-4a,5-dihydrolumiflavin, a model for flavin 4a-hydroperoxide.

FIGURE 12. Superposition of FMN in glycolate oxidase (thin line) with 4a,5-epoxy-ethano-3-methyl-4a,5-dihydro lumiflavin (thick line).

orienting what corresponds to the hydroperoxide part on the re-face. Superposition of this analogue on FMN in glycolate oxidase (Figure 12) shows that the hydroperoxide would fit excellently in the water pocket with the distal oxygen in hydrogen bond distance to mainchain atom O78, Flavin O(2) and Oy of Ser-106. This suggests that the water pocket close to the C(4)O positions of the isoalloxazine ring on the re-side (Figure 9) in the oxidized enzyme is the oxygen pocket of the reduced enzyme. The water molecule, which makes several hydrogen bonds, could be replaced by oxygen, which would be suitably located for receiving electrons from reduced FMN and forming a covalent bond at position O(4a). If this hypothesis is correct, the cofactor in glycolate oxidase would receive electrons from the substrate on the si side and deliver them to oxygen on the re-side. There cannot be any objection a priori to this transfer since no oxygen is incorporated into the product by glycolate oxidase.17 A corresponding structure for the 4a-hydroperoxide intermediate in /?-hydroxybenzoate hydroxylase has been suggested.52 A comparison to the homologous enzyme flavocytochrome b2, discussed in more detail below, supports this theory. Flavocytochrome b2 (see also Lederer, Chapter 7 of Vol. II of this series) is an electron transferase and is thus not reoxidized by oxygen and accordingly the suggested oxygen pocket corresponding to the internal water on the re-side in glycolate oxidase does not exist in this enzyme. Some inhibitor binding studies have been made to mammalian glycolate oxidase. The results should be applicable in a qualitative manner to the plant enzyme since they seem to have very similar properties. Based on the tight binding of oxalate, it has been proposed that there should be at least two positively charged residues in the active site and that these should also bind the substrate.18 There are indeed two such residues, Arg-164 and Arg-257, that can form a good binding site for oxalate. However, this site is too far from FMN to simulate a productive substrate binding site. It is possible however, that this represents only an initial binding site, and that oxalate then in a slower process, accompanied by proton uptake to His-254, is bound as a transition state analogue. Such a two-step binding process

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for oxalate has been demonstrated in the case of L-lactate oxidase53 a functionally related enzyme. Inhibitor studies with monocarboxylic acids of different chain lengths have shown that there must be a hydrophobic pocket big enough to accommodate a chain of at least 5 alkyl residues.18 From the structure it is seen that hydrophobic interactions, albeit not very extensive, could be made along the substrate channel which is sufficiently big to accommodate substrates of this length.

IIL COMPARISON WITH OTHER FMN-BINDING ENZYMES A. p/a-BARREL SIMILARITY BETWEEN TRIMETHYLAMINE DEHYDROGENASE AND GLYCOLATE OXIDASE The eight-stranded (3/a-barrel structure is a common structural motif that has been found in a variety of different enzymes. This motif occurs among enzymes of nonhomologous amino acid sequences and quite different functions. They are built up from a common core of eight parallel strands and eight helices connected by loop regions that vary considerably in length and conformation. When these structures of nonhomologous sequences are superimposed and compared, it is found54 that the common core comprises about 160 residues with an rms deviation of around 3.0 A in their Ca-atom positions. Trimethylamine dehydrogenase (EC 1.5.99.7) is a flavin enzyme whose structure has been determined55 (see Steenkamp and Mathews, Chapter 15 of Vol. II, this series). It is composed of three structural domains of which the largest, N-terminal domain, is a (3/otbarrel that binds FMN and an iron-sulfur cluster [4Fe-4S]. When the flavin-binding domain of trimethylamine dehydrogenase is compared to glycolate oxidase, the number of equivalenced Ca-atoms is 168 with an rms deviation of 2.6 A. This is slightly closer than what is obtained comparing glycolate oxidase and triosephosphate isomerase, 167 equivalences with an rms deviation of 2.9 A. In the latter case the similarity between the structures resides in the core of the p/a-structure, i.e., in the eight strands and helices, while the connecting loops are quite different. The similarities between trimethylamine dehydrogenase and glycolate oxidase lies also in the p/a-barrel core, but in addition, both structures have two short antiparallel strands closing off the barrel at the N-terminal end of the strands of the barrel. The FMN molecules are positioned in a similar way in both enzymes with an rms deviation of 1.6 A when the rotation matrix obtained by superimposing the Ca-atoms is used. In trimethylamine dehydrogenase the FMN molecule is covalently bound through its 6-position to a cysteine side chain of the protein. The phosphate groups are, in both enzymes, in approximately the same position as the substrate phosphate group in triosephosphate isomerase, at the N-terminal end of an a-helix in one of the loops that connects the strands with the helices. Thus the functional basis for FMN-binding involves the same folding framework but does not require extensive structural similarity between the barrel structures. No chemical sequence is known for trimethylamine dehydrogenase but with the rms deviation obtained (2.6 A) no significant sequence homology between these two enzymes is expected. The conformation of FMN when it is bound to these barrel structures is quite different from that observed in the electron carrier flavodoxin56-57 where the protein moiety has a different structure, an open twisted pleated sheet of parallel p-strands with helices on both sides. The ribityl side chain of FMN is more extended in glycolate oxidase and points in a different direction relative to the ring system. Thus different overall protein structures forces FMN to bind in different conformations, whereas proteins of the same structural type bind FMN in similar conformations. A similar situation has been observed for NAD binding to various proteins.58

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FIGURE 13. Superposition of the Ca-atoms of one subunit of flavocytochrome b2 (thin line) on glycolate oxidase (thick line).

B. STRUCTURAL COMPARISON OF GLYCOLATE OXIDASE AND FLAVOCYTOCHROME b2 1. Overall Similarities Flavocytochrome b2 from yeast (EC 1.1,2.3) is a tetrameric enzyme, carrying one FMN and one protoheme IX per subunit, that catalyzes the oxidation of lactate to pyruvate.59 In this process, FMN is first reduced in a reaction mechanism which is similar to that of ahydroxy acid oxidases42-60 including glycolate oxidase. However, in flavocytochrome b2, FMNH2 is reoxidized by transferring its electrons through the heme group to cytochrome c which is the natural electron acceptor. This sequence of steps involves the transient formation of a catalytically competent flavin semiquinone.61 The X-ray structure of flavocytochrome b2 from baker's yeast has been determined and refined62 (see Lederer, Chapter 27, Vol. II, this series). The N-terminal 99 residues form a separate heme-binding domain, the cytochrome b2 core. The following 365 residues are folded into an eight stranded p/a-barrel that binds FMN, preceded by a small helical domain, thus similar to the glycolate oxidase structure. Finally, a 25 residue extended tail makes contact with each of the other 3 subunits of the tetramer. The sequence of glycolate oxidase15'16 exhibits 37% identity to the sequence of the flavin-binding domain in flavocytochrome b2,63'64 thus indicating that the two proteins are homologous. Superimposing flavocytochrome b2 on glycolate oxidase gives 311 equivalent Ca-atoms with an rms deviation of 0.93 A. This implies that the three-dimensional structures are largely identical. Thus not only the common core, but also virtually all loop regions exhibit very similar conformations. Even the N-terminal regions (residues 1 to 70 in glycolate oxidase; 123 to 192 in flavocytochrome b2), which form part of the small additional helical domain outside the barrel, superimpose well (Figure 13). The only large significant difference is that the loop in glycolate oxidase between strand four and helix four of the barrel, involving

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29 residues which cover the active site, is moved away from the active site in flavocytochrome b2 and is replaced by the cytochrome domain. Other minor differences between the two structures are (1) this loop is five residues shorter in flavocytochrome b2; (2) helix six of the barrel has a slightly different tilt; and (3) three loops at the N-terminal end of the IBstrands (32, |33 and [37 in the barrel are two to five residues longer in flavocytochrome b2. These inserted residues in flavocytochrome b2 are not involved in any packing interactions, and the affected loops are in both structures facing the solvent and are of no functional significance. Flavocytochrome b2 has also a one residue insertion at the beginning of helix 1 and its extended tail of about 25 residues at the C-terminal end does not exist in glycolate oxidase. A sequence alignment based on the structural alignment is shown in Figure 14. A striking observation is that in spite of this strong similarity in the backbone structure, the FMN orientation and binding mode in the two structures, although similar, are significantly different (Figure 15). Superimposing the main-chain atoms of ten residues in the loops involved in FMN-binding or catalysis gives an rms deviation of 0.57 A. Using this rotation matrix, the rms deviation between the atoms of the bound FMN ring systems is 1.1 A and 0.54 A for the ribityl and phosphate atoms. The packing of the subunits of the two enzymes has also been compared. The fourfold packing arrangement, employing a crystallographic axis in the case of glycolate oxidase and a local axis in flavocytochrome b2, is very similar in the two enzymes. Superposition of half the tetramer of flavocytochrome b2 on glycolate oxidase gave an rms deviation of 0.98 A for 616 atoms. There are very tight subunit contacts made around the fourfold axis in flavocytochrome b2 by the tail region which does not exist in glycolate oxidase. The subunit interactions in glycolate oxidase about the twofold axes normal to the fourfold axis, which form the octameric structure, are not conserved in flavocytochrome b2. 2. Active-Site Geometry and Substrate Binding The substrates for glycolate oxidase and flavocytochrome b2 are very similar, both being small a-hydroxy acids, glycolate and lactate, respectively. The structure of the complex of pyruvate, the product of lactate oxidation, bound to flavocytochrome b2 in the semiquinone form has been determined. Using the superposition of the Cot-atoms, the glycolate model in glycolate oxidase falls on top of pyruvate in flavocytochrome b2. There is only one side chain in the active site which is not conserved, Trp-108 in glycolate oxidase which is Leu-230 in flavocytochrome b2. Part of the volume of the tryptophan ring is occupied by the propionate side chain of the heme in flavocytochrome b2. In loop 1 close to the active site, Pro-77 in glycolate oxidase is Ala-196 in flavocytochrome b2 but the main chain conformations of these residues are the same. One important difference in FMN binding is a hydrogen bond from the side chain of Ser-195 in (3-strand 1 to the ribityl side chain in flavocytochrome b2, which is not present in glycolate oxidase where the corresponding residue is Ala-76. This difference in hydrogen bonding contributes to the ease with which FMN can be abstracted from glycolate oxidase by raising the salt concentration as compared to flavocytochrome b2. A second difference in loop 1 (Figure 16), is the main chain conformations of Thr-78/ 197 which has an energetically unfavorable Rvalue in the case of glycolate oxidase. The side chain conformations of these threonines are also different in the two structures. This might be caused by Leu-436 in flavocytochrome b2, which corresponds to Val-312 in glycolate oxidase. This residue could restrain the position of the methyl group of Thr-197 in flavocytochrome b2. The only other difference in the vicinity of this Thr is Gln-81 in glycolate oxidase which corresponds to Cys-200 in flavocytochrome b2. The third major difference in the active site is the buried water molecule, close to the C(H)O positions of the isoalloxazine ring (Figures 15,16) in glycolate oxidase, which is not

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FIGURE 14. Sequence alignment of flavocytochrome b2 and glycolate oxidase based on the structural alignment. The placement of a deletion between flavocytochrome b2 positions 313 and 314 is arbitrary since there is no sequence or structural similarity between positions 300 to 324.

present in flavocytochrome b2. In the latter, the FMN ring system lies closer to strand one and the amide of Ala-198 forms a hydrogen bond to the flavin N(5). The different locations of the flavin rings are also reflected in a slight but pronounced perturbation in the same direction of some of the side chains (mainly, Tyr-129/254, His-254/373, and Lys-230/349) around the FMN, resulting in a slightly different hydrogen-bond network in the active site.

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FIGURE 15. Stereo diagram of the superimposed active sites of glycolate oxidase (thick line) and flavocytochrome b2 (thin line). The cross behind the flavin is a buried water molecule in glycolate oxidase which is not present in flavocytochrome b2.

FIGURE 16.

Stereo diagram of FMN and loop 1 in (A) glycolate oxidase; (B) flavocytochrome b2.

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3. Mechanistic Implications The similarities at the active sites support the idea of an identical first step, carbanion formation, for the chemical reaction catalyzed by the two enzymes. There is no evidence as to the nature of the second step, electron transfer proper, and whether it is identical or not in the two classes of enzymes (electron transferase vs. oxidase). The flavin oxidative half-reaction is clearly very different in the two enzymes. In flavocytochrome b2, solvent and hence oxygen access to the flavin re-side is prohibited by the hydrogen bonds from the side chains of Gln-252 and Ser-228 to FMN N(3) and O(4), respectively, and from the main-chain nitrogen to Ala-198 to N(5). The difference in FMN orientation in the two structures is thus the cause of the functional difference between the enzymes in the flavin oxidative half reaction. The emergence of the flavooxidase and flavodehydrogenase function seem to have proceeded by divergent evolution from a common precursor. During this evolutionary process, the occurrence of a few mutations, fine-tuning the FMN orientation, have had a decisive influence on determining the ultimate function of the enzymes.

REFERENCES 1. Miziorko, H. M. and Lorimer, G., Ribulose-l,5-bisphosphate carboxylase-oxygenase, Ann. Rev. Biochem., 52, 507, 1983. 2. Frigerio, N. A. and Harbury, H. A., Preparation and some properties of crystalline glycolic acid oxidase of spinach, J. Biol Chem., 231, 135, 1958. 3. Kerr, M. W. and Groves, D., Purification and properties of glycolate oxidase hompisum sativum leaves, Phytochemistry, 14, 359, 1975. 4. Nishimura, M., Akhmedov, Y. D., Strzalka, K., and Akazawa, T., Purification and characterization of glycolate oxidase from pumpkin cotyledons, Arch. Biochem. Biophys., 222, 397, 1983. 5. Behrends, W., Rausch, U., Loeffler, H.-G., and Kindl, H., Purification of glycolate oxidase from greening cucumber cotyledons, Planta, 156, 566, 1982. 6. Pandey, O. P. and Sanwal, G. G., Purification and properties of glycolate oxidase fromNopalea dejecta, a CAM plant, Plant Physiol. Biochem., 9, 80, 1982. 7. Schuman, M. and Massey, V., Purification and characterization of glycolic acid oxidase from pig liver, Biochim. Biophys. Acta, 221, 500, 1971. 8. Duley, J. A. and Holmes, R. S., L-a-Hydroxyacid oxidase isozymes purification and molecular properties, Eur. J. Biochem., 63, 163, 1976. 9. Dupuis, L., DeCaro, J., Brachet, P., and Puigserver, A., Purification and some characteristics of chicken liver L-2-hydroxyacid oxidase A, FEBS Lett., 266, 183, 1990. 10. Schwam, H., Michelson, S., Randall, W. C., Sondey, J. M., and Hirschman, R., Purification and characterization of human liver glycolate oxidase. Molecular weight, subunit and kinetic properties, Biochemistry, 18, 2828, 1979. 11. Fry, D. W. and Richardson, K. E., Isolation and characterization of glycolic acid oxidase from human liver, Biochim. Biophys. Acta, 568, 135, 1979. 12. Liao, L. L. and Richardson, K. E., The inhibition of oxalate biosynthesis in isolated perfused rat liver by DL-phenyl lactate and n-heptanoate, Arch. Biochem. Biophys., 154, 68, 1973. 13. Williams, H. E. and Smith, L. A., Jr., Disorders of oxalate metabolism, Amer. J. Med., 45, 715, 1968. 14. Zelitch, I. and Ochoa, S., Oxidation and reduction of glycolic and glyoxylic acids in plants. I. Glycolic acid oxidase, J. Biol. Chem., 201, 707, 1953. 15. Cederlund, E., Lindqvist, Y., Soderlund, G., Branden, C.-L, and Jornvall, H., Primary structure of glycolate oxidase from spinach, Eur. J. Biochem., 173, 523, 1988. 16. Volokita, M. and Somerville, C. R,, The primary structure of spinach glycolate oxidase deduced from the DNA sequence of a cDNA clone, J. Biol. Chem., 262, 15825, 1987. 17. Richardson, K. E. and Tolbert, N. E., Oxidation of glyoxylic acid to oxalic acid by glycolic acid oxidase, J. Biol. Chem., 236, 1280, 1961. 18. Schuman, M. and Massey, V., Effect of anions on the catalytic activity of pig liver glycolic acid oxidase, Biochim. Biophys. Acta, 227, 521, 1971. 19. Zelitch, L, a-Hydroxysulfonates as inhibitors of the enzymatic oxidation of glycolic and lactic acids, J. Biol. Chem., 224,251, 1957.

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20. Corbett, J. R. and Wright, B. J., Inhibition of glycolate oxidase as a rational way of designing a herbicide, Phytochemistry, 10, 2015, 1971. 21. Jewess, P. J., Kerr, M. W., and Whitaker, D. P., Inhibition of glycolate oxidase from pea leaves, FEBS Lett., 53, 292, 1975. 22. Schonbrunn, A., Abeles, R. H., Walsh, C. T., Ghisla, S., Ogata, H., and Massey, V., The structure of the covalent flavin adduct formed between L-lactate oxidase and the suicide substrate 2-hydroxy-3butynoate, Biochemistry, 15, 1798, 1976. 23. Randall, W. C., Streeter, K. B., Cresson, E. L., Schwam, H., Michelson, S. R., Anderson, P. S., Cragoe, E. J., Jr., Williams, W. R., Eichler, E., and Rooney, C. S., Quantitative structure-activity relationships involving the inhibition of glycolic acid oxidase by derivatives of glycolic and glyoxylic acids, J. Med. Chem., 22, 608, 1979. 24. Rooney, C. S. , Randall, W. C., Streeter, K. B., Ziegler, C., Cragoe, E. J., Jr., Schwam, H., Michelson, S. R., Williams, W. R., and Eichler, E., Inhibition of glycolic acid oxidase. 4-Substituted 3-hydroxy-lH-pyrrole-2,5-dione derivatives, J, Med. Chem., 26, 700, 1983. 25. Williams, W. R., Eichler, E., Randall, W. C., Rooney, C. S., Cragoe, E. J., Jr., Streeter, K. B., Schwam, H., Michelson, S. R., Patchett, A. A., and Taub, D., Inhibition of glycolic acid oxidase. 4Substituted-2.4-dioxobutanoic acid derivatives, J. Med. Chem., 26, 1196, 1983. 26. Massey, V., Miiller, F., Feldberg, R., Schuman, M., Sullivan, P. A., Howell, L. G., Mayhew, S. G., Matthews, R. G., and Foust, G. P., The reactivity of flavoproteins with sulfite. Possible relevance to the problem of oxygen reactivity, /. BioL Chem., 244, 3999, 1969. 27. Fendrich, G. and Ghisla, S., Studies on glycolate oxidase from pea leaves: determination of stereospecificity and mode of inhibition by a-hydroxybutynoate, Biochim. Biophys. Acta, 702, 242, 1982. 28. Macheroux, P., Massey, V., Thiele, D. J., Soderlind, E., and Lindqvist, Y., Characterization of glycolate oxidase and an active site mutant, in Flavins and Flavoproteins, Curti, B., Ronchi, S., and Zanetti, G., Eds., W. de Gruyter, Berlin, 1991, 119. 29. Massey, V., Ghisla, S., and Moore, E. G., 8-Mercaptoflavins as active site probes of flavoenzymes, /. Biol. Chem., 254, 9640, 1979. 30. Pace, C. and Stankovich, M., Oxidation-reduction properties of glycolate oxidase, Biochemistry, 25, 2516, 1986. 31. Lindqvist, Y. and Branden, C.-I., Preliminary crystallographic data for glycolate oxidase from spinach, J. Biol. Chem., 254, 7403, 1979. 32. Lindqvist, Y. and Branden, C.-L, Structure of glycolate oxidase from spinach at a resolution of 5.5 A, J. MoL BioL, 143, 201, 1980. 33. Lindqvist, Y. and Branden, C.-I., Structure of glycolate oxidase from spinach, Proc. Natl. Acad. Sci. U.S.A., 82, 6855, 1985. 34. Lindqvist, Y. and Branden, C.-L, The active site of spinach glycolate oxidase, J. BioL Chem., 264, 3624, 1989. 35. Lindqvist, Y., Refined structure of spinach glycolate oxidase at 2 A resolution, /. MoL BioL, 209, 151, 1989. 36. Lindqvist, Y., Branden, C.-L, Mathews, F. S., and Lederer, F., Spinach glycolate oxidase and yeast flavocytochrome b2 are structurally homologous and evolutionary related enzymes with distinctly different function ana FMN-binding, J. BioL Chem., 266, 3198, 1991. 37. Luzatti, V., Traitement statistique des erreurs dans la determination des structures cristallines, Acta Cryst., 5, 802, 1952. 38. Phillips, D. C., Sternberg, M. J. E., Thornton, J. M., and Wilson, L A., An analysis of the structure of triose phosphate isomerase and its comparison with lactate dehydrogenase, /. MoL BioL, 119, 329, 1978. 39. McLachlan, A. DM Gene duplication in the structural evolution of chymotrypsin, J. MoL BioL, 128, 49, 1979. 40. Kabsch, W. and Sander, C., Dictionary of protein secondary structure: pattern recognition of hydrogen bonded and geometrical features, Biopolymers, 22, 2577, 1983. 41. Miller, S., Janin, J., Lesk, A. M., and Chotia, C., Interior and surface of monomeric proteins, J. MoL BioL, 196, 641, 1987. 42. Ghisla, S., Dehydrogenation mechanisms in flavoprotein catalysis, in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Eds., Elsevier/North Holland, Amsterdam, 1982, 133. 43. Walsh, C. T., Flavin coenzymes: At the crossroads of biological redox chemistry, Ace. Chem. Res., 13, 148, 1980. 44. Bruice, T. C., Mechanisms of flavin catalysis, Ace. Chem. Res., 13, 256, 1980. 45. Ghisla, S. and Massey, V., Mechanisms of flavoprotein-catalyzed reactions, Eur. J. Biochem., 181, 1, 1989. 46. Ghisla, S. and Massey, V., L-lactate oxidase, in Chemistry and Biochemistry of Flavoenzymes, Vol. 2, Muller, F., Ed., CRC Press, 1991, 243.

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47. Lederer, F., Flavocytochrome b2, in Chemistry and Biochemistry of Flavoenzymes, Vol. 2, Miiller, F., Ed., CRC Press, 1991, 153. 48. Bruice, T. C. ? Flavin oxygen chemistry brought to date, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 45. 49. Ballou, D. P., Flavoprotein monooxygenases, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 605. 50. Wierenga, R., Drenth, J., and Schulz, G. E., Comparison of the three-dimensional protein and nucleotide structure of the FAD-binding domain of p-hydroxybenzoate hydroxylase with the FAD- as well as NADPHbinding domains of glutathione reductase, J. Mol. BioL, 167, 725, 1983. 51. Bolognesi, M., Ghisla, S., and Incoccia, L., The crystal and molecular structure of two models of catalytic flavo (co) enzyme intermediates, Acta Cryst., B34, 821, 1978. 52. Schreuder, H. A., Hoi., W. G. J., and Drenth, J., Analysis of the active site of the flavoprotein phydroxybenzoate hydroxylase and some ideas with respect to its reaction mechanism, Biochemistry, 29, 3101, 1990. 53. Ghisla, S. and Massey, V., Studies on the mechanism of action of the flavoenzyme lactate oxidase. Proton uptake and release during the binding of transition state analogs, J. BioL Chem., 252, 6729, 1977. 54. Lebioda, L., Hatada, M. H., Tulinsky, A., and Mavridis, I. M., Comparison of the folding of 2-keto3-deoxy-6-phosphogluconate aldolase, triosephosphate isomerase and pyruvate kinase. Implications in molecular evolution, J. Mol. BioL, 162, 445, 1982. 55. Lim, L. W., Shamala, N., Mathews, S. F., Steenkamp, D. J., Hamlin, R., and Xuong, N., Threedimensional structure of the iron-sulfur flavoprotein trimethylamine dehydrogenase at 2.4 A resolution, J. BioL Chem., 262, 15140, 1986. 56. Watenpaugh, K. D., Sicker, L. C., and Jensen, L. H., The binding of riboflavin-5'-phosphate in a flavoprotein: flavodoxin at 2.0 A resolution, Proc. Natl. Acad. Sci. U.S.A., 70, 3857, 1973. 57. Burnett, M. R., Darling, G. D., Kendall, D. S., LeQuesne, M. E., Mayhew, S. G., Smith, W. W., and Ludwig, M. L., The structure of the oxidized form of clostridial flavodoxin at 1.9 A resolution, J. BioL Chem., 249,4383, 1974. 58. Eklund, H. and Branden, C.-L, Crystal structure, coenzyme conformation and protein interactions, in Pyridine Nucleotide Coenzymes Chemical, Biochemical, and Medical Aspects, Vol. 2, Dolphin, D., Poulson, R., and Avramovic, O., Eds., John Wiley & Sons, New York, 1987, 51. 59. Jacq, C. and Lederer, F., Cytochrome b2 from baker's yeast (L-lactate dehydrogenase): A double headed enzyme, Eur. J. Biochem., 41, 311, 1974. 60. Urban, P. and Lederer, F., Intermolecular hydrogen transfer catalyzed by a flavodehydrogenase, baker's yeast flavocytochrome b2, J. BioL Chem., 260, 11115, 1985. 61. Capeillere-BIandin, C., Bray, R. C., Iwatsubo, M., and Labeyrie, F., Flavocytochrome b2: kinetic studies by absorbance and electron-paramagnetic-resonance spectroscopy of electron distribution among prosthetic groups, Eur. J. Biochem., 54, 549, 1975. 62. Xia, Z.-X. and Mathews, F. S., Molecular structure of flavocytochrome b2 at 2.4 A resolution, J. Mol. BioL, 212, 837, 1990. 63. Lederer, F., Cortial, S., Becam, A. M., Haumont, P. YM and Perez, L., Complete amino acid sequence of flavocytochrome b2 from baker's yeast, Eur. J. Biochem., 139, 59, 1985. 64. Guiard, B., Structure, expression and regulation of a nuclear gene encoding a mitochondrial protein: the yeast L(-I-)-lactate cytochrome c oxidoreductase (cytochrome b2), EMBO J.t 4, 3265, 1985.

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Chapter 14 GENERAL PROPERTIES OF FLAVODOXINS Stephen G. Mayhew and Gordon Tollin

TABLE OF CONTENTS I.

Introduction

390

II.

Distribution and Functions

390

III.

Purification, Molecular Weight, Chemical Composition

395

IV.

Structures

396

V.

Oxidation-Reduction Properties

398

VI.

Interaction of Flavin and Protein

401

VII.

Electron Transfer Mechanisms A. Kinetic Evidence That the Exposed Dimethylbenzene Ring of FMN is Involved in Electron Transfer B. Evidence for Electrostatic Modulation of the Redox Kinetics C. Relation Between Rate Constants for Electron Transfer and Redox Potentials D. Static Effects on Rates of Electron Transfer E. Structural Aspects of Intermediate Protein-Protein Complexes F. Specificity of Protein-Protein Electron Transfer Reactions

406

References

407 408 410 411 415 417 417

390

Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION The flavodoxins are a group of small flavoproteins which contain a single molecule of FMN and function as electron carriers in low potential oxidation-reduction reactions. The first member of the group was originally termed "phytoflavin" and it was isolated in 1963 from Anacystis nidulans (Synechococcus PCC 6301). K2 Proteins with similar properties were subsequently discovered in other microorganisms, and by the time of the first detailed review of the field in 1975,3 the crystal structures of flavodoxins from Clostridium beijerinckii MP (formerly Clostridium MP)4 and Desulfovibrio vulgaris had been determined. In addition, several protein sequences were available, and extensive studies had been carried out on the redox and catalytic properties of native flavodoxins and on the flavin-binding properties of apoflavodoxins. Since then, additional flavodoxins have been isolated and characterized, the crystal structures of two more flavodoxins (A. nidulans and Chondrus crispus) have been determined, flavodoxin structures have been probed by a variety of NMR techniques, extensive studies have been carried out on the mechanisms of electron transfer to and from flavodoxins, and the techniques of molecular biology have begun to be used to study flavodoxin structure and function. The flavodoxins have been studied intensively because of their small size, their high stability, and the ease with which they can be isolated from several microbial sources and as recombinant proteins expressed by cloned genes, and also because of the notion that they may be models for larger flavoproteins, especially those flavoproteins which like flavodoxins stabilize the flavin semiquinone in its neutral form and have a low reactivity with oxygen. This chapter gives a general review of work on this group of flavoproteins with an emphasis on recent developments; further chapters in the present series (see Chapter 15, this Volume) provide detailed reviews of the X-ray crystallographic and NMR studies of flavodoxins. The reader is also referred to earlier reviews, which deal with various aspects of the subject.3-5"8

II. DISTRIBUTION AND FUNCTIONS Flavodoxins have been isolated from a range of strictly anaerobic, facultatively anaerobic, obligately aerobic, and photosynthetic bacteria, from blue-green algae, and from the eukaryotes, Chlorella fusca, a green alga, and the seaweed Chondrus crispus, a red alga (Table 1). They have not been reported in higher plants and animals. Their synthesis in most of the organisms of Table 1 is known to be regulated by iron; in such organisms, flavodoxin is either synthesized only during growth in iron-poor medium, or synthesis is enhanced by iron-deficiency. This regulation by iron, and the reciprocal regulation of the synthesis of the iron-sulfur electron-transfer protein, ferredoxin, has been discussed elsewhere.3 Flavodoxins are constitutive in Escherichia coli and Chondrus crispus, and their synthesis also appears to be independent of iron in the nitrogen-fixing bacteria Azotobacter vinelandii, Azotobacter chroococcum and Klebsiella pneumoniae; in contrast, flavodoxin synthesis is induced by iron-deficiency in nitrogen-fixing organisms that are strict anaerobes and/or which carry out photosynthesis. Flavodoxin is synthesized by sulfate-reducing bacteria of the genus Desulfovibrio even when the organisms are grown in an iron-rich medium, but because iron is precipitated from the growth medium as iron sulfide, it was not possible to decide whether flavodoxin synthesis by these organisms is controlled by iron.58 However, recent work using an antibody preparation to flavodoxin from D. vulgaris (Hildenborough) has shown that flavodoxin synthesis in this sulfate-reducing bacterium does indeed respond to iron in the culture medium.59 Very little is known about the genetic mechanisms that organisms use to regulate flavodoxin and ferredoxin synthesis in response to the iron concentration in the growth medium. Laudenbach et al.60 have shown that when iron is added

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to cultures of iron-deficient A. nidulans R2, the concentration of RNA transcripts that encode flavodoxin is rapidly reduced to a very low level, suggesting that control of flavodoxin synthesis by iron occurs at the level of transcription. Flavodoxins function as electron carriers between other redox proteins. Most of the reactions in which they have been found to function operate at physiological redox potentials similar to that of the hydrogen electrode (Em7 = — 420 mV). The redox properties of purified flavodoxins have suggested that the oxidized flavin is not directly involved in such reactions; rather the flavin in the protein cycles between the semiquinone and the hydroquinone, transferring single electrons to and from other proteins. However, there is evidence that flavodoxin semiquinone is the electron donor for the activation of methionine synthetase61-62 and the semiquinone is known to be oxidized readily to the quinone by a number of nonphysiological oxidants; therefore a wider physiological role for the semiquinone as an electron donor cannot be excluded. The activities of flavodoxins in reactions involved in the metabolism of pyruvate, hydrogen, sulfite, nicotinamide nucleotides, and dinitrogen have been discussed elsewhere.3'5 Recent work has extended the earlier observations to other organisms. In many cases, a flavodoxin from a particular microorganism functions interchangeably with a ferredoxin from the same organism in reactions catalyzed by cell-free extracts, although the kinetics of the reactions with the two electron carriers are usually different. However, unique roles have been ascribed to flavodoxins in some organisms. For example, flavodoxin has been implicated63 as the electron donor for trithionate reductase from D. vulgaris (Hildenborough) which catalyzes the reaction.

and in a different strain of D. vulgaris (Miyazaki F) flavodoxin is thought to be the natural electron acceptor for a pyruvate dehydrogenase that catalyzes the oxidation of pyruvate to acetyl-CoA.64 The most convincing demonstration of a specific role for flavodoxin has been made, however, with the nitrogen-fixing system of the faculative anaerobe, K. pneumoniae. A wealth of biochemical and genetic work has established flavodoxin as the sole physiological electron donor for nitrogenase in this organism.65"67 First, flavodoxin is synthesized only during growth under nitrogen-fixing conditions; second, the gene for flavodoxin (n//F) forms part of the m/gene cluster; third, mutant strains impaired in niJF do not fix nitrogen in vivo, although extracts from them contain high levels of nitrogenase and they reduce dinitrogen when dithionite ion is used as the nonphysiological source of electrons to by-pass the requirement for an electron carrier; finally, a pyruvate-flavodoxin oxido-reductase, the nifl gene product, has been purified from extracts and shown to regenerate a nitrogen-fixing system with flavodoxin and the two protein components of nitrogenase.

In contrast to the well-documented role for flavodoxin in K. pneumoniae, the role of flavodoxins in the aerobic nitrogen-fixing bacteria of the group Azotobacter is less clear. Fully-reduced flavodoxins isolated from these organisms serve as electron donors to purified nitrogenase, and flavodoxin synthesis by Az. chroococcum is repressed by ammonium ion.31 Klugkist et al.36 detected three flavodoxins in Az. vinelandii (ATCC 478). One of them

ATCC 478 Protein 1 Protein 2 Protein 3 K. pneumoniae R. rubrwn

— 22,800

21,500 20,500 21,500

— — —

274 274 274 — 272(54.2)

— — — —

378 372 371 370(9.6) 376(11.3)

— —

— 370(6.6) 370(9) 371(9.5)

— 274(50)

E. coli A, chroococcum A. vinelandii OP

— 14,500 —

458(11.6) 452(11.3) 461(10.6) 454(10.1) 460(11.2)

— 467(8.25) 452(10) 450(10.6) — —

456.5(10.2) 456(11.26) 456.5(10.7)

374(8.2) 374(10.05) 375.5(8.7) 273(47) 272(48.88) 273(48)

16,000 16,500 16,300

456(10.03)

374(8.46)

274(45.8)

15,400

444(7.7) 445(10.13) 443(9.1) 443(10.4) 446 445(10.2)

373(6.5) 377(9.22) 372(7.9) 374(8.47) 372 377(8.75)

Desulfovibrio desulfuricans BerreEau D. gigas D. salexigens D. vulgaris

272(38.6) 272(53.6) 272(40) 272(45.8) 270 272(47.6)

14,000 20,100 14,600

445(10.4)

376(9.1)

— 15,000

272(46.8)

15,800

Absorption maxima (e) nm (mA/-! cm'1)

C. tyrobutyricum Megasphaera elsdenii (Peptostreptococcus elsdenii}

Clostridium beijerinckii MP (Clostridium MP) C. formicoaceticum C. kluyveri C. pasteurianum

Source

MWof holoprotein

TABLE 1 Properties of Flavodoxins

7 7 7 7 —

— — 7.8 7 7.7 8.5 8.2 7.7 —

-320 -500 -506 -412 —

— -440 -435 -410 -518 -495 -464 -515

—

—

— — — -158 —

— — -150 -143 -240 -198 + 50 -270 -215

—

36 36 36 31 37,38

24 25 24,26,27 28,143 29,79 30,31 32,33 34 35

20 21 22 23

— — —

-390 -405 -395 7.05 8 7.4 —

— -372

— -419

10 11,12 13—15 9 16 17—19

— -130 — -132 — -115

— -407

— 7 — 7 — 7

9

-92

-399 7

Ref.

E2 (mV) Ei (mV)

pH

Redox potentials

22,000 20,000 — 21,000 22,000

Nostoc MAC Anabaena PCC 7119 Anabaena PCC 7120 Chondrus crispus Chlorella fusca

377

270 — 274(47.5) — — 275(50.9) 275(54.6) — 376(9.02) 377(8.4) 373(8.5) 386(10.7) 379(9)

— 377

— 275

465(10.2) — 466(9.5) 465(9.4) 464(9.2) 464(10.7) 464(10.2)

— 465(9.2)

—

7 7 —

7 7 8 — 7

-450 -447 -450 — -414 -414 -390 -425 — — — —

-50 -221 -281 — -215 -210 -195 -196

39,40 41,42 41,42 43 44 45, 46 47,48 49 50 51

Note: Flavodoxins have also been reported in Thiocapsa roseopersicina,52 Rhodopseudomonas capsulata,53 Synechocystis PCC 6714 (Aphanocapsa PCC 6714)54 Ankistrodesmus braunii,55 Micrococcus aeruginosa,56 and Fusobacterium nucleatum.57

17,000 20,300

Synechococcus lividus Synechoccus PCC 6301 (Anacystis nidulans)

394

Chemistry and Biochemistry of Flavoenzymes

(flavodoxin 3, Table 1) was present only when the organism was grown with ammonium ion as nitrogen source; the other two were also found in cells grown under N 2 -fixing conditions, but the synthesis of one (flavodoxin 2, Table 1) was enhanced tenfold by growth with dinitrogen. It was concluded that flavodoxin 2 is probably a m/gene product in this organism. The three purified flavodoxins from this strain of Az. vinelandii differ from each other in their molecular properties (see later); it is not clear whether they can be distinguished by their catalytic activities with nitrogenase. The nifF gene that codes for flavodoxin in a different strain of Az. vinelandii (OP) has been cloned and sequenced.68 Mutants of the organism that lack this flavodoxin still fix dinitrogen, showing that flavodoxin is not the only electron carrier to nitrogenase. The alternative electron carrier has not been identified, and, in contrast to the ATCC 478 strain,36 there is no evidence for more than one type of flavodoxin in the OP strain.68 The source of electrons for the reduction of flavodoxins in Azotobacter spp. is also uncertain. Haaker and co-workers69"71 have shown that a high proton motive force across the cell membrane is required for nitrogenase activity in Az. vinelandii, and they have detected a linear relationship between the rate of electron transfer through the respiratory chain and nitrogenase activity in whole cells. They interpret these and other observations as evidence that the reducing equivalents for nitrogenase are generated in the cytoplasmic membrane, and that they are transferred to flavodoxin via an NAD(P)H:flavodoxin oxidoreductase in the membrane; dinitrogen was found to induce the synthesis of an NADPH dehydrogenase,71 but it is not known whether this enzyme functions in nitrogen fixation. The redox potential of the nicotinamide nucleotides at pH 7 ( — 0.32 V) is not sufficiently negative to allow reduction by NAD(P)H of flavodoxin to the hydroquinone, the redox form required for electron transfer to nitrogenase. It was argued earlier that because the effects of pH on the potentials of NAD(P)H and flavodoxins are different, reduction could occur at about pH 5, a condition that might be attained on the membrane surface.72 Reduction of flavodoxin by NAD(P)H is catalyzed by extracts and purified enzymes from a variety of organisms, and reduction under anaerobic conditions proceeds to give a mixture of flavodoxin semiquinone and hydroquinone according to the redox potentials of the nicotinamide nucleotide and the particular flavodoxin. With extracts of Clostridium tyrobutyricum, however, the extent of reduction is influenced by acetyl-CoA.16 In the absence of acetyl-CoA, flavodoxin is reduced only as far as the semiquinone; in the presence of an acetyl-CoA-regenerating system, complete reduction to the hydroquinone occurs. The explanation of this observation is not known. The effect is possibly related to an earlier observation, also unexplained, that an acetyl-CoA-regenerating system allows the formation of hydrogen from NAD(P)H by extracts of C. kluyveri.73'75 The observation of full reduction of flavodoxin by NADH and extracts of C. tyrobutyricum could result from complexation between acetyl-CoA and flavodoxin, and a consequent modification of the redox potentials of the electron carrier. However, spectroscopic evidence for the formation of such a complex could not be obtained with either C. tyrobutyricum or M. elsdenii flavodoxins, and acetylCoA does not have an appreciable effect on the redox potentials of these proteins.76 Flavodoxins are active in a number of reactions in which their precise roles are unclear. For example, the exchange of [1-14C-] of acetyl-CoA into CO by CO dehydrogenase from Clostridium thermoaceticum is stimulated by flavodoxin from C. formicoaceticum71 Flavodoxin functions as an electron acceptor for CO dehydrogenases78 and is converted to the semiquinone form.10 It has been proposed that in the exchange reaction the electron carrier plays a role in internal electron transfer during the cleavage of the methyl and CoA bonds to the carbonyl of acetyl-CoA, and the resynthesis of acetyl-CoA. In Escherichia coli, flavodoxin is required for the reductive activation of pyruvateformate lyase79'80 and of methionine synthetase.61-62 The activation of pyruvate-formate lyase requires fully reduced flavodoxin generated either by the oxidation of pyruvate, catalyzed

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by pyruvate:flavodoxin(ferredoxin) oxidoreductase (CoA-acylating), or from NADPH and an NADPH-flavodoxin oxidoreductase. The mechanism of the activation by flavodoxin is not understood, although the requirement for flavodoxin in the activation appears to be specific; E. coli ferredoxin is reduced by these systems but it does not activate the pyruvateformate lyase. In contrast, the activation of methionine synthetase is known to involve methylation of coenzyme B12 in the enzyme by adenosylmethionine;61'62 it appears that flavodoxin semiquinone is the electron donor for this reaction. Yoshino81 reported that Az. vinelandii flavodoxin is a potent allosteric inhibitor of AMP nucleosidase isolated from the same organism (Kj approximately 10 |xAf); FMN was found to be a weaker inhibitor (Ki = 4.7 mM) while apoflavodoxin had no effect. It was proposed that through its inhibitory effects on AMP nucleosidase, flavodoxin could be involved in the regulation of riboflavin biosynthesis in this organism.

III. PURIFICATION, MOLECULAR WEIGHT, CHEMICAL COMPOSITION Flavodoxins are acidic proteins (pi values of 3.12 to 5.05 have been reported8'82) that bind strongly to anion exchange resins such as diethylaminoethyl cellulose. Most of the procedures that have been described for their purification exploit this property (see References, Table 1). The procedures usually involve chromatography on DEAE-cellulose, salt fractionation with ammonium sulfate, and gel filtration; use has also been made of hydroxyapatite23 and of hydrophobic chromatography on DEAE-cellulose83 or Toyopearl.84 The strong hydrophobic interaction with DEAE-cellulose is especially useful for recovering flavodoxin from dilute solutions that contain high concentrations of ammonium sulfate; the protein is quantitatively adsorbed to the resin when a small amount of DEAE-cellulose is added, and it can be recovered by elution with sodium chloride.76 Most of the enzyme assays that have been described for flavodoxins are of limited use to measure purification because they lack specificity.3-85-86 Enzyme assays that involve reactions with nitrogenases30'87'88 or which assay the activation of E. coli enzymes61 '62-79-80 appear to be more specific, but they are not suitable for routine use because they require purified enzymes that are unstable. It is often more convenient to estimate purity from the optical spectrum, measuring the ratio of absorbance at the visible maximum to the absorbance at about 275 nm due to the protein (Table I). 3 Purified flavodoxins are stable for long periods in solution at 4°C or -20°C. Several of them crystallize readily and produce crystals suitable for X-ray diffraction analysis; in some cases, better crystals result when the reduced protein is used. One species of Az. vinelandii flavodoxin dimerizes during storage.89 Since dimerization is prevented by addition of a thiol such as mercaptoethanol, it is thought that the dimer involves the formation of a disulfide from the single cysteine sulfydryl group in the protein. The flavodoxins are usually monomeric. Their molecular weights are in the range 14,000 to 23,000, and according to their size, they fall into two groups, one in the range 14,000 to 17,000 and the other in the range 20,000 to 23,000 (Table 1). In contrast to other clostridial flavodoxins which are small, the protein from C. kluyveri falls in the group of higher molecular weight, and the protein from S. lividus is an exception among the cyanobacteria in having a low molecular weight. All flavodoxins contain a single molecule of FMN bound tightly but noncovalently to the protein, and this flavin functions as the only known redoxactive group in the protein. Edmondson and co-workers90"93 have observed covalently bound phosphate in Az. vinelandii flavodoxin; the group is not present in other flavodoxins, and its function in the Az. vinelandii protein is unknown. The content of this kind of phosphate has been reported to vary with the strain of Az. vinelandii and with the growth conditions.36 The flavin extracted from flavodoxin purified from a mutant strain of Az. vinelandii (str.

396

Chemistry and Biochemistry of Flavoenzymes

FIGURE 1. Comparison of amino acid sequences of flavodoxins. Az.v. = Azotobacter vinelandii'^ K.p. = Klebsiella pneumoniae;105 A.n. = Anacystis nidulans (Synechococcus PCC 6301);60 Ch.c. = Chondrus crispus;107 An.v. = Anabaena variabilis (PCC 7120);113 D.v. - Desulfovibrio vulgaris (Hildenborough);99-101-102 C.b. - Clostridium beijerinckii MP;% M.e. = Megasphaera elsdenii?7'9S C.p. = Clostridium pasteurianum.112

T2N 200) has different chromatographic properties from those of FMN and the absorption maximum in the visible spectrum is at 450 nm rather than 445 nm.94

IV. STRUCTURES The amino acid compositions of several flavodoxins are known and the sequence of amino acids in the protein has been reported for three of the smaller flavodoxins95102 and five of the larger molecules (Figure 1)^8,87,103-ioe an(j ^^^{^1 sequences are known for five others,95-107'112 including additional strains of An. nidulans (Synechococcus PCC 6301)107-110 and Az. vinelandii (strain O). HI All but three of the sequences were determined by sequencing the protein. The sequences for K. pneunmonmiae*1'104 Anacystis nidulans R2,60 and Ana-

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baena variabilis 7120, 113 flavodoxins were deduced from the base sequences of the genes. The base sequences of the genes for flavodoxins in D. vulgaris*00'102 and Az. vinelandii (strain OP)68 have also been reported; these confirmed the amino acid sequences and helped to resolve asn/asp and gln/glu assignments. Laudenbach et al.60 point out that the partial sequences of Synechococcus PCC 6301 (a k a Anacystis nidulans) differ from the corresponding sequences of A. nidulans R2 only at position 54 which is serine in PCC 6301 and cysteine in R2. They speculate that such a small difference might be due to a difference in strain. Eren and Swenson114 have chemically synthesized an artificial gene for flavodoxin from C. beijerinckii MP, expressed the gene in E. coli, and shown that the amino acid composition, the amino terminal sequence and other properties of the protein coded by the artificial gene are identical with those of the wild-type flavodoxin. The long-chain flavodoxins from Az. vinelandii, K. pneumoniae and A. nidulans R2 have been aligned in Figure 1, according to Drummond,104'115 with the complete sequence of A. nidulans R2 flavodoxin as given by Laudenbach et al.,60 and additional deletions in the 50 residues at the N-terminus to accommodate the sequences of the proteins from Ch. crispus106 and Anabaena variabilis.113 The alignment of the short-chain protein from C. beijerinckii MP with that from D. vulgaris is based on their known crystal structures.3 Spectroscopic measurements suggest that the structures of the M. elsdenii and C. pasteurianum proteins more closely resemble that of C. beijerinckii MP flavodoxin, rather than D. vulgaris flavodoxin, and the amino acid sequences of the three proteins are homologous over much of their length. The two groups of flavodoxins have been aligned with each other according to Dubourdieu and Fox" with additional deletions to accommodate the more recent sequences for the long chain molecules. Homology between the two groups in the carboxylterminal sequences is poor, and although minimal mutation frequencies were used to align this region, the alignment is not unambiguous." The alignment requires several spaces in the long-chain molecules and 41 spaces in the short-chain molecule. Dubourdieu and Fox" placed 22 of these spaces in the short-chain molecules in a block beginning at residue 134 of Figure 1; the crystal structure of An. nidulans (Synechococcus PCC 6301) flavodoxin suggests that the insertion in this region is in fact 20 residues long and, as shown in Figure 1, that it begins at residue 133.110 Other disparities between the alignments predicted from genetic considerations and the X-ray structures of flavodoxins have been noted.3 Inspection shows that the greatest region of homology occurs in the N-terminal sequence, with several residues being invariant, and there are two additional regions that are highly conserved (53 to 65 and 91 to 98 of Figure 1). In the known crystal structures, residues near the N-terminus contribute to the binding site for the phosphate of FMN (S or T(10), T(12) and T(15)), and the two other regions of high conservation either provide residues that directly contact the flavin or they immediately precede sequences which provide residues at the flavin-binding site. Crystal structures have been determined for oxidized flavodoxins from C. beijerinckii MP, D. vulgaris, A. nidulans and Ch. crispus. Structures have also been determined for the semiquinone and hydroquinone of C. beijerinckii MP flavodoxin, and for the semiquinones of the D. vulgaris and An. nidulans proteins. The structures are reviewed in detail elsewhere in this series.116 They all consist of a central parallel p-sheet consisting of five strands linked by four a-helical segments that occur in pairs on the outside of the molecule. The longerchain flavodoxin is distinguished chiefly by an insertion of 20 residues in a loop in the fifth strand of the p-sheet. In all of the structures, the FMN is bound on one side of the molecule with the ribityl phosphate side chain extending towards the center of the molecule. The negative charge on the phosphate is not neutralized by positively charged amino acid side chains; instead the phosphate is hydrogen bonded to hydroxy amino acids and backbone amide groups. There is much homology of amino acid sequence in the area of the flavin site, but the detailed structure around the flavin varies among the flavodoxins. For example,

398

Chemistry and Biochemistry of Flavoenzymes

although the isoalloxazine structure is sandwiched between hydrophobic amino acid side chains in all of the structures, different amino acids are involved. In A. nidulans flavodoxin, Trp-57 is adjacent to the face of the flavin nearer the interior of the molecule, while Tyr94 flanks the other face of the flavin (in Ch. crispus flavodoxin these residues are Trp-56 and Tyr-98);117 the corresponding residues in D. vulgaris flavodoxin are Trp-60 and Tyr98 respectively, while in C. beijerinckii flavodoxin they are Met-56 and Trp-90. The amino acid sequences suggest that in the long-chain flavodoxins from Az. vinelandii and K. pneumoniae the side chains of leucine residues (57 and 56, respectively) flank the inner face of the flavin. It is clear that these and other differences in the structures of the flavin-binding sites of the flavodoxins account for differences in their physical properties such as optical spectra and redox potential. Solvent access to the flavin appears to be restricted to the dimethyl benzene region. Consequently, it is often presumed that electron transfer to and from the flavodoxins occurs through this region of the FMN. The distribution of charged amino acids in flavodoxins has been extensively studied in recent years because electrostatic dipoles induced by such residues have been found to be important for electron transfer in other redox proteins, such as the cytochromes. The arrangement of negatively charged residues on the surface of C. beijerinckii flavodoxin is asymmetric with most of the charge clustered near to the flavin. This arrangement induces a dipole moment along an axis passing through the FMN118 and it is thought to be important in complex formation between flavodoxins and electron acceptors/donors (see later). The charge distribution in the longer chain flavodoxins is similarly asymmetric. In the proteins from the nitrogen-fixing organisms there is a cluster of positive charge (on a-helix 1) that is not present on the other proteins. Drummond104'115 has proposed that this region has a role in the interaction with other nif proteins.

V. OXIDATION-REDUCTION PROPERTIES The oxidation-reduction properties of the FMN in the flavodoxins differ markedly from those of FMN in free aqueous solution (see References 3 and 5 for a discussion), and much recent work has attempted to explain how the protein structure at the flavin-binding site effects these changes. As noted earlier, the flavin semiquinone is highly stabilized in the flavodoxins, and, moreover, the ionization of FMN semiquinone is suppressed so that the semiquinone is stabilized in its neutral form. The oxidized protein is converted to the semiquinone by addition of one reducing equivalent under anoxic conditions; addition of a further reducing equivalent generates the hydroquinone. The reducing equivalents can be provided electrochemically with dyes to mediate electron transfer from the metallic electrode to the protein,9-19-119 by pulse radiolysis to generate e aq " and CO2~~,120'123 chemically as dithionite ion,3 photochemically with 5deaza-flavins123 or plant chloroplasts as catalysts, or enzymically with such systems as hydrogen and hydrogenase and reduced nicotinamide nucleotides in the presence of ferredoxin-NADP+ reductase.3'5-6-8 All of these agents reduce flavodoxins to the semiquinone, but the extent of reduction beyond the semiquinone depends on the relative redox potentials of the flavodoxin and the electron donor. Photochemical reduction with 5-deaza-flavin provides a convenient and rapid method of reducing flavodoxins fully to the hydroquinone. Anderson et al.122 showed that the anionic flavin semiquinone is formed within 2 ms of pulse radiolysis of M. elsdenii flavodoxin at pH 6.1 or pH 9.15 in aerobic solution. The anion then protonates rapidly to the neutral semiquinone (rate constant = 2.6 X 105 s" 1 at pH 6.1). Earlier pulse radiolysis experiments on this flavodoxin121 and on flavodoxin from C. beijerinckii MP120 reported fast reactions which were attributed to protein conformational changes. It now seems likely that both were due to formation of the semiquinone anion and its subsequent protonation. Faraggi and Klapper120 reported a slow reaction that

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occurred over the millisecond timescale and they also attributed this reaction to a conformational change; however, it was pointed out later that such changes are artifacts that result from the use of terf-butanol to scavenge hydroxyl radicals.121 Mayhew and Massey124 have noted that contamination by oxygen in a photoreduction experiment (and also in experiments using hydrogen as reductant with crude preparations of hydrogenase76) can lead to the formation of a novel red derivative of FMN in flavodoxins. The rate and extent of conversion to the red derivative depends on the oxygen concentration and on the pH (it is enhanced at low pH), and full formation requires a high concentration of a carboxylic acid or an alcohol. The absorption spectrum of the modified flavodoxin has a broad maximum at 505 nm (e505 = 7.3 mM" 1 cm" 1 ) with additional new peaks at 308, 322, and 350 nm. The visible absorbance decreases reversibly at high pH. The derivative is very stable in air, but since an excess of ferricyanide ion slowly converts it to oxidized flavodoxin, it nevertheless appears that the derivative is a reduced flavin. The modification does not involve a covalent interaction with the protein because the chromophore remains in solution when the apoprotein is precipitated with acid, and the red complex is restored by addition of fresh apoflavodoxin to the free chromophore. The detailed chemical nature of this derivative is not known, but NMR measurements suggest that it is a 1,8-dihydroflavin with an alkyl residue added at C(8).125 The derivative also seems to occur naturally in preparations of flavodoxin from C. kluyverii11 and some preparations of C. beijerinckii MP flavodoxin.76 Dithionite ion has been widely used as the reductant for flavodoxins and its use has resulted in a number of anomalous observations on the reactions of flavodoxins (and other low potential electron carriers). Many of these became explicable when it was shown that the reductant of flavodoxin in solutions of sodium dithionite ion is most probably SO2_ formed by dissociation of S2O42",126'127 and that the redox potential of this radical changes with pH and the concentration of dithionite ion (see References 128 to 130 for a discussion). The redox potentials of some of the flavodoxins are so negative that sodium dithionite at pH 7 is not sufficiently reducing to convert them completely to the hydroquinone, and the difficulty is compounded if, as is usually the case, the dithionite ion is contaminated with its oxidation product (bi)sulfite.128*130 The kinetics of reduction of several flavodoxins by sodium dithionite have been studied;126-131 Thorneley & Deistung131 showed that in keeping with the Marcus theory of electron transfer,132 there is a linear relationship between the log of the rate constant for reduction of flavodoxin semiquinones by SO2_ and the redox potential of the semiquinone/hydroquinone couple. Mixtures of fully oxidized and fully reduced flavodoxin from M. elsdenii rapidly cornproportionate to generate the semiquinone,17 making it clear that the semiquinone is in equilibrium with the other two redox species and that stabilization of the semiquinone is truly thermodynamic, rather than simply a result of a low rate of intermolecular one-electron transfer in solutions of the protein. A similar thermodynamic stabilization of the semiquinone is likely for other flavodoxins. The high stability of the semiquinone in flavodoxins allows the oxidation-reduction potentials for the two one-electron steps to be determined readily by a variety of electrochemical, spectrophotometric and enzymic methods.3-19 The values of E2 (quinone-semiquinone) and E, (semiquinone-hydroquinone), and the difference between them (E 2 -E I ), vary with pH and with the source of the flavodoxin (Table 1). The effects of pH on the potentials have been determined for several proteins, including those from M. elsdenii,17J9 C. pasteurianum,9 C. beijerinckii MP,9 A. nidulans (Synechococcus PCC 6301),42 D. vulgaris^3 and two species of Anabaena (PCC 7119134 and PCC 712049). They show that the plot E2 vs. pH is linear over the physiological range of pH with a slope of about -60 mV, consistent with the addition of a single electron and a single proton, while over similar pH range, the plot of E t vs. pH is either independent of pH or it changes from pH-independent at the upper end of the range to a slope of about - 60 mV

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Chemistry and Biochemistry of Flavoenzymes

at low pH. The slope change for E! vs. pH, first observed with flavodoxin from M. elsdenii,17 but since observed with other flavodoxins,9'42'133'134 indicates that there is a redox-linked pK associated with the hydroquinone. The value for this pK in M. elsdenii flavodoxin is 5.8, and values determined for the other flavodoxins are not too different. The pK was ascribed to N(1)H of FMNH2 in flavodoxin 17 because in protein-free FMNH 2 , a pK associated with N(1)H occurs at 6.7.135 However, NMR measurements of 15N-enriched FMN in several flavodoxins136"139 have shown that the chemical shifts are characteristic of the flavin hydroquinone anion, with no evidence for protonation at N(l), even at pH values lower than the pK indicated by the redox measurements. These observations imply that the pK for the flavin hydroquinone is shifted to a very low value when FMNH2 is bound to apoflavodoxins, and that the change in slope of E/pH for some flavodoxins must be due to the redox-linked protonation of another group, most probably the side chain of an amino acid residue. An elegant model has been proposed by Ludwig et al.116-140 to account not only for the shift to a smaller value of pK in the hydroquinone, but also for the shifts to greater values of the pKs of the semiquinone and quinone. Important features of this model include a steric restriction to protonation at N(l) of the hydroquinone, repulsion of the negative charge at N(l) by neighboring charged groups on the protein and a resultant destabilization of the hydroquinone relative to the quinone and semiquinone, and shifts of the pK of a neighboring carboxylate (Glu-60 and Glu-59 in M. elsdenii and C. beijerinckii MP flavodoxins, respectively) to a smaller value in the quinone and semiquinone, but to a greater value in the hydroquinone. It is proposed that it is the ionization of this group that is detected in the measurements of E^ The measured potentials can be used together with the measured dissociation constant for dissociation of FMN from flavodoxin and reported values of the redox potentials of protein-free FMN to calculate dissociation constants for the dissociation of the semiquinone and fully reduced forms of the flavin.3-141 These calculations show that the semiquinone is bound more tightly than the oxidized flavin; binding of the hydroquinone, although still strong, is even weaker than the oxidized flavin. For example, KD for flavodoxin from M. elsdenii is approximately 0.4 nM;125 using potentials of —372 mV and — 115 mV for El and E2 (Table 1), the calculated KD values for the semiquinone and hydroquinone are 3.4 fM and 9.1 nM, respectively, based on potentials for FMN determined by Draper and Ingraham,135 and 0.18 fM and 2.9 nM, respectively, using the potentials for FMN determined more recently by pulse radiolysis.142 Protein-flavin interactions that stabilize the semiquinone, that shift the pK of the semiquinone from about pK 8,6 in free FMN to greater than 10.5 in the flavodoxins, and which dramatically change the affinity of flavin and protein with redox state have been suggested from the known crystal structures. For example, it has been proposed that the stabilization of the semiquinone in its neutral form is due in part to a hydrogen bond between N(5)H of the flavin and a carbonyl oxygen atom of the protein backbone.110'143 The carbonyl is provided by glycine residues in C. beijerinckii MP (G57) and D. vulgaris (G61) flavodoxins but by an asparagine (N58) in A. nidulans (Synechococcus PCC 6301) flavodoxin. A protein conformational change accompanies reduction of oxidized FMN in the protein to the semiquinone, and brings the carbonyl into close contact with N(5)H of the flavin. The presence of a hydrogen bond at N(5) is supported by NMR measurements on fully reduced flavodoxins. 137139 The semiquinone of the A. nidulans (Synechococcus PCC 6103) flavodoxin is somewhat less stable than the semiquinones of the other two proteins, and it has been suggested that this might be caused by the relatively bulky side chain of Asp-58 hindering the conformational change.110 Mutant proteins produced by site-directed mutagenesis are now becoming available, and they will allow this idea and other suggestions about the roles of proposed protein-flavin interactions to be tested by varying the nature of the side chains of specified amino acids. The semiquinone of a mutant of D. vulgaris flavodoxin which

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has an asparagine residue instead of glycine at position 61 is less stable than the semiquinone of the native protein, and measurements of the redox potentials and FMN dissociation constant suggest that there is a relatively large effect on the binding of the semiquinone; the K D values for FMN, FMNH- and FMNH~ were decreased by factors of 13.3, 923, and 21.2fold, respectively, compared with the corresponding values for the native protein.29-144 Tyr98 has been substituted by a variety of amino acids;145 the neutral semiquinone is stabilized in the mutants but E} depends on the properties of the substituent for Tyr-98. Early studies by X-ray crystallography on model flavins suggested that the flavin hydroquinone in free solution is bent along the N(3)-N(10) axis146 with a "butterfly" motion of the molecule about this axis.147 The large change in the redox potential of the semiquinone/ hydroquinone couple of FMN in the flavodoxins was originally attributed to the protein constraining the flavin hydroquinone to a planar conformation.148 Moonen et al.149'150 have since shown that fully reduced flavins in solution are, in fact, planar, and therefore the planarity of protein-bound FMN can have little effect on the redox potential E,. They proposed that interactions between the negative charge on N(l) of the hydroquinone and the two negative charges on the flavin phosphate could account for the relative instability of the hydroquinone-protein complex and the consequent large shift of the redox potential to a more negative value.151 Values calculated for the redox potentials of flavodoxins on the basis of this theory were found to be in fair agreement with measured values, and an increase of negative charge on the flavin side chain by addition of a second phosphate group at the C(3') position (riboflavin 3',5'-bisphosphate) shifted the redox potential to even more negative values, as predicted.20 However, the theory requires a value for the electrical permitivity of the protein, and in the absence of an experimental value, it was necessary to use an assumed value. It seems likely that the destabilization of the hydroquinone is due partly to interaction of the negative charge at N(l) and negatively charged side chains of nearby amino acids, as proposed in the model of Ludwig et al.116-140

VI. INTERACTION OF FLAVIN AND PROTEIN The FMN of flavodoxins is bound tightly but noncovalently to the apoprotein and these two components of the holoprotein can be separated by a variety of methods, as summarized by Mayhew and Ludwig.3 Extraction and precipitation of the apoprotein with trichloroacetic acid, using methods similar to those first described for Az. vinelandii flavodoxin,152'153 has come to be widely used because it it rapid, and, despite its strongly denaturing conditions, it provides apoprotein in high yield from several flavodoxins. These can be fully reconstituted to their holoproteins by subsequent addition of FMN. The use of EDTA throughout the procedure is recommended to minimize metal-catalyzed oxidation of thiol groups in the protein and the associated loss of FMN binding activity that occurs with certain apoflavodoxins.141 The reassociation of FMN with apoflavodoxin from several sources leads to complexes that are indistinguishable from the native flavodoxins. The fluorescences of FMN and apoflavodoxin are quenched in the complex, and there are also qualitative and quantitative changes in the visible and near UV absorption and CD spectra. These spectroscopic changes provide convenient methods for following the association and dissociation reactions. Early experiments to investigate the kinetics and thermodynamics of the interaction of FMN and other flavins with apoflavodoxins, mainly with proteins prepared from Az. vinelandii and A/, elsdenii, have been discussed elsewhere.3-5 More recently, Gast et al.154 have explored the effects of pH and ionic strength on the interaction of FMN with apoflavodoxin from M. elsdenii by measuring the change in fluorescence following dilution of the holoprotein into buffer of known ionic strength, and in the pH range 2 to 5. The fluorescence increased with time until a new equilibrium was reached:

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Chemistry and Biochemistry of Flavoenzymes

Values for the dissociation rate constant, koff, were determined from the progress curves, and values for the dissociation constant, KD, were determined from the position of the new equilibrium at completion of the reaction. Since K D = koff/kon, values for the association rate constant could be calculated from the experimental values for koff and K D . All three constants were found to vary with pH and ionic strength (I), and plots of the logarithm of each constant vs. the square root of I were linear. The effects of a change in ionic strength varied with the pH. An increase of ionic strength at pH values smaller than 3.8 increased koff and K D and therefore destabilized the holoprotein; in contrast, similar increases in ionic strength above pH 4.2 decreased koff and KD and led to a greater stabilization. The calculated values for kon showed that the rate of association is much more sensitive to ionic strength than the rate of dissociation. The data, interpreted according to Bronsted theory, indicated that the apoprotein reaches its maximum net positive charge at pH 2.0 to 2.6 and that its isoelectric point is at about pH 4. The net charge calculated was 11 to 12, in agreement with a theoretical value of 12 deduced from the primary structure. Extrapolation of the data to zero ionic strength showed that kon is 3.2 x 105 M~ l s^ 1 at 22° and independent of pH; the calculated value agreed well with an earlier direct measurement of kon when the earlier data were also corrected to zero ionic strength and corrected for a difference in temperature using a published value for the activation energy of the reaction.141 After extrapolation to zero ionic strength, koff and KD still depend on pH, and the pH profile was consistent with two protein ionizations with a pK value of 3.4. It was suggested that the groups might be the carboxyls of two glutamic acid residues (60 and 61) which are in the neighborhood of the flavin if structural homology is assumed between flavodoxins from M. elsdenii and C. beijerinckii MR More recently, it was proposed that the pK for Glu60 in M. elsdenii flavodoxin varies with the oxidation state of the flavin. 116 - 140 A study using temperature-jump techniques with monitoring of the flavin fluorescence suggested that two relaxations occur with Az. vinelandii and M. elsdenii flavodoxins following perturbation of the temperature.155 The observations were consistent with two steps in the binding of FMN to the apoproteins. However, Gast and Miiller156 could not detect a relaxation under the conditions used in the earlier study, and they suggested that the temperature jump technique cannot be applied to native flavodoxins because the interaction between apoprotein and FMN is too strong. Thy therefore carried out similar experiments using the weaker complex of M. elsdenii apoflavodoxin and deoxy-FMN, a compound that lacks the hydroxyl groups on the ribityl side chain of FMN. A single relaxation was observed with this complex, and also with complexes of Az. vinelandii apoflavodoxin and a series of model flavins that carry charged or uncharged side chains of differing length. It was concluded that the binding reactions occur in a single step. A similar conclusion has been drawn from temperature-jump and stopped-flow spectrophotometry experiments of flavodoxin from D. vulgaris.157 Visser et al.158 studied the effects of pressure in the range 1 x 10~ 3 to 11 kbar on the flavin and protein fluorescence of a series of flavodoxins and of apoflavodoxin from M. elsdenii. Both reversible and irreversible increases in the flavin fluorescence occurred, and the extent of the irreversible change was greater at pH 7.5 than at pH 5. The sensitivities of the flavodoxins to pressure were found to vary with the source of the protein, and to decrease in the order D. vulgaris, M. elsdenii, C. beijerinckii and Az. vinelandii. It was suggested that the irreversible change involved a covalent modification of the flavin-binding region of the protein, but physicochemical analysis of the inactive product was not reported. Very few of the amino acids that interact with FMN in native flavodoxins can be chemically modified under sufficiently mild conditions to avoid denaturation of the protein, and early experiments (reviewed in Reference 3) to explore the effects of modifying tyrosine and tryptophan have not been extended. Site-directed mutagenesis should allow the introduction of groups that can be more readily modified and used as centers for the introduction

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of structural probes. Although thiol groups do not directly interact with the flavin in those flavodoxins for which crystal structures are known, their modification in several of the smaller flavodoxins by a variety of reagents was found to prevent FMN binding. The modifying groups that were introduced were quite large and it was suggested that their inhibition of flavin binding arises indirectly by effects on the secondary and/or tertiary structure of the protein (see Reference 3 for a review). It was observed recently that when the two thiols of M. elsdenii apoflavodoxin are converted to thiocyanate groups by cyanylation of the mixed disulfide of apoprotein and thionitrobenzoate, the resulting protein still binds flavin, albeit 50,000 times more weakly than the native protein.159-160 Evidently in this case the small size of the modifying group causes much less desruption of the protein structure. The use of 13C-enriched KCN in the cyanolysis allowed the introduction of groups that can be detected by NMR. Comparison of the two main NMR signals observed in the apo- and holoproteins with those of model compounds in solvents of differing polarity allowed the conclusion that the two thiocyanates in the apoprotein are in a hydrophobic environment. The environment of one of the groups changes drastically when FMN is bound so that it is now much more polar, and possibly exposed to solvent. The changes suggest that one of the thiols in a region of the protein that undergoes significant conformational change when FMN is bound. 16° On the basis of circular dichroism and fluorescence measurements, D'Anna and Tollin161 also concluded that protein conformational changes accompany the binding of FMN. Substitution of FMN in flavodoxins with other flavins has been used widely to explore the flavin-binding site. Such studies have employed flavins modified at almost every atom in the molecule, and, as reviewed by Ghisla and Massey,162 a wide range of modified flavins is now available to probe many properties of the binding site. Early experiments to investigate the specificity of apoflavodoxins for the flavin prosthetic group showed that they can be divided into two groups based on their requirement for a terminal phosphate on the ribityl side chain of FMN; one group, that includes the apoproteins of Az. vinelandii, D. vulgaris, and A. nidulans flavodoxins, binds riboflavin, while apoproteins in the other group, including those of flavodoxins from M. elsdenii and the Clostridium spp., have a strict requirement for the 5'-phosphate group of FMN. Apoproteins in the second group do not interact with riboflavin to an appreciable extent, or with riboflavin 4'-phosphate,163'164 but they bind FMN as avidly as proteins from the first group, and they also form a strong complex with riboflavin 3', 5'-bisphosphate.20*163'164 The apoproteins of all of the larger flavodoxins examined so far bind riboflavin, and, with one exception, the smaller apoflavodoxins do not. However, the division according to flavin specificity does not coincide with the division based on size, since D. vulgaris apoflavodoxin, a small protein, forms a relatively strong complex with riboflavin. The explanation for the requirement for the phosphate group in stabilizing the complexes is not known. The relatively high specificity of proteins in the second group was instrumental in the recognition that many commercial preparations of FMN contain much phosphorylated flavin impurity,165 and prompted the analytical use of these apoproteins in the determination of FMN by fluorescence titration, and of FAD after enzymic hydrolysis,165'167 and in the assay of flavokinase.168 A preparative procedure has also been described for the purification of FMN by column chromatography using immobilized apoflavodoxin from M. elsdenii as an affinity ligand.169 However, it is clear now that although this procedure provides a rapid and convenient way of separating riboflavin 5'-phosphate (FMN) from most side products of the chemical phosphorylation of riboflavin, it does not remove the small amount of riboflavin 3',5'-&/sphosphate that is formed in the reaction and which contaminates commercial supplies. The very high affinity of M. elsdenii apoflavodoxin for FMN (Kd approximately 0.4 nM) has been shown to provide a uniquely nondisruptive method for extraction of flavin from the FMN-containing diaphorase of C. kluyveri.170

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Chemistry and Biochemistry of Flavoenzymes

TABLE 2 Flavins Modified in the Isoalloxazine System That are Bound by Apoflavodoxins Flavin modification 1 -Carba- 1 -deaza-FMN 2-Thio-FMN 4-Thio-, 4-NHCH3-, 4-NOH-, 4N(CH 3 ) 2 - f and 4-S-CONH2-FMN 4a(n-Propyl)4a,5-dihydro-FMN 5-Carba-5-deaza-FMN 5-Thia-5-deaza-FMN 5,6,7,8-Tetrahydro-FMN 6-Hydroxy-FMN 6-Thio-, 6-azido-, 6-cyano-, and 6amino-FMN 6,7-Dimethyl-FMN (iso-FMN) 7-Chloro-FMN 7,8-Dichloro-FMN 8-Hydroxy-FMN 8-Mercapto-FMN 8-Chloro-FMN 8-Azido-FMN 8a-[5(N-acetyl)-L-cysteinyl]-FMN 8a-[S(yV-acetyl)-L-cysteinyl]-tyrosinyl-riboflavin and FMN 8a-Imidazoyl-FMN a

Source of apoflavodoxin

Ref."

D. vulgaris, M, elsdenii, C. beijerinckii M. elsdenii M. elsdenii

116, 140, 172, 173 174, 175 176

M. elsdenii Az. vinelandii, M. elsdenii M, elsdenii C. pasteurianum M. elsdenii M. elsdenii

177 178, 179 180 181 182 183—185

M. elsdenii C. pasteurianum M. elsdenii, C, pasteurianum M. elsdenii, D. vulgaris, Az, vinelandii M. elsdenii M. elsdenii, C. pasteurianum M. elsdenii Az. vinelandii Az. vinelandii

141 181, 186, 187 3, 181, 186, 187 188—190 191 192, 181, 186, 187 193, 194 195, 196 195, 196

Az. vinelandii, C. pasteurianum

197—199

Additional flavin complexes are given in Reference 3.

Gast and Miiller156 studied the effects on the kinetics and thermodynamics of interaction with Az. vinelandii apoflavodoxin of systematically changing the length of the side chain from 1 to 6 carbon atoms in flavins with either no terminal charged group, or with a phosphate or carboxylate at the terminus. They confirmed an earlier finding153 that the rate of association is lower with flavins that carry a terminal ionizable group, and suggest that this might result from a requirement for dehydration of the ionized group prior to binding. A further notable finding in this work was that the complexes with the charged side chains on the flavin become weaker as the length of the side chain decreases, until there is no detectable binding of the derivatives that have only two carbon atoms separating the charged group from the isoalloxazine. In contrast, the flavin with an uncharged side chain of only two carbon atoms is bound almost as tightly as riboflavin. These observations may be an indication of strong repulsive forces between the charged flavin side chains with 1 or 2 carbon atoms and negatively charged groups in the binding site for the isoalloxazine of FMN in Az. vinelandii flavodoxin. Carlson and Langerman171 used calorimetry to study the binding of FMN, riboflavin, 8-carboxylic acid riboflavin and FAD to apoflavodoxin from Az. vinelandii. They concluded that the interaction with FMN is pH-independent in the pH range 6 to 9, confirming that the phosphate mono- and dianions are bound, and that there is no proton flux associated with the binding. The dependence of the enthalpy change on temperature was -220 cal deg" 1 mol~ l below 20°C and -820 cal deg"1 mol" 1 above this temperature, suggesting that the protein exists in two conformations, both of which bind FMN. Comparison of the enthalpy change due to FMN binding (-28 kcal mol" 1 ) with that for riboflavin (-16.7 kcal mol" 1 ) indicated that the ribityl phosphate moiety provides approximately half of the enthalpy of binding of FMN. Apoflavodoxins bind a wide range of flavins that are modified in the isoalloxazine structure (Table 2). The use of such modified flavins has provided information about the

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environment of the FMN-binding site and about the way in which it changes with redox state and pH, and also information about the exposure of different parts of the isoalloxazine structure to solvent and to external modifying groups and electron donors and acceptors. Much of this information is consistent with the known crystal structures. For example, the strong binding by Az. vinelandii apoflavodoxin of flavins which contain bulky substituents at the 8a-position,195'197'198 and the high chemical reactivity of 8-mercapto-FMN in M. elsdenii flavodoxin with the external reagents iodoacetamide and iodoacetic acid191-192 is in keeping with the crystal structures of other flavodoxins which show that the edge of the dimethyl benzene of the flavin is accessible to solvent. Access to the 8-position is not completely unrestricted in M. elsdenii flavodoxin, however, because thiophenol reacts poorly with 8-CI-FMN in this protein.192 Similarly poor reactions of 2-thio-FMN-flavodoxin with the thiol reagent methyl methanethiolsulfonate,174 and of 4-thio-FMN-flavodoxin with hydroxylamine176 are consistent with the crystal structure of the homologous flavodoxin from C. beijerinckii that the 2- and 4-positions of the native protein are shielded from direct contact with solvent. The relatively high reactivity of 6-cyano- and 6-mercapto-FMN-flavodoxin from M. elsdenii with modifying reagents, and the low yield of covalently bound flavin when 6-azido-FMN flavodoxin is photoirradiated implies that the 6-position is accessible to solvent.183"185 Computer graphics analysis of the homologous protein from C. beijerinckii showed that solvent can reach this position through a groove on the si face.185 Ludwig et al. 116J4 ° employed 1-carba-l-deaza-FMN to study the protein-flavin interactions at the 1-position of C. beijerinckii and M. elsdenii flavodoxins, using X-ray crystallography and thermodynamic measurements of the flavin-protein association constants and redox potentials as a function of pH. They showed that in the crystal structure, the protein in the region of Gly-89 moves to accommodate the C(1)H group, and they observed that while the redox potentials of this derivative vary in a similar way with pH to those of the native flavodoxin, the pK detected in the hydroquinone is associated with the flavin itself, rather than with a group on the protein. Information about the charge distribution in the flavin-binding sites has been obtained from studies with 6-hydroxy-,182 8-hydroxy-,18819° 6-mercapto-,183 and 8-mercapto-FMN.191t192 These flavins exist in different tautometric and/or mesomeric forms, depending on the ionization state and the environment, and the absorption spectra of the different tautomers and mesomers are sufficiently different to allow them to be identified in complexes with proteins. In addition, the pK values of their ionizable groups are in a range (3.8 to 7.1) that is appropriate for studies with proteins. Ghisla and Massey162 have pointed out that these ionizable flavins mimic the ionization behavior of normal reduced flavin. The first detailed studies of interactions with the flavodoxins were carried out with the hydroxy-flavins. These showed that apoflavodoxin stabilizes the orange 8-OH-FMN in the neutral form, and comparison of the shape of the absorption spectra with model compounds indicates that the proton is on the hydroxyl at C(8)-O rather than at N(1)-C(2).188 The pK for the hydroxyl group is shifted from 4.8 in the free flavin to 6.1 in the complex with M. elsdenii apoflavodoxin, in keeping with the flavodoxin crystal structures which show that the flavin-binding sites are negatively charged. Most studies with modified flavins have employed only one apoflavodoxin (often from M. elsdenii or Az. vinelandii). However, in the case of 8-OHFMN, the properties of the complexes of the modified flavin and apoflavodoxins from three species have been compared.189 The shifts of the pK that occur when 8-OH-FMN is bound to the apoproteins from Az. vinelandii and D. vulgaris flavodoxins are similar to but smaller than those observed with M. elsdenii apoflavodoxin, but the absorption spectra of the flavin anions in the three complexes are different, as are the spectra of the native proteins, and they clearly reflect differences in the local environment provided by the protein. In contrast to the spectra of the complexes with the other two proteins, which are unresolved, the spectrum of the complex with D. vulgaris apoflavodoxin is resolved into two distinct maxima,

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Chemistry and Biochemistry of Flavoenzymes

possibly reflecting a particularly hydrophobic environment provided by the side chains of tyrosine and tryptophan which sandwich the isoalloxazine in this protein. The redox properties of the complexes of the flavins of Table 2 with apoflavodoxins are generally consistent with those of the native protein and of the protein-free flavin (see also References 3, and 121). With only a few exceptions, the complexes with oxidized flavins are reduced stepwise to the neutral semiquinone and then to the hydroquinone. The complexes with 5-deaza-flavins undergo a two-electron reduction, consistent with the redox properties of the protein-free flavin, 178J79 and the flavins with bulky substituents at the 4position also fail to stabilize a semiquinone, possibly because the substituent blocks the formation of a hydrogen bond to the N(5)H of the semiquinone.162 The visible absorption spectra of the hydroquinones of the complexes of 1-deaza-FMN140 and 4-thio-FMN176 with M. elsdenii apoflavodoxin at neutral pH are characteristic of their protonated forms. This is in contrast to the hydroquinone of the FMN complex which is stabilized as the anion. The hydroquinones of the modified flavins can be protonated at the 2- or 4-oxygen (1-deazaFMN) or the 4-sulfur atoms, however, and it is likely that in the flavodoxins protonation at these positions is still possible.140 Harzer and co-workers200-201 and Visser et al.202 showed that apoflavodoxins bind lumazine derivatives and that the properties of the complexes of 6,7-(2,3-dimethyl)-A^(8)ribityllumazine-5'-phosophate200202 and 6,7-(2,3-dimethylbutano)-Ar(8)-ribityllumazine-5'phosphate201 resemble those of native flavodoxin; one-electron-reduced intermediates are formed and the complexes have catalytic activity in the photosynthetic reduction of NADP + by chloroplasts that is surprisingly high given current ideas regarding the entry of electrons via the dimethylbenzene of FMN in native flavodoxins (see later) and the structures of these lumazine derivatives in which the corresponding region is either lacking or not conjugated. Time-resolved fluorescence studies of flavodoxin have provided some insights into the dynamic features of the FMN-protein interaction.203 The FMN fluorescence in flavodoxins is strongly quenched, amounting to about 1 to 5% of the fluorescence of free FMN. This is due to interactions which occur in both the ground and excited states of the flavin. Interactions in the ground state with aromatic amino acids are the main source of the static quenching (which does not change the fluorescence lifetime), whereas dynamic interactions occurring in the first excited singlet state lead to shortening of the fluorescence lifetime. The experiments revealed a flavin fluorescence component with a lifetime of 30 to 40 ps in D. vulgaris flavodoxin, and one with a longer lifetime of 5.6 ns. Similar results were obtained with the D. gigas flavodoxin, with lifetime components of 20 ps and 4.8 ns. The short-lived component from D. vulgaris flavodoxin was associated with dynamic quenching that resulted from the formation of an exciplex with the side chain of the binding-site tryptophan, and electron transfer from the indole ring to the isoalloxazine structure. The longer-lived emission could be due either to fluorescence from the exciplex itself, or to emission from a population of flavodoxin molecules with a transiently more open and more fluorescent conformation. This latter explanation is consistent with the fact that the 5.6 ns lifetime of this component is longer than that of free FMN, which could be a consequence of the more apolar environment provided by the aromatic residues in the FMN binding site. It is also consistent with the observation that the long-lived component of D. gigas flavodoxin has a shorter lifetime than is the case for the D. vulgaris protein, an observation which correlates with the lower aromatic environment of the FMN binding site (absence of Trp) in the D. gigas protein. The shorter lifetime component in the D. gigas flavodoxin is presumably also due to exciplex formation.

VII. ELECTRON TRANSFER MECHANISMS The mechanisms by which electrons are transferred between the FMN of flavodoxins and other redox centers have been studied intensively using flavodoxin semiquinone as the

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TABLE 3 Effect of Chemical Modification of the FMN Cofactor of C. pasteurianum Flavodoxin on Second-Order Rate Constant for Electron Transfer from the Semiquinone to Horse Cytochrome c Cofactor

FMN 8-IMF

FMN 7,8-diClFMN 5,6,7, 8-Tetrahydro FMN a

Ionic strength (mA/)

90 90 100 100 100

k2 (A/'1 s-1) 5.1 2.4 1.5 1.1 3.9

x x x x x

1053 104 106 105 105

Ref.

199 199 204 181 181

This value is somewhat smaller than expected based on the earlier experiments of Simondsen and co-workers.205 This probably reflects differences in experimental conditions and in flavodoxin preparations. However, values within each set of experiments are comparable.

electron donor and a variety of biological and nonbiological compounds as electron acceptors. Several considerations make flavodoxins useful objects for the study of biological electron transfer. First, crystal structures are available for flavodoxins from four species, and their structures at the FMN-binding site show substantial variation. Second, the semiquinone of flavodoxins is readily formed by photochemical reduction of the oxidized molecule, and although the semiquinone is fairly unreactive with dioxygen, its redox potential is low enough to allow it to reduce many other redox compounds of biochemical interest. Third, as noted above, there is an area of negative charge associated with the surface of the molecule near to the FMN binding site, and this makes flavodoxins particularly suitable for investigating the influence of electrostatic interactions on electron transfer processes. Fourth, the FMN redox center of flavodoxins can be replaced by a wide variety of flavin analogues to generate stable, redox-active complexes with properties different from those of the native protein. These considerations make flavodoxins useful not only as models for other flavoproteins, but also to establish the general principles of biological redox processes. This section provides an overview of these studies. A. KINETIC EVIDENCE THAT THE EXPOSED DIMETHYLBENZENE RING OF FMN IS INVOLVED IN ELECTRON TRANSFER The crystal structures of flavodoxins showed that the dimethylbenzene ring of FMN is exposed to solvent, and it was suggested that electron transfer to and from the flavin probably occurs through this region of the molecule.3-5 This proposal has been investigated by substituting FMN in flavodoxins with FMN analogues, including the 7-chloro-, 8-chloro- 7,8dichloro-, 5,6,7,8-tetrahydro-, and 8-a-N-imidazolyl (8-IMF) derivatives, and using stopped flow spectrophotometry and laser flash photolysis methods to compare the kinetics of electron transfer from these complexes with those from the native protein.186'187-199 Table 3 presents second order rate constants for electron transfer from the semiquinone of C. pasteurianum flavodoxin to horse cytochrome c. The dependency of the rate constants on the concentration of cytochrome c is nonlinear, especially at low ionic strength, and a two-step kinetic mechanism that involves formation of a protein-protein complex, followed by electron transfer from FMNH- to the heme, has been proposed (see References 205 and 206 for discussion). The rate constants of Table 3 are for the step in which the complex is formed, and they show that when the dimethylbenzene ring of FMN is modified, the rate of formation of the complex decreases. The rate constant also decreases with increasing

408

Chemistry and Biochemistry of Flavoenzymes TABLE 4 Effect of Chemical Modification of the FMN Cofactor of C. pasteurianum Flavodoxin on Limiting First-Order Rate Constant for Electron Transfer from the Semiquinone to Horse Cytochrome c Cofactor

FMN 8-IMF

FMN 7,8-Dichloro-FMN 5,6,7, 8-Tetrahydro-FMN a

Ionic strength (mAf)

k. (s-1)

45 45 65 65 65

236a

7 78 23 3

Ref.

199 199 205 181 181

This value is somewhat larger than expected based on the earlier experiments of Simondsen and co-workers (cf. footnote to Table 3).

ionic strength, and this ionic strength dependency is not appreciably affected by changes in the structure of the flavin, in accordance with the conclusion that the ionic strength effects are due to the protein rather than the flavin (see below). The association constant of the complex is also independent of the flavin structure for the analogues of Table 3. It is concluded that in the intermediate complex of the two proteins, the dimethylbenzene moiety of the flavin and a surface of the cytochrome that is most likely the exposed heme edge, as described later, are within van der Waals distance of one another. The limiting first order rate constants for the reactions of Table 3 also decrease when the dimethylbenzene ring is modified (Table 4), indicating that the second phase of the reaction, the electron transfer step and any associated structural rearrangements, is also directly affected by alterations in this region of the flavin. The modifications made to the flavin in the analogues of Tables 3 and 4 cause changes in the flavin redox potentials, but such changes cannot account for the kinetic effects observed (see References 181 and 199 for discussion). Therefore, the simplest interpretation of the data is that outer sphere electron transfer occurs between the two proteins, and that it involves orbital overlap between the TT electron systems of the two prosthetic groups. The extent of such overlap will depend on both the distance between the heme and the flavin and their mutual orientation, and these two factors must also help to determine the values for the rate constants. De Francesco et al.199 showed that the rate constants for electron transfer from the semiquinones of C. pasteurianum and Az. vinelandii flavodoxins to horse cytochrome c are independent of pH. They concluded that electron transfer from the neutral semiquinones is not coupled to proton release, and that the proton is lost before the processes associated with electron transfer occur. The conclusion is consistent with the observations of van Leeuwen et al.121 that the rate of reduction of Az. vinelandii flavodoxin to the semiquinone by methyl viologen radical is also independent of pH, and with pulse radiolysis experiments with M. elsdenii flavodoxin122 in which the rate constant for protonation of the anionic semiquinone was found to change from 1.1 x 105 s" 1 to 2.6 x 105 s"1 as the pH was changed from 9.2 to 6.1. B. EVIDENCE FOR ELECTROSTATIC MODULATION OF THE REDOX KINETICS The possible involvement of charged amino acid side chains in the interaction of flavodoxins with oxidants has been investigated by measuring the effects of a change in ionic strength on the kinetics of electron transfer. The flavodoxins for which crystal structures

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TABLE 5 Electrostatic Analysis of Kinetic of Oxidation of C. pasteurianum Flavodoxin Semiquinone by Various c-Type Cytochromes204 Cytochrome source Rsp. tenue 3761 Euglena sp. Rm. vanniellii Rps, capsulata Rsp. rubrwn Ps. aeruginosa Candida krusei Tuna Horse P. denitrificans Chi. thiosulphatophilum E. halophila a

b

Net charge

Va« (kcal/mol)

Z,b

4+

-5.6 + 7.0 -10.3 -11.8 -13.3 + 3.2 + 16.2 -16.0 -23.5 -12.3 -9.5 + 6.5

+ 1.6 -2.0 + 3.0 + 3.4 + 3.9( + 4) -0.9(+1) + 4.7 + 4.7( + 4) + 6.8( + 4) + 3.6( + 4) + 2.8( + 2) -1.9

2+ 1+ 0 25+ 7+ 7+ 7_ 6+ 10-

Vjj the electrostatic free energy for the charge-charge interaction, was obtained by least squares fitting to the experimental ionic strength data. A positive sign indicates a repulsive interaction and a negative sign an attractive interaction. Zlt the active-site charge on the cytochrome, was calculated using the fitted values of V il( Z2 = - 4, and assumed values for the radius of the interaction domain and the dielectric constant. Values in parentheses are those expected for the heme-edge region from the crystal structure and sequence information.

Data from Tollin et al.204

are available show that the surfaces of the proteins adjacent to the exposed part of the FMN have a high concentration of negatively charged side chains. The likelihood that electrostatic interactions are important in electron transfer to charged oxidants was confirmed by theoretical calculation,118-207 and by experiments in which electron transfer was measured from the semiquinones of several flavodoxins to a variety of other electron-transfer proteins, including cytochromes, blue copper proteins, and high potential ferredoxins,199'204-205'208"211 for which sufficient structural information is available to permit a reasonable estimate of the net charge surrounding the redox prosthetic group. The effects of ionic strength on the rate constants for electron transfer were consistent in all cases with the notion that negative charges on the flavodoxin interact with the charges on the oxidant (Figure 1, Table 5). The results of theoretical fits to the data for a group of otype cytochromes (Table 5) permit an extrapolation to infinite ionic strength to determine k^ values which should reflect intrinsic reactivities uninfluenced by electrostatic interactions, as well as allowing estimates to be made of the active site charges (Z,,^). Similar results have been obtained for the other groups of redox proteins. A decrease of rate constant with an increase of ionic strength implies that there is an attractive electrostatic interaction between the reactants, and the slope of the dependence is determined by the size of the charge product, Z,Z2, a reflection of the electrostatic free energy of interaction (V H ). If it is assumed tht the active site charge (Z2) or C. pasteurianum flavodoxin is -4, then the data for the reaction between this flavodoxin and cytochrome c indicate an active site charge (Z L ) of + 4.7 for the cytochrome. This value is in good agreement with the charge at the exposed heme edge that can be calculated from the crystal structure. It should be noted from Figure 3 that the electrostatic effects can be quite large. For example, the relative reactivities at an ionic strength of 50 mAf of two cytochromes

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Chemistry and Biochemistry of Flavoenzymes

with the same redox potential, horse cytochrome c and Pseudomonas aeruginosa cytochrome c551, with C. pasteurianum flavodoxin are 100:1. In addition, a change in ionic strength can cause electrostatic interactions to reverse the relative reactivities of two proteins with a flavodoxin. Thus at an ionic strength of 360 mM, P. aeruginosa cytochrome c551 reacts approximately four times faster than horse cytochrome c. When electrostatic calculations were made using the structure for C. beijerinckii MP to determine the free energy of association as a function of ionic strength,118-207 good agreement was found with the rate constant vs. ionic strength profiles observed experimentally with C, pasteuranium flavodoxin; subsequent work showed that the effects of ionic strength on the redox reactions of the two flavodoxins are the same.210 Simondsen et al.205 showed that the limiting first order rate constant for electron transfer from the semiquinone of C. pasteurianum flavodoxin to horse cytochrome c decreases with increasing ionic strength. Taken together, the data and the theoretical calculations suggest that both attractive and repulsive electrostatic interactions between the charged groups on the surfaces of the reacting proteins act to orientate the molecules prior to actual physical contact as they diffuse together, and that this preorientation facilitates the formation of a complex that is optimal for electron transfer. It is important to note, however, that this conclusion does not apply to all complexes between electron transfer proteins; the electrostatically stabilized complex of cytochrome c and cytochrome c peroxidase, for example, is not the most active in electron transfer.212 In such cases, other forces must ensure that the two proteins are orientated in the appropriate way for electron transfer. A further notable conclusion from the studies of the effects of ionic strength on the reactions with flavodoxin semiquinone is that although the sign of the charge on the oxidant that is calculated from the kinetic data is sometimes the same as the net charge on the protein, in many cases it is not. For example, the protein net charge for Paracoccus denitrificans cytochrome c2 is -7, but the active site charge that is calculated from the kinetic data with flavodoxin is +3.6 (Table 4). The difference is due to the way in which the charge on the cytochrome is distributed asymmetrically over the molecule; the positive charge is localized around the exposed edge of the heme, while the negative charge is largely on the opposite side of the molecule. In general it appears that the local charge close to the electron transfer site is more important than the overall charge, although when the local charge is small, the apparent charge observed in the kinetic experiments can be strongly influenced by the protein net charge.206'213 Electrostatic interactions might also be utilized as a control mechanism for electron transfer. Thus, Cheddar and Tollin214 have noted that binding of polylysine to C. pasteurianum flavodoxin modifies the reactivity towards small inorganic oxidants such that positively charged molecules react more slowly and negatively charged molecules react more rapidly. The effect can be quite large in some instances (up to 200-fold). Whether or not this effect is utilized biologically remains to be determined.

C. RELATION BETWEEN RATE CONSTANTS FOR ELECTRON TRANSFER AND REDOX POTENTIALS

Marcus electron transfer theory132 requires that the rate constants for the reaction between two redox moieties should depend on the difference between their redox potentials (i.e., the thermodynamic driving force), as well as on changes in structure and solvation that must occur prior to electron transfer to allow the transition state to be reached (often referred to as the reorganizational energy). Initial studies showed that the predicted relationship between the rate constant and the difference in midpoint redox potential holds for a wide range of heme, copper, and iron-sulfur proteins in their reactions with the semiquinone of proteinfree flavin.206-213 However, when similar comparisons were done with the semiquinone of C. pasteurianum flavodoxin, the patterns of reactivity were much more complex (e.g.,

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Figure 2). It is apparent from these data that the relatively tight relationships among the various structurally homologous groups of redox proteins found with lumiflavin semiquinone are only partly retained in the reactions with flavodoxin semiquinone. Furthermore, the range of values for the rate constants with flavodoxin are 1000 to 10,000 times greater than with lumiflavin. These differences suggest that the steric properties of the reaction site, and the reorganizational energy contribution, play a much greater role in electron transfer reactions between two proteins than between a protein and a relatively small molecule such as lumiflavin.206 Thus, when two proteins form a complex, interactions occur over a relatively large fraction of each of their surfaces, and they penetrate each other much less than a complex of protein and a small molecule. As a consequence, relatively small differences in surface topography, which are without effect on the interaction with lumiflavin, can have a large effect on the distance between the redox centers and their orientation in a complex with flavodoxin (see below). Additional factors which influence the reorganizational energy, such as a restricted access of solvent to the region between two proteins and/or restructuring of the solvation shells of the two molecules, may make a larger contribution to the energetics of protein-protein interactions than is the case in a complex of a protein with a small molecule. Regardless of the quantitative explanation for these striking differences, it is clear that flavodoxin semiquinone discriminates more readily than lumiflavin between members of an homologous series of redox proteins, an essential requirement for establishing biological specificity in protein-protein electron transfer. D. STERIC EFFECTS ON RATES OF ELECTRON TRANSFER In the present discussion, the term steric effects refers to kinetic factors that involve features of the surface topography of the protein which affect access of an incoming reactant to the redox center in the protein. The exposure to solvent of various atoms of the prosthetic group is one measure of such effects. Tollin et al.215 have shown that calculations of the solvent exposure of the redox centers of a variety of cytochromes, iron-sulfur proteins and blue copper proteins correlate well with their relative reactivities with the semiquinone of free flavin, but correlate poorly with their reactivities with flavodoxin semiquinones. In order to explain the reactivities with flavodoxins, it was necessary to consider more global features of the active site regions. For example, the relative reactivities of several structurally homologous c-type cytochromes that have similar redox potentials, with the semiquinone of C. pasteurianum flavodoxin (Table 6) can be correlated with amino acid side chains which project from the front surface of the cytochrome. When these are present, they interfere with the approach of flavodoxin, but they do not form significant barriers to the approach of lumiflavin or solvent (cf. References 203 and 216 for further discussion). A similar situation was found to exist for cytochrome c' as compared to cytochrome c (cf. Chlorobium cytochrome c555 vs. Alcaligenes cytochrome c' in Table 6). In this case, the barrier to flavodoxin is a deep cleft in the surface of cytochrome cf which restricts access of flavodoxin to the solvent-exposed heme edge.209 It is interesting that cytochrome cf is approximately three times as reactive towards lumiflavin semiquinone as cytochrome c (Figure 2); the cleft is large enough to allow access of the smaller lumiflavin molecule, and therefore the relative reactivities are dominated by the extent of solvent exposure of the heme itself, which is greater for cytochrome c' than for cytochrome c, and independent of the surrounding topography. Similar results were obtained when the reactivities of the semiquinones of free flavin and of flavodoxin with azurins, plastacyanins, and stellacyanin were compared.210 In this case, the reactivity of stellacyanin with lumiflavin was higher, and the reactivity with flavodoxin lower than the corresponding reactivities of the other two copper proteins (Figure 3). A crystal structure of stellacyanin is not available, but it can be predicted from the results of the kinetic measurements that the copper center in this protein lies in a cleft or depression, analogous to the one found in cytochrome c', which restricts the access of protein molecules

412

Chemistry and Biochemistry of Flavoenzymes TABLE 6 Second-Order Rate Constants, Extrapolated to Infinite Ionic Strength, for Electron Transfer from C. pasteurianum Flavodoxin Semiquinone to Various Cytochromes Cytochrome Horse c Tuna c Candida c Pseudomonas c551 R. rubrum c2 Chlorobium c555 Alcaligenes c'

Em7 (mV)

260 260 260 270 324 150 130

k^A/^s- 1 ) 0.2 0.9 1.5 11.5 20.8 0.3

x x x x x x

4.1

104 104 104 104 104 104

Ref. 204 204 204 204 204 204 209

but not of molecules as small as free flavins. This result emphasizes the value of using flavodoxin as a probe in the study of other electron transfer proteins. Cheddar et al. 211 have compared the reactivities of flavodoxins from different species with a variety of c-type cytochromes and with ferricyanide ion (Table 7). In all of the reactions studied, the semiquinone of flavodoxin from C. pasteurianum was more reactive than the semiquinone of flavodoxin from C. beijerinckii MP, and these reactivities are in the order predicted by their redox potentials. In contrast, flavodoxins from A. nidulans and from Az. vinelandii were less reactive than expected from thermodynamic considerations, and in these cases it is likely that the FMN is less accessible. This conclusion is consistent with the known differences in the crystal structures of flavodoxins from C. beijerinckii MP and A. nidulans110 and with computer graphic modeling which indicates that because of differences in the way in which FMN is oriented in their flavin binding sites, the two flavodoxins must orientate themselves differently in the electron transfer complex with cytochrome c.211 A crystal structure is not available for flavodoxin from Az. vinelandii, but the kinetic data suggest that the steric constraints at the flavin-binding site of this protein must be similar, though not identical, with those of flavodoxin from A. nidulans. The ionic strength dependence of the rate constant for electron transfer between flavodoxins from A. nidulans and Az. vinelandii and P. denitrificans cytochrome c551 was of opposite sign to the effects seen with the clostridial flavodoxins.211 This marked difference between the flavodoxins was seen only in their reactions with the cytochrome c551 from P. denitrificans. It is probably due to the fact that the surface of this cytochrome at which the heme edge is exposed is virtually devoid of charge, while the corresponding surfaces of all of the other cytochromes used have a high positive charge.207 It was argued that the steric constraints on complex formation with the A. nidulans and Az. vinelandii flavodoxins caused them to interact at sites of cytochrome c551 that were different from the sites at which the clostridial flavodoxins interact, and hence to experience a different charge. The electrostatic attraction of the positively charged regions on the other cytochromes for the negative surface of the flavodoxins is so dominant that steric effects are manifested mainly in the magnitude of the rate constant, rather than in the ionic strength effects. This emphasizes, in a quite direct way, the strong influence that surface topography can exert on the nature of proteinprotein interactions during electron transfer. A study by De Francesco et al.199 on the effects of substituting the Sa-position of FMN of flavodoxins from C. pasteurianum and Az. vinelandii with the relatively bulky imidazole function provides a further direct demonstration of the influence of steric factors. The crystal structures and other studies have indicated that the 8a-position of FMN in these flavodoxins is highly exposed, so that the position can be substituted with bulky groups such as imidazole and the resulting flavin can still be accommodated by the protein.195 The substitution caused

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FIGURE 2. Ionic strength dependence of the second order rate constants for electron transfer from C. pasteurianum flavodoxin semiquinone to oxidized c-type cytochromes. Solid lines are theoretical fits to data points using an electrostatic model as described in Tollin et al.204 (A) Rsp. tenue 3761 r553; (B) Euglena sp. c552; (C) Rm. vanniellic2; (D)Rsp. capsulata c2; (E)Rsp. rubrum c2; (F) Ps. aeruginosa c551; (G) P. denitrificans c2; (H) Candida krusei c, (I) tuna c, (J) horse c, (K) Chi. thiosulfatophilum r555, (L) E. halophila c551. (Data from Tollin et al.204)

a 10 to 30-fold decrease in the second-order rate constant and limiting first-order rate constant for electron transfer (Table 3). Such decreases cannot be explained by a change in redox potential of the bound flavin, and it is likely that the effect is due to a steric effect of the imidazole substituent and a consequent increase in the distance between the flavin and heme. If correct, the explanation suggests that the imidazole ring cannot function as a bridge for

414

Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. Left:Semilog plots of second order rate constants for electron transfer from lumiflavin semiquinone to various oxidized redox proteins vs. difference in midpoint potential between reactants. Each point corresponds to an individual protein (for identification of specific proteins, see Meyer et al., 216 Meyer et al.,209 Przysiecki et al.,208 Tollin et al.206). (o) cytochromes c ; (•) cytochromes c\ (X) high potential ferredoxins; (D) blue copper proteins (Pc = plastocyanin; Az = azurin; NR = nitrite reductase; La = laccase; St = stellacyanin). Curves are theoretical plots using the Marcus exponential equation (see Tollin et al.206). Right: Semilog plot of electrostatically corrected second order rate constants for electron transfer from C. pasteurianum flavodoxin semiquinone to various oxidized redox proteins. Symbols as in left-hand plot. (A) Euglena c552; (B) Rhodospirillum c553; (C) Candida c\ (D) tuna c; (E) horse c. Curves are theoretical plots using the Marcus exponential equation; steeper curves, which would fit more of the data points, can only be obtained using physically unreasonable parameters. (Data from Tollin et al.,204 Przysiecki et al.,208 Meyer et al.,209 and Tollin etal., 206 .)

TABLE 7 Relative Reactivities of Various Flavodoxins Towards c-Type Cytochromes Flavodoxin Anacystis Azotobacter C. pasteurianum C, beijerinckii MP Anacystis Azotobacter C. pasteurianum C. beijerinckii MP

Cytochrome Tuna c Tuna c Tuna c Tuna c Pseudomonas Pseudomonas Pseudomonas Pseudomonas

c551 c55 1 c551 c551

k. (M~l s'1)

0.74 1.3 8.7 4.6 1.2 0.31 1.0 0.35

x x x x x x x x

103 103 103 103 103 103 105 105

Data from Cheddar et al.211

electron transfer between the flavin and heme. This might have implications for those enzymes which contain covalently bound flavin that is linked to the protein through an 8a-histidyl linkage. It was also observed199 that protonation of the imidazole ring of 8-IMF caused a further

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FIGURE 4. Stereo view of a computer-modeled electron transfer complex between tuna cytochrome c (upper molecule) and C. beijerinckii MP flavodoxin (lower molecule). Only those side-chain residues involved in intermolecular charge pairs (dashed lines) are shown.

20- to 50-fold decrease in the second order rate constant. It was suggested that this effect could be due to an energetically unfavorable interaction between the cytochrome surface and the positively charged imidazolium ring, in keeping with the relatively large electrostatic effects note above. E. STRUCTURAL ASPECTS OF INTERMEDIATE PROTEIN-PROTEIN COMPLEX There is much kinetic and other evidence that 1:1 complexes of redox proteins form during catalysis by redox enzymes that transfer electrons to or from other proteins (e.g., ferredoxin and ferredoxin-NADP+ reductase; adrenodoxin and adrenodoxin reductase; cytochrome P450 and cytochrome P450 reductase, cytochrome c and cytochrome c peroxidase). However, although an X-ray analysis of a disordered crystal of the cytochrome c:cytochrome c peroxidase complex has been reported,217 a crystal structure of such a complex is not yet available, and it has not been possible to directly relate the kinetic results to structural features in the complex. Nevertheless, it has been possible to obtain likely structures for the complexes by using computer graphics techniques and the known structures of the two component proteins in the complex. The first model of this kind was made for the complex of cytochrome c and cytochrome b5,218 and subsequent models used the principles established with this complex by Salemme.219 These can be summarized as follows: the complex is stabilized primarily by electrostatic forces; prosthetic groups in the two partners are as far as possible oriented with their macrocyclic rings parallel, and they are as close together as possible; water molecules are excluded from the area of interaction. Simondsen et al.205 used the principles outlined above and known crystal structures to construct a model of a 1:1 complex of C. beijerinckii MP flavodoxin and tuna cytochrome c, and they used the model to explain the results of their kinetic measurements on the electron transfer between C. pasteurianum flavodoxin semiquinone and oxidized horse cytochrome c. A stereo view of this complex is shown in Figure 4. The molecules are oriented so that the solvent-exposed parts of the flavin and heme are facing the interface between the molecules; the edge of the redox centers are in van der Waals contact, with the heme and flavin planes inclined to each other at approximately 30°. Hydrophobic contacts are formed

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Chemistry and Biochemistry of Flavoenzymes

between residues of a polypeptide loop (residues 7 to 10) on flavodoxin and residues in a depression on cytochrome c into which the loop fits, and salt links are formed between the side chains of lysine residues 13, 25, 27, and 79 on cytochrome c and the side chains of glutamate residues 120, 65, and 62, and aspartate residue 58 on flavodoxin. The charge interactions are consistent with the effects of ionic strength on the reaction kinetics, although, as noted above, such electrostatic interactions may not always be necessary for optimal electron transfer in complexes, and they are also consistent with the results of theoretical calculations118-205-207 and with the results of cross-linking studies.220 The overall structure of the complex is compatible with the observed effects on the kinetics of modifying the structure of the dimethylbenzene ring of FMN (see above).181'199 An analogous computer graphics model has been generated for the complex of flavodoxin and cytochromre c3 from D. vulgaris.221 Five salt links were found in this complex, involving acidic residues on the flavodoxin and lysines on the cytochrome, and again the FMN and one of the four heme prosthetic groups were nearly coplanar and in van der Waals contact. Complexes have also been demonstrated for more physiological complexes of flavodoxin, e.g., flavodoxin and ferredoxin-NADP+ reductase from Anabaena PCC 7119.222 In this latter case, a covalently cross-linked complex has been generated from the electrostatically stabilized complex. Although the covalent complex has biological activity, kinetic studies have shown that the rate constant for intracomplex electron transfer between the two proteins is significantly diminished compared to that for the electrostatic complex.223 This illustrates the danger of using covalent cross-linking procedures to stabilize protein-protein electron transfer complexes. Az. vinelandii flavodoxin has been found to contain an as yet unidentified "labile" phosphorus-containing residue, which can be removed from the protein by acid treatment or by denaturants such as urea or guanidinium chloride,91 and whose presence is dependent on the growth phase of the organism.92 The presence of this group does not affect to oneelectron redox potential of the flavodoxin. However, a major difference between the two types of flavodoxin is that the FMN phosphorus NMR signal is broadened beyond detection upon flavin semiquinone formation for the "phospho" form,91 but not for the "dephospho" form.224 Complex formation of "dephospho" oxidized flavodoxin with spinach ferredoxin-NADP+ reductase produces no perturbation of the 31P resonances from the FMN or FAD phosphates of the two proteins, or of the covalent, disubstituted, phosphate resonance of the flavodoxin. Formation of the flavodoxin semiquinone within the complex results in extensive broadening of the FMN phosphate resonance, in contrast to the results with the free "dephospho" flavodoxin.224 These observations suggest a perturbation of the flavin ring-side chain configuration of the "dephospho" flavodoxin semiquinone on binding to the spinach reductase, in which the side chain phosphate is in closer proximity to the isoalloxazine ring in the complex than when free in solution. Difference absorption spectra of complexes of "phospho" and "dephospho" Az. vinelandii flavodoxin with the ferredoxin reductase from Anabaena variabilis show that both forms are bound in a 1:1 stoichiometry with similar binding affinities.233 However, the spectral perturbations (which reflect the flavin cofactors of either or both proteins) are not the same, and thus there are probably differences in the binding geometries in the two complexes. Difference CD spectra are also consistent with these results. NMR studies of the complex of C. pasteurianum flavodoxin and horse cytochrome c showed that there is a perturbation of the heme resonances in the complex, and that the perturbation is greater than would be anticipated from a change in the rotational relaxation.225 The observations indicate that structural changes occur within the heme crevice. Large changes also occur in the circular dichroism of the Soret region of the heme spectrum when the complex is formed,226 and these changes are also consistent with a perturbation of the

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heme environment, possibly due to a loosening of the heme crevice and uncoupling of the dipole interactions between segments of the polypeptide chain and the heme. Neither of these spectroscopic changes occur to an appreciable extent in the complex of Az. vinelandii flavodoxin and horse cytochrome c, and the heme CD effects were largely eliminated in the complex with C. pasteurianum flavodoxin, when 8-IMF was substituted for FMN. Since both of these complexes have low catalytic activity, it is possible that the heme perturbation is necessary for rapid electron transfer. It may be significant that perturbations of the CD spectrum also occur in complexes between ferredoxin and its reductase,227 and between cytochrome c and its oxidase,228 and for both cytochrome c oxidase229'230 and cytochrome c peroxidase,231 reduction is facilitated by formation of the complex with cytochrome c. It is possible, therefore, that structural modifications are a general consequence of proteinprotein interactions, and are an important feature in electron transfer between redox proteins. Dynamic motion at the interface of the two proteins in the complex may also be important in electron transfer. Evidence has been given that the heme and flavin in the complex of horse cytochrome c and C. pasteurianum flavodoxin are more open for reaction with external reductants that would be expected from the structure of the computer graphics model.232 The kinetics of reduction of the cytochrome component by the flavin semiquinone are nonlinear; this was attributed to dynamic processes that have first order rate constants of 1000 to 3000 s" 1 at low ionic strength, and which open up the interface region to allow greater access to the redox groups. Such dynamic changes may be important in multistep mechanisms which involve complexes of the type described here. It was shown in the same study that the small redox protein rubredoxin is able to reduce cytochrome c at much the same rate whether or not the cytochrome is complexed to flavodoxin, again implying that dynamic processes play an important role in the mechanism. F. SPECIFICITY OF PROTEIN-PROTEIN ELECTRON TRANSFER REACTIONS It is essential in biological electron transfer processes that reactions occur with a high degree of specificity, so that undesirable redox events which are favorable from a thermodynamic point of view do not occur. The three principal factors which influence the reaction kinetics of protein-protein electron transfer identified in the studies outlined above on electron transfer from flavodoxin semiquinones, are thermodynamic driving force, protein surface topography, and electrostatic interactions. Each of these factors is capable of producing a 1000-fold variation in rate constant, and therefore kinetic variations of up to nine orders of magnitude can occur in a family of structurally related redox proteins by changing the nature of the amino acid side chains which determine these parameters. Evolution uses this mechanism to optimize the driving force, the steric and electrostatic interactions for a particular electron transfer process, and in this way ensures that a physiological function can proceed with high specificity and efficiency.

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34. Yoch, D. C., The electron transport system in nitrogen fixation by Azotobacter. IV. Some oxidationreduction properties of azotoflavin, Biochem. Biophys. Res. Commun., 49, 335, 1972. 35. Watt, G. D., An electrochemical method for measuring redox potentials of low potential proteins by microcoulometry at controlled potentials, Anal. Biochem., 99, 399, 1979. 36. Klugkist, J., Voorberg, J., Haaker, H., and Veeger, C., Characterization of three different flavodoxins from Azotobacter vinelandii, Eur. J, Biochem., 155, 33, 1986. 37. Cusanovich, M. A. and Edmondson, D. E., Isolation and characterization of Rhodospirillum rubrwn flavodoxin, Biochem. Biophys. Res. Commun., 45, 327, 1971. 38. Macknight, M. L., Gray, W. R., and Tollin, G., N-terminal amino acid sequences of Azotobacter vinelandii and Rhodospirillum rubrum flavodoxins, Biochim. Biophys, Ada, 59, 630, 637. 39. Crespi, H. L., Smith, C., Gajda, L., Tisue, T., and Ameraal, R. M., Extraction and purification of 'H, 2H and isotope hybrid algal cytochrome, ferredoxin and flavodoxin, Biochim. Biophys. Acta, 256, 611, 1972. 40. Crespi, H. L., Norris, J. R., Bays, J. P., and Katz, J. J., ESR and NMR studies with deuterated flavodoxin, Ann. N.Y. Acad. Sci., 222, 800, 1973. 41. Smillie, R. M. and Entsch, B., Phytoflavin, in Methods Enzymol., 23, 504, 1971. 42. Entsch, B. and Smillie, R. M., Oxidation-reduction properties of phytoflavin, a flavoprotein from bluegreen algae, Arch. Biochem. Biophys., 151, 378, 1972. 43. Bothe, H., Hemmerich, P., and Sund, H., Some properties of phytoflavin isolated from the blue-green algae Anacystic nidulans, in Flavins and Flavoproteins, Kamin, H.,Ed., University Park Press, Baltimore, 1971, 211. 44. Sykes, G. A. and Rogers, L. J., Redox potentials of algal and cyanobacterial flavodoxins, Biochem. J., 217, 845, 1984. 45. Hutber, G. N., Hutson, K. G., and Rogers, L. J., Effect of iron deficiency on levels of two ferredoxins and flavodoxin in a cyanobacterium, FEMS Microbiol. Lett., 1, 193, 1977. 46. Hutber, G. N., Smith, A. J., and Rogers, L. J., Flavodoxin from the blue-green algae Nostoc strain MAC, Phytochemistry, 20, 383, 1981. 47. Filial, M. F., Sandmann, G., and Gomez-Moreno, C., Flavodoxin from the nitrogen-fixing cyanobacterium Anabaena PCC 7119, Arch, Microbiol, 150, 160, 1988. 48. Fillat, M. F., Edmondson, D. E., and Gomez-Moreno, C., Structural and chemical properties of a flavodoxin from Anabaena PCC 7119, Biochem. Biophys. Res. Commun., 1040, 301, 1990. 49. Paulsen, K. E., Stankovich, M. T., Stockman, B. J., and Markley, J. L., Redox and spectral properties of flavodoxin from Anabaena 7120, Arch. Biochem. Biophys., 280, 68, 1990. 50. Fitzgerald, M. P., Husain, A., and Rogers, L. J., A constitutive flavodoxin from a eukaryotic alga, Biochem. Biophys. Res. Commun., 81, 630, 1978. 51. Zumft, W. G. and Spiller, H., Characterization of a flavodoxin from the green alga Chlorella, Biochem. Biophys. Res. Commun., 45, 112, 1971. 52. Korsunskii, O. F. and Smolygina, L. D., Properties of low potential electron exchangers isolated from the nitrogen-fixing purple sulfur bacteria Thiocapsa roseopersicina, in Biol. Fiksatsiya Mol, Azota, Kertovich, V. L., Ed., Naukova Dumka, Kiev, 1983, 219. 53. Yakunin, A. F., Lauvinaviciene, T., and Kulakova, S. M., Electron transporters and enzymes interacting with nitrogenase and hydrogenase in nitrogen-fixing phototrophs, in Biol. Fiksatsiya Mol. Azota, Kretovich, V. L., Ed., Naukova Dumka, Kiev, 1983, 227. 54. Sandman, G. and Malkin, R., Iron-sulfur centers and activities of the photosynthetic electron transfer chain in iron-deficient cultures of the blue-green alga Aphanocapsa, Plant. Physiol., 73, 724, 1983. 55. Pshenova, K. V., Mutishin, A. A., Shatilov, V. R., and Sofin, A. V., Isolation of proteins of the photosynthetic electron transport chain bound with photosystem I from Ankistrodesmus braunii, Prikl. Biokhim. Mikrobiol.t 17, 773, 1981. 56. Cohn, C. L., Alam, J., and Krogmann, D. W., Multiple ferredoxins from cyanobacteria, Physiol. Veg>, 23, 659, 1985. 57. Chen, J.-S., Flavodoxin of an anaerobic bacterium Fusobacterium nucleatum sensitive to metronidazole but not producing hydrogen, Fed. Proc. Fed. Am. Soc. Exp. Biol., 45, 1890, 1986. 58. Peck, H. D., Jr. and LeGall, J., Biochemistry of dissimilatory sulfate reduction, Phil. Trans. R. Soc. Lond. Ser. B, 298, 443, 1982. 59. Curley, G. P., personal communication. 60. Laud en bach, D. E., Reither, M. E., and Straus, N. A., Isolation, sequence analysis and transcriptional studies of the flavodoxin gene from Anacystic nidulans R2 J. BacterioL, 170, 258, 1988. 61. Fujii, K. and Huennekens, F. M., Activation of methionine synthetase by a reduced triphosphopyridine nucleotide-dependent flavoprotein system, J. Biol. Chem., 249, 6745, 1974. 62. Fujii, K., Galivan, J. H., and Huennekens, F. M., Activation of methionine synthetase: a further characterization of the flavoprotein system, Arch. Biochem. Biophys., 178, 662, 1977.

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63. Kim, J.-H. and Akagi, J. M., Characterization of a trithionate reductase system from Desulfovibrio vulgaris, J . Bacteriol., 163, 472, 1985. 64. Ogata, M. and Akagi, T., Pyruvate dehydrogenase and the path of lactate degradation in Desulfovibrio vulgaris Miyazaki K , J. Biochem., 100, 311, 1986. 65. Nieva-Gomez, D., Roberts, G. P., Klevickis, S., and Brill, W. J., Electron transport to nitrogenase in Klebsiellapneumoniae, Proc, Natl. Acad. Sci. U.S.A., 77, 2555, 1980. 66. Shah, V. K., Stacey, G., and Brill, W. J., Electron transport to nitrogenase. Purification and characterization of pyruvate:flavodoxin oxidoreductase, the nifl gene product, J. Biol. Chem., 258, 12064, 1983. 67. Beynon, J., Cannon, M., Buchanan-Wollaston, V., and Cannon, F., The nif promoters of Klebsiella pneumoniae have a characteristic primary structure, Cell, 34, 6654, 1983. 68. Ben net, L. T., Jacobson, M., and Dean, D. R., Isolation, sequencing and mutagenesis of the nif F gene encoding flavodoxin from Azotobacter vinelandii, J. Biol. Chem., 263, 1364, 1988. 69. Haaker, H., de Kok, A., and Veeger, C., Regulation of dinitrogen fixation in intact Azotobacter vinelandii, Biochim. Biophys. Acta, 357, 344, 1974. 70. Laane, C., Krone, W., Konings, W., Haaker, H., and Veeger, C., Short term effect of ammonium chloride on nitrogen fixation by Azotobacter vinelandii and by bacteroids of Rhizobium leguminosa, Eur. J. Biochem., 103, 39, 1980. 71. Klugkist, J., Haaker, H., and Veeger, C., Studies on the mechanism of electron transport to nitrogenase in Azotobacter vinelandii, Eur. J. Biochem., 155, 41, 1986. 72. Haaker, H. and Veeger, C., Involvement of the cytoplasmic membrane in nitrogen fixation by Azotobacter vinelandii, Eur. J. Biochem., 77, 1, 1977. 73. Thauer, R. K., Jungermann, K., Rupprecht, E., and Decker, K., Hydrogen formation from NADH in cell-free extracts of Clostridium kluyveri. Acetyl CoA requirement and ferredoxin dependence, FEBS Lett., 4, 108, 1969. 74. Jungermann, K., Rupprecht, E., Ohrloff, C., Thauer, R. K., and Decker, K., Regulation of the reduced nicotinamide adenine dinucleotide-ferredoxin reductase system in Clostridium kluyveri, J. Biol. Chem., 246, 960, 1971. 75. Thauer, R. K., Rupprecht, E., Ohrloff, C., Jungermann, K., and Decker, K., Regulation of the reduced nicotinamide adenine dinucleotide phosphate-ferredoxin reductase system in Clostridium kluyveri, J. Biol. Chem., 246, 954, 1971. 76. Mayhew, S. G., unpublished results. 77. Ragsdale, S. W. and Wood, H. G., Acetate biosynthesis by acetogenic bacteria. Evidence that carbon monoxide dehydrogenase is the condensing enzyme that catalyzes the final steps of synthesis, J. Biol. Chem., 260,3970, 1985. 78. Ragsdale, S. W., Ljungdahl, L. G., and DerVartanian, D. V., Isolation of carbon monoxide dehydrogenase from Acetobacterium woodii and comparison of its properties with those of the Clostridium thermoaceticum enzyme, J. Bacteriol., 155, 1224, 1983. 79. Knappe, J. and Blasckowski, H. P., Pyruvate formate-lyase from Escherichia coll and its activation system in Methods Enzymoi., 61, 508, 1975. 80. Blasckowski, H. P., Neuer, G., Ludwig-Festl, M., and Knappe, J., Routes of flavodoxin and ferredoxin reduction in Escherichia coli. CoA-acylating pyruvate:flavodoxin and NADPH:flavodoxin oxidoreductases participating in the activation of pyruvate formate-lyase, Eur. J. Biochem., 123, 563, 1982. 81. Yoshino, M., Murakami, K., and Tsushima, K., Flavodoxin: An allosteric inhibitor of AMP nucleosidase from Azotobacter vinelandii, J. Biochem. (Tokyo), 80, 839, 1976. 82. Dutton, J. E. and Rogers, L. J., Isoelectric focusing of ferredoxins, flavodoxins and rubredoxin, Biochim. Biophys. Acta, 537, 501, 1978. 83. Mayhew, S. G. and Howell, L. G., Chromatography of proteins on diethylaminoethyl cellulose in concentrated ammonium sulphate, Anal. Biochem., 41, 466, 1971. 84. Sakihama, N., Kitagawa, Y., Kitazume, Y., and Shin, M., Application of Toyopearls to purification of Azotobacter flavodoxin, Agric. Biol. Chem., 47, 2917, 1983. 85. Chen, J.-S. and Blanchard, D. K., A simple hydrogenase-linked assay for ferredoxin and flavodoxin, Anal. Biochem., 93, 216, 1979. 86. Blusson, H, Petitdemange, H., and Gay, R., A new fast and sensitive assay for NADH-ferredoxin oxidoreductase in clostridia, Anal. Biochem., 110, 176, 1981. 87. Deistung, J., Cannon, F. C., Cannon, M. C., Hill, S., and Thorneley, R. N. F., Electron transfer to nitrogenase in Klebsiella pneumoniae; nifF gene cloned and the gene product flavodoxin, purified, Biochem. J., 231,743, 1985. 88. Hill, S. and Kavanagh, E. P., Roles of nifF and nif J gene products in electron transport to nitrogenase in Klebsiella pneumoniae, J. Bacteriol., 141, 470, 1980. 89. Yoch, D. C., Dimerization of Azotobacter vinelandii flavodoxin (azotoflavin), Arch. Biochem. Biophys., 170, 326, 1975.

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90. Edmondson, D. E. and James, J. L., Covalently bound non-coenzyme phosphorus residues in flavoproteins. M P NMR studies on Azotobacter flavodoxin, Proc. Natl. Acad. Sci. U.S.A., 76, 3786, 1979. 91. Edmondson, D. E. and James, T L., Physical and chemical studies on the FMN and non-flavin phosphate residues in Azotobacter flavodoxin, in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Jr., Eds., Elsevier, New York, 1982, 111. 92. Edmondson, D. E. and Peleato, M. L., Properties of protein-bound phosphorus groups in flavodoxin from Azotobacter vinelandii, in Inorganic Nitrogen Metabolism, Ullrich, N. R., Aparicio, P. J., Syrett, P. J., and Castillo, R, Eds., Springer Verlag, Berlin, 1987, 187. 93. BovIan, M. H. and Edmondson, D. E., Studies on the incorporation of a covalently bound disubstituted phosphate residue into Azotobacter vinelandii flavodoxin in vivo, Biochem. J., 268, 745, 1990. 94. Hofstetter, W. and DerVartanian, D. 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117. Fukuyama, K., Wakabayashi, S., Matsubara, H., and Rogers, L. J., Tertiary structure of oxidized flavodoxin from an eukaryotic red alga Chondrus crispus at 2.35 A resolution. Localization of charged residues and implication for interaction with electron transfer partners, /. BioL Chem., 265, 15804, 1990. 1 1 8 . Matthew, J. B., Weber, P. C., Salem me, F. R., and Richards, F. M., Electrostatic orientation during electron transfer between flavodoxin and cytochrome, c, Nature (London), 301, 169, 1983. 119. Armstrong, F. A., Hill, H. A. O., and Walton, N. J., Reactions of electron-transfer proteins at electrodes, Q, Rev. Biophysics, 18, 261, 1985. 120. Faraggi, M. and Klapper, M. H., One electron reduction of flavodoxin, a fast kinetic study, /. Biol. Chem., 254, 8139, 1979. 121. Mayhew, S. G. and van Arem, E. J. F., Studies on the redox properties of flavodoxin and flavodoxin derivatives, Abstr, 10th Inter, Congr. Biochem., 236, 1976. 122. Anderson, R. F., Massey, V., and Schopfer, L. M., Pulse radiolysis studies on flavodoxin, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 279. 123. Massey, V. and Hemmerich, P., Photoreduction of flavoproteins and other biological compounds catalyzed by deazaflavins, Biochemistry, 17, 9, 1978. 124. Mayhew, S. G. and Massey, V., Photochemical formation of a stable red derivative of flavodoxin, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 261. 125 Muller, F., Vervoort, J., van Mierlo, C. P. M., Mayhew, S. G., van Berkel, W. J. H., and Bacher, A., C-13, N-15 and two-dimensional NMR techniques in flavoprotein research, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 279. 126. Mayhew, S. G. and Massey, V., Studies on the kinetics and mechanism of reduction of flavodoxin from Peptostreptococcus elsdenii by sodium dithionite, Biochim. Biophys. Acta, 315, 181, 1973. 127. Lambeth, D. O. and Palmer, G., The kinetics and mechanism of reduction of electron transfer proteins and other compounds of biological interest by sodium dithionite, J. Biol. Chem., 248, 6095, 1973. 128. Mayhew, S. G., The redox potential of dithionite and SO 2 ~ from equilibrium reactions with flavodoxins, methyl viologen and hydrogen plus hydrogenase, Eur. J. Biochem., 85, 535, 1978. 129. Mayhew, S. G., van Arem, E. T. F., Strating, M. J. J., and Wassink, J. H., The effects of pH on dithionite-reduced flavodoxin from Peptostreptococcus elsdenii and the use of apoflavodoxin to determine and purify FMN, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 411. 130. Mayhew, S. 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144. Carr, M. C., Curley, G. P., Mayhew, S. G., and Voordouw, G., Effects of substituting asparagine for glycine-61 in flavodoxin from Desulfovibrio vulgaris (Hildenborough), Int. Biochem., 20, 1025, 1990. 145. Swenson, R. P., Krey, G., and Eren, M., Site-directed mutagenesis of the flavin-binding site of bacterial flavodoxins, J. Cell. BioL, 107, 621a, 1989. 146. Kierkegaard, P., Norrestam, R., Werner, P.-E., Csoregh, I., Van Glehn, M., Karlsson, R,, Leijonmarck, M., Ronnquist, O., Stensland, B., Tillberg, O., and Torbjornsson, C., X-ray structure investigation of flavin derivatives, in Flavins and Flavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 1. 147. Tauscher, L., Ghisla, S., and Hemmerich, P., NMR study of nitrogen inversion and conformation of 1,5-dihydro-isoalloxazines ("reduced flavin")* Helv. Chim. Acta, 56, 630, 1973. 148. Ludwig, M. L., Burnett, R. M., Darling, G. D., Jordan, S. J., Kendall, D. S., and Smith, W. 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170. Madigan, R. and Mayhew, S. G., 1990, unpublished results. 171. Carlson, R. and Langermann, N., The thermodynamics of flavin binding to the apoflavodoxin from Azotobacter vinelandii, Archiv. Biochem. Biophys., 229, 440, 1984. 172. Pompom, D., Guiard, B., and Lederer, F., Reconstitution of liver HADH:cytochrome b5 oxidoreductase and of Desulfovibrio vulgaris flavodoxin with 1-carba-l-deazaflavin, Eur. J. Biochem., 128, 377, 1982. 173. Entsch, B., Husain, M., Ballou, D. P., Massey, V., and Walsh, C., Oxygen reactivity of/>-hydroxybenzoate hydroxylase containing 1-deaza-FAD, J. Biol. Chem., 255, 1420, 1980. 174. Claiborne, A., Massey, V., Fitzpatrick, P. F., and Schopfer, L. M., 2-Thioflavins as active site probes of flavoproteins, 7. Biol. Chem,, 257, 174, 1982. 175. Choong, Y. S. and Massey, V., 2-Thioriboflavins 5'-phosphate (2-thioFMN) lactate oxidase, Eur. J. Biochem., 131, 501, 1983. 176. Massey, V., Claiborne, A., Biemann, M., and Ghisla, S., 4-Thioflavins as active site probes of flavoproteins; general properties, J. Biol. Chem., 259, 9667, 1984. 177. Scola-Nagelschneider, G., Brustlein, M., and Hemmerich, P., 4a-alkyldihydroflavin: coenzyme synthesis and modification of flavodoxin, Eur. J. Biochem., 69, 305, 1976. 178. Edmondson, D. E. and Tollin, G., On the importance of the N-5 position in flavin coenzymes. Properties of free and protein-bound 5-deaza analogs, Biochemistry, 11, 1133, 1972. 179. Walsh, C., Flavin coenzymes: at the crossroads of biological redox chemistry, Ace. Chem. Res., 13, 148, 1980. 180. Fenner, H., Grauert, R., Hemmerich, P., Michel, H., and Massey, V., 5-Thia-5-deazaflavin, a le transferring flavin analog, Eur. J. Biochem., 95, 183, 1979. 181. Simondsen, R. P. and Tollin, G., Transient kinetics of redox reactions of flavodoxin: effects of chemical modification of the flavin mononucleotide prosthetic group on the dynamics of intermediate complex formation and electron transfer, Biochemistry, 22, 3008, 1983. 182. Mayhew, S. G., Whitfield, C. D., Ghisla, S., and Jorns, M., Identification and properties of new flavins in electron-transferring flavoprotein from Peptostreptococcus elsdenii and pig liver glycolate oxidase, Eur. J. Biochem., 44, 579, 1974. 183. Ghisla, S., Massey, V., and Yagi, K., Preparation and some properties of 6-substituted flavins as activesite probes for flavin enzymes, Biochemistry, 25, 3282, 1986. 184. Massey, V., Ghisla, S., and Yagi, K., 6-Azido- and 6-aminoflavins as active-site probes of flavin enzymes, Biochemistry, 25, 8089, 1986. 185. Massey, V., Ghisla, S., and Yagi, K., 6-Thiocyanoflavins and 6-mercaptoflavins as active-site probes of flavoproteins, Biochemistry, 25, 8103, 1986. 186. Shiga, K. and Tollin, G., Studies on the mechanism of electron transfer in flavodoxins, in Flavins and Flavoproteins, Singer, T. P., Eds., Elsevier, Amsterdam, 1976, 422. 187. Jung, J. and Tollin, G., Transient kinetics of electron-transfer reactions of flavodoxins, Biochemistry, 20, 5124, 1981. 188. Ghisla, S. and Mayhew, S. G., Identification and properties of 8-hydroxy flavin adenine dinucleotide in electron transferring flavoprotein from Peptostreptococcus elsdenii, Eur. J. Biochem., 63, 373, 1976. 189. Ghisla, S., Massey, V., and Mayhew, S. G., Studies on the active centers of flavoproteins: binding of 8-hydroxy-FAD and 8-hydroxy FMN to apoproteins, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 334. 190. Ghisla, S. and Mayhew, S. G., Isolation, synthesis and properties of 8-hydroxyflavins, Methods Enzymol., 66, 241, 1980. 191. Massey, V., Ghisla, S., and Moore, E. G., 8-Mercaptoflavins as active site probes of flavoenzymes, J. Biol. Chem., 254, 9640, 1979. 192. Schopfer, L. M., Massey, V., and Claiborne, A., Active site probes of flavoproteins. Determination of the solvent accessibility of the flavin position 8 for a series of flavoproteins, J. Biol. Chem., 256, 7329, 1981. 193. Ghisla, S., Fitzpatrick, P. F., and Massey, V., 8-Azidoflavins: Photoaffinity labels for flavoproteins, in Flavins and Flavoproteins, Bray, R. C., Engel, R C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 751. 194. Fitzpatrick, P. F., Ghisla, S., and Massey, V., 8-Azidoflavins as photoaffinity labels for flavoproteins, J, Biol. Chem., 260, 8483, 1985. 195. Oestreicher, G., Edmondson, D. E., and Singer, T. P., Binding of 8a-substituted flavins and flavin peptides to Azotobacter flavodoxin, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 447. 196. Shiga, K., Tollin, G., Falk, M. C., and McCormick, D. B., Binding and oxidation-reduction of monoamine oxidase-type 8ot-(S-peptidyl)flavins with Azotobacter (Shethna) flavodoxin, Biochem. Biophys. Res. Commun., 66, 227, 1975.

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197. Edmondson, D. E. and De Francesco, R., 8a-imidazoylflavins: Application of their redox and spectral properties to flavoenzyme systems, in flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 653. 198. De Francesco, R., Tollin, G., Brown, K., and Edmondson, D. E., Factors influencing the rate of electron transfer from flavodoxin to cytochrome c, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 305. 199. De Francesco, R., Tollin, G., and Edmondson, D. E., Influence of 8a-imidazole substitution of the FMN cofactor on the rate of electron transfer from the neutral semiquinones of two flavodoxins to cytochrome, c, Biochemistry, 26, 5036, 1987. 200. Harzer, G., Rokos, H., Otto, M. K., Bacher, A., and Ghisla, S., Biosynthesis of riboflavin. 6,7Dimemyl-8-ribityllumazine-5 '-phosphate is not a substrate for riboflavin synthetase, Biochim. Biophys, Acta, 540, 48, 1978. 201. 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218. Salem me, F. R., An hypothetical structure for an intermolecular electron transfer complex of cytochromes c and b 5J J. Mol. Bioi, 102, 563, 1976. 219. Salemme, F. R., Structure and function of cytochrome c, Ann. Rev. Biochem., 46, 299, 1978. 220. Dickerson, J. L., Kornuc, J. J., and Rees, D. C., Complex formation between flavodoxin and cytochrome c. Cross-linking studies, J, Biol Chem., 260, 5175, 1985. 221. Stewart, D. E., Le Gall, J., Peck, H. D., Jr., Wampler, J. E., Moura, L, Moura, J. J. G., Xavier, A. V., and Weiner, P. K., A computer graphics model of the complex formed between flavodoxin and the tetraheme cytochrome c3, in Flavins and Flavoproteins, Edmondson, D. E., and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 311. 222. Gomez-Moreno, C., Sancho, J., Fillat, M., Pueyo, J. J., and Edmondson, D. E., Complex formation between ferredoxin-NADP+-oxidoreductase and flavodoxin, in Flavins and Flavoproteins, Edmondson, D. E., and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 335. 223. Walker, M. C., Pueyo, J. J., Gomez-Moreno, C., and Tollin, G., Comparison of the kinetics of reduction and intramolecular electron transfer in electrostatic and covalent complexes of ferredoxin-NADP+ reductase and flavodoxin from Anabaena PCC 7119, Arch. Biochem. Biophys., 2881, 76, 1990. 224. Bonants, P. J. M., Miiller, F., Vervoort, J., and Edmondson, D. E., A 31P-nuclear-magnetic-resonance study of NADPH-cytochrome -P-450 reductase and of the Azotobacter flavodoxin/ferredoxin-NADP+ reductase complex, Eur. J. Biochem., 190, 531, 1990. 225. Hazzard, J. T. and Tollin, G., Proton NMR study of the cytochrome c: flavodoxin electron transfer complex, Biochem. Biophys. Res. Commun., 130, 1281, 1985. 226. Tollin, G., Brown, K., Francesco, R. D., and Edmondson, D. E., Flavodoxin-cytochrome c interactions: circular dichroism and nuclear magnetic resonance studies, Biochemistry, 26, 5042, 1987. 227. Zanetti, G. and Curti, B., Properties of a cross-linked complex between ferredoxin-NADP+ reductase and ferredoxin, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G,, Eds., Walter de Gruyter, Berlin, 1984, 179. 228. Weber, C., Michel, B., and Bosshard, H. R., Spectroscopic analysis of the cytochrome c oxidasecytochrome c complex: circular dichroism and magnetic circular dichroism measurements reveal change of cytochrome c heme geometry imposed by complex formation, Proc. Natl. Acad. Sci. U.S.A., 84, 6687, 1987. 229. Ahmad, L, Cusanovich, M. A., and Tollin, G., Laser flash photolysis studies of electron transfer between semiquinone and fully-reduced free flavins and the cytochrome ocytochrome oxidase complex, Biochemistry, 21, 3122, 1982. 230. Bickar, D., Turrens, J. F., and Lehninger, A. C., Possible direct electron transport from cytochrome c to cytochrome a3, Fed. Proc. Fed. Am. Soc. Exp. Biol., 44, 678, 1985. 231. Hazzard, J. T., Poulos, T. L., and Tollin, G., Kinetics of reduction by free flavin semiquinones of the components of the cytochrome c-cytochrome c peroxidase complex and intracomplex electron transfer, Biochemistry, 26, 2836, 1987. 232. Hazzard, J. T., Cusanovich, M. A., Tainer, J. A., Getzoff, E. D., and Tollin, G., Kinetic studies of reduction of a 1:1 cytochrome oflavodoxin complex by free flavin semiquinones and rubredoxin, Biochemistry, 25, 3318, 1986. 233. Edmondson, D. E., personal communication.

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Volume III Chapter 15

STRUCTURE AND REDOX PROPERTIES OF CLOSTRIDIAL FLAVODOXIN Martha L. Ludwig and Catherine L. Luschinsky

TABLE OF CONTENTS I.

Introduction

428

II.

Structure-Function Studies of Flavodoxins: An Overview

430

III.

The X-Ray Structures of Flavodoxin from C. beijerinckii A. Structure Determination B. The Polypeptide Fold C. The FMN Binding Site 1. Description of the FMN-Protein Interactions in Reduced Clostridial Flavodoxin 2. The Conformation of Residues 56 to 60 in Oxidized C. beijerinckii Flavodoxin 3. Comparisons of Oxidized C. beijerinckii Flavodoxin with the Reduced and Semiquinone Structures 4. Comparisons between Reduced and Semiquinone Forms of C. beijerinckii Flavodoxin 5. NMR Measurements of Protein-FMN Interactions 6. The Conformation of the Isoalloxazine Ring D. Comparison with the Structure of M, elsdenii Flavodoxin

433 433 435 437

IV.

The Oxidized-Semiquinone Equilibrium A. A Summary of the Energetics B. The Turn Conformations C. The Kinetics of the Conformation Change

448 448 449 450

V.

The Semiquinone-Reduced Equilibrium A. Reduced Flavodoxins are Anionic Species B. Estimates of the Magnitude of Electrostatic Interactions C. A Review of the Energetics of Ring Bending in the Reduced Flavins

452 452 453 456

Some Comparisons of C. beijerinckii with A. nidulans and Other Flavodoxins A. The Phosphate Binding Loop B. Isoalloxazine Binding Site: the Bend Near Residue 60 C. Isoalloxazine Binding Site: the Structure Near Residue 90

457 457 458 459

Perspectives

460

VI.

VII.

437 440 442 444 446 446 447

Acknowledgments

461

References

462

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Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION Flavodoxins are the smallest members of the flavoprotein family, with molecular weights ranging from 14 to 23 kDa. Their size, stability, and ease of isolation have made these electron transfer proteins ideal subjects for a variety of structural and physical-chemical studies. Flavodoxins from at least 13 species of microorganisms (including two eukaryotes, Chondrus crispus and Chlorella fusca,) have been isolated and characterized. Flavodoxins are able to substitute for ferredoxins in a variety of electron transfer reactions at low potential,' including the phosphoroclastic splitting of pyruvate2 and the transfer of reducing equivalent to NADP in photosynthesis.3 They act as donors to both hydrogenase and nitrogenase: analysis of mutations in the M/gene cluster of Klebsiella pneumoniae has demonstrated the requirement for flavodoxin (ni/F) as electron donor to nitrogenase.4'5 Despite the broad distribution of flavodoxins, other unique functions have been difficult to ascribe to these FMN-containing electron carriers. However, activation of methionine synthase6-7 and pyruvate formate lyase8-9 may prove to be specific functions for flavodoxin in E. coli. In some species (clostridia, cyanobacteria), the expression of flavodoxins is tightly controlled by the levels of iron in the medium, but in other species such as C. crispus, expression appears constitutive.' ° In Chapter 14 (this volume), May hew and Tollin treat more fully the occurrence and physiological roles of flavodoxins.11 In this chapter we describe the three-dimensional structures of flavodoxins and discuss their relationship to the oxidation-reduction potentials, taking clostridial flavodoxin as our primary example. Crystal structures for oxidized flavodoxins from Clostridium beijerinckii MP12 (formerly Clostridium MP), Desulfovibrio vulgaris,13'14 and Anacystis nidulans15 were reported between 1972 and 1983. More recently the X-ray structure of flavodoxin from Chondrus crispus, a close structural relative of A. nidulans, has been determined,16 and structures of the reduced states of D. vulgaris flavodoxin have been reported.17 The semiquinone18'19 and reduced forms20'21 of C. beijerinckii flavodoxin have also been described, the latter only in brief summaries. With the advent of multidimensional NMR techniques, several groups have undertaken the determination of the three-dimensional structures of flavodoxins in solution. The methods have been applied most completely to the proteins from M. eisdenii22'24 and Anabaena 7120;25-26 partial structural information is also available for flavodoxins from D. vulgaris21 and A. nidulans;28 In vivo, flavodoxins act as one-electron carriers. To perform this function, the proteins perturb the two one-electron potentials of FMN by as much as 225 mV, if one takes the values of Draper and Ingraham29 as valid for free FMN (see Table 1.) The effects of the protein moiety on the ox/sq (oxidized semiquinone) potentials vary somewhat with species, but the potential for addition of the second electron, Esq/red, is typically -400 mV or lower at pH 7.0. Thus the outstanding property of these proteins, which display the lowest potentials known for protein-bound FMN or FAD, is their ability to destabilize the fully reduced form of the flavin cofactor. As shown in Scheme 1, the effects of the protein on redox potentials are linked to changes in association constants and correspond to free energies as large as 5 Kcal/mol. In the thermodynamic cycles of Scheme 1, the perturbation of the potentials by the protein and the change in affinity of the protein for FMN with redox state are equivalent expressions of the same phenomenon. Thus the control of potentials is conveniently discussed in the framework of relative free energies of association. The three-dimensional structures provide some indication of the chemical basis for such free energy differences. Comparison of the structures of oxidized and semiquinone forms of C, beijerinckii flavodoxin demonstrated that a structural change accompanies introduction of the first electron:19 a peptide unit in a reverse turn adjoining the flavin ring changes orientation to form a hydrogen bond with the N(5)H of the neutral FMN semiquinone (Sections III.C.3 and IV). This new interaction between the protein and the prosthetic group should increase the

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SCHEME 1. Thermodynamic cycles relating potentials and association constants for M. elsdenii flavodoxin at pH 7.0 and 20°C. Ka for oxidized FMN was determined from the fluorescence of mixtures of FMN and apoprotein;30 Ka values for FMN semiquinone and reduced FMN were calculated from Ka for oxidized FMN and the differences in potential between free and bound FMN. The potentials for flavodoxin are from Mayhew31 and the potentials for free FMN are from Draper and Ingraham.29 The one-electron FMN potentials determined by Anderson32 lead to much larger values of Ka for binding of the semiquinone. Ka for fully reduced FMN is smaller by a factor of 20 than Ka for binding of oxidized FMN; the ratio Kred/Kox depends on the value of the two electron midpoint potential for FMN, which is not in dispute.

protein-FMN association constant and is considered to be an important factor determining the ox/sq redox potential. A similar structural rearrangement occurs in D. vulgaris flavodoxin.14'17 In contrast, structures of the semiquinone and reduced forms of flavodoxin from C. beijerinckii are very similar; the hydrogen bond at N(5)H remains intact and no further rearrangements occur on addition of the second electron (Sections III. C.4 and V). Thus, changes in conformation are not the primary determinant of the sq/red potential. For a time it was presumed that the protein forced the reduced isoalloxazine ring into an unfavorable planar conformation, and that this distortion accounted for the low redox potential.20 However, NMR measurements33 subsequently demonstrated that unsubstituted free reduced flavins are essentially planar (Section V.C). Thus the "ring-strain" rationale for the low sq/red potential has been abandoned. Instead, speculation has focused on electrostatic effects: the reduced flavin is an anionic species, and the distributed negative charge34-35 is surrounded by a partly hydrophobic environment lacking compensating countercharges. This chapter begins by summarizing some of what is known about the structures and redox potentials of the whole family of flavodoxins. After this brief overview, we describe the structures of flavodoxin from C. beijerinckii obtained from recent refinement of the Xray data, concentrating on the structure of the fully reduced form, which has not been reported in detail elsewhere. We compare the structure of fully reduced C. beijerinckii flavodoxin with the solution structure of reduced Megasphaera elsdenii flavodoxin, determined by NMR methods, and with the semiquinone and oxidized forms of C. beijerinckii flavodoxin, and discuss the protein-FMN interactions and how they affect the observed oxidation-reduction potentials. A concluding section examines some structural variations among flavodoxins and their effects on oxidation-reduction potentials.

430

Chemistry and Biochemistry of Flavoenzymes TABLE 1 Redox Potential Values of Flavodoxins at pH 7.0a in mV Source Anabaena 7120 Anacystis nidulans Azotobacter chroococcum Azotobacter vinelandiP Chondrus crispus Clostridium beijehnckii Clostridium pasteurianum Desulfovibrio vulgaris Escherichia coll Klebsiella pneumoniae Megasphaera elsdenii Nostoc strain MAC Synechococcus lividus FMN a

b

E2 (ox/sq)

E, (sq/red)

Ref.

-196 -221 -215 -103 -229 -165 -222

-425 -447 -414 -522 -464 -458 -370 -399 -419 -438 -455 -422 -372 -414 -450 -172 -124

48 49 50 51 52 53 50 56 56 57 58 51 31 50 59 29 32

-92 -132 -103 -244 -170 -115 -210

-50 -238 -314

Some potentials were not measured at pH 7.0. For these, a —59 mV change per pH unit increase was used to estimate E2 at pH 7.0. It was assumed that El values did not change at pH values above 7.0. Additional values have been reported for the potential of A. vinelandii flavodoxin by Barman and Tollin,54 and Klugkist et al.55 Several flavodoxins with differing potentials have been isolated from strains ATCC 478 and OP.11 Values from Reference 53 are for the recombinant, dephospho enzyme.

II. STRUCTURE-FUNCTION STUDIES OF FLAVODOXINS: AN OVERVIEW From the X-ray and NMR structure analyses, it is evident that flavodoxins share a common polypeptide fold and are classic examples of proteins in which a central parallel sheet is surrounded on both sides by helices. In the flavodoxins for which tertiary structures have been determined,12'13'15'16-24 the central sheet consists of five strands whose order along the amino acid sequence may be denoted 2-1-3-4-5 (Figure 1). In the flavodoxins from A. nidulans15 and C. crispus16 a substantial insert divides the fifth strand into two parts. The stereo drawings of Figure 1 compare the folds of the flavodoxins from C. beijerinckii, a short chain of 138 residues, and from A. nidulans, a representative of longer chains, with 169 residues. From these two examples the similarity of the core of the folds is obvious, but differences in the vicinity of FMN are also apparent. Three-dimensional superpositions of the structures of C. beijerinckii, A. nidulans, and D. vulgaris flavodoxins, together with descriptions of the flavodoxin from C. crispus,16 define several loci at which insertions or deletions have been accepted in the course of evolution. Figure 2 displays the sequence alignments derived from matching of the threedimensional structures. A major insertion of 20 amino acids, beginning at residue 120 of A. nidulans flavodoxin, clearly distinguishes A. nidulans from the other flavodoxins in Figure 2. Alignments based on amino acid sequences40 indicate that a similar insertion has occurred in the flavodoxins from A. vinelandii and K. pneumoniae, and suggest that flavodoxins with chains longer than A. nidulans also incorporate as many as ten extra residues

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FIGURE 1. Stereodrawings of the polypeptide folds of (A) C. beijerinckii and (B) A. nidulans flavodoxins. The structures are oriented to give identical views of the sheet and the first helix. The major insert in A. nidulans flavodoxin is at the lower right of panel (B), starting at residue 120; the shorter insert near residue 90 is just below the flavin ring. This view displays the different orientation of the flavin ring in the two structures. In each drawing, residues that are charged at neutral pH are indicated with + or - signs at the Cot positions. The charge on Asp-90 of A. nidulans flavodoxin (panel B) is partly obscured.

somewhere between positions 65 and 80 of A. nidulans flavodoxin.41 These major differences in molecular size result from insertions distant from the FMN. While they do not directly affect protein-flavin contacts, these variations may influence formation of electron transfer complexes. The postulated roles of specific residues in facilitating electron transfer are not considered in this chapter, but are discussed by Mayhew and Tollin11 in Chapter 14 of this volume. Two shorter gaps or inserts are observed near residue 60 and near residue 90, in portions of the polypeptide which are close to the FMN prosthetic group (Figure 2); insertions in these regions produce conformational variations which affect FMN-protein interactions. From differences in the circular dichroism and optical spectra of flavodoxins, Edmondson and Tollin42 and D'Anna and Tollin43 proposed that the flavodoxin family be subdivided into two major groups, one providing a more apolar environment for the FMN ring and able to bind riboflavin, the other more polar with little affinity for riboflavin. This division seems

432

Chemistry and Biochemistry of Flavoenzymes

FIGURE 2. Alignments of the residues of A. nidulans™ C. beijerinckii^1 and D. vulgaris3*'39 flavodoxins, based on the correspondence of the Ca coordinates. The structures were superimposed by fitting the sheet strands and the first helix, and structural equivalences were determined from Ca separations and by viewing on a graphics display. The numbers overhead refer to the C. beijerinckii sequence; numbers at the ends of the individual lines refer to the individual sequences. Vertical lines denote structural equivalences, where Cot atoms are almost superimposed. The structures around positions 40, 70, and 130 (C. beijerinckii numbering) are considered to be similar, i.e., the chain folds are congruent in these regions although the chains are displaced from one another in the different structures. These similarities are denoted by connecting dots. At insertions or locations where the chains follow different paths, no symbols connect the sequences. The flavin binding sequences at 56 to 59 and 89 to 91 are not highly conserved; inserts and substitutions occur in these regions.

to correlate well with the conformation of the sequences between residues 90 and 100. In C. beijerinckii and M. elsdenii flavodoxins, representatives of the more polar class, the residues beyond position 91 do not wrap around the flavin as they do in A. nidulans, C. crispus, or Anabaena 7120 flavodoxins. The variations of potentials and other properties that may result from substitutions near 60 and near 90 are discussed in Section VI (cf. Figures 16 to 18). The affinity of apoflavodoxins for FMN can be measured by monitoring spectral or fluorescence changes as apoprotein is added to FMN.1*30 The association constants at neutral pH are large; representative values are: 2 x 109 M" 1 for M. elsdenii™ 5 X 109 M"1 for D. vulgaris,44 and 6 x 108 M" 1 for A. nidulans.45 A. nidulans46 and D. vw/garw1-43 flavodoxins also bind riboflavin with an association constants of the order of 106. The kinetics of the association and dissociation reactions have been discussed elsewhere.' It has sometimes been assumed that significant changes in protein conformation accompany the binding of FMN. Studies of the structures of the apoproteins are now feasible, using NMR methods. The initial results47 suggest that the apoprotein retains its characteristic fold in the absence of the cofactor, but provide evidence for changes in the conformations of residues constituting the FMN binding site. Oxidation-reduction potentials have been measured for flavodoxins from a number of species (Table 1). Significant species-dependent variations are observed, especially for the oxidized/semiquinone (ox/sq) potential. Despite these individual variations (from -50 to -240 mV for Eox/sq), the pattern of shifts in graphs of potential vs. pH, relative for free

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FIGURE 3. Graphs of potential vs. pH for FMN and for M. elsdenn flavodoxin. The one-electron potentials for flavodoxin (•) were taken from Mayhew31 and from Stankovich;62 the one-electron potentials for free FMN (—) are from Draper and Ingraham,29 simplified as in their paper. Linear pH-dependent regions of the FMN potentials have been extended to intersect with horizontal segments at the experimentally determined pK values of 6.5 for reduced FMN and 8.6 for FMN semiquinone. Because of the increased pK of flavodoxin semiquinone, the flavodoxin graphs do not change slope at pH 8.6.

FMN, is similar for all members of the flavodoxin family.60 The graph of the ox/sq potential vs. pH has a slope of - 60 mV throughout the accessible range of pH, and in most species the value at pH 7.0 is higher than that for free FMN (Figure 3). The dependence of Eox/sq on pH beyond pH 8.6 reflects an increase in pK of the neutral flavin semiquinone that results from interaction with the protein (see Section IV of this chapter). The potential for the sq/ red equilibrium is decreased dramatically by association of FMN with apoflavodoxins; it is this shift which is primarily responsible for the ability of flavodoxins to function as oneelectron reagents. In free FMN the graph of sq/red potential vs. pH shows inflections near pH 6.5, the pK for formation of the reduced N(l) anion, and near pH 8.6, the pK of N(5) in the semiquinone. In flavodoxins, the sq/red potential is constant with pH from approximately pH 7.0 upward. NMR measurements25-27'63 establish that the reduced flavin in all flavodoxins examined is anionic at pH 7.0, consistent with the pH-independent behavior of the potential above pH 7.0. An inflection point below pH 7.0 reflecting a redox-linked pK of 5.8 has been observed in M. elsdenn flavodoxin;31 the corresponding pK in C. beijerinckii flavodoxin is 6.7.1'56 Recent studies of M. elsdenn flavodoxin suggest that the pK observed in measurements of the sq/red potential corresponds to ionization of a group on the protein rather than the titration of bound reduced FMN61 (Section V).

III. THE X-RAY STRUCTURES OF FLAVODOXIN FROM C. beijerinckii A. STRUCTURE DETERMINATION Table 2 summarizes the crystallographic data and the results of refinement for each oxidation state. The cell parameters for crystals of C. beijerinckii flavodoxin vary with oxidation state as shown. Differences between the measured X-ray intensities for the oxidized

434

Chemistry and Biochemistry of Flavoenzymes TABLE 2 Structure Analysis of Flavodoxin from C. beijerinckii A. X-Ray Data (Space Group P3l2l)

Cell a, b Cellc Resolution Unique reflections measured Fraction of unique reflections

Oxidized

Semiquinone

Reduced

61.56 70.36 10.0-1.9 A 12435 98%

61.36 70.98 10.0-1.8 A 14833 98%

61.68 71.05 10.0-1.8 A 12916 86%a

B. Crystallographic Refinement

R-factor for starting model Cycles Solvents R-factor for current model Overall B rms Shift (main-chain atoms) a

Oxidized

Semiquinone

Reduced

0.265 36 98

0.274 10 96 0.204

0.296 40 97

0.186 21.0 A2 0.38 A

17.3 A2 0.32 A

0.190 17.3 A2 0.36 A

The flow cell limited the accessible range of diffraction space.

and semiquinone forms were large enough to suggest nonisomorphism, and hence the structures were determined independently by multiple isomorphous replacement.12-18 In retrospect, the intensity differences have been shown to result from small rotations of the molecules involving changes in intermolecular contacts.19 The interactions between Asn-137 of one molecule and residues 57 to 59, near the FMN in a neighboring molecule, alter when the flavin is reduced to the semiquinone form (see Section III.C, Figures 7 and 10). Crystals of the semiquinone and fully reduced species proved sufficiently isomorphous to use the semiquinone structure as the starting point for analysis of fully reduced flavodoxin. Crystals of the semiquinone form were grown from ammonium sulfate in a nitrogen atmosphere at 4°C; for analysis of oxidized flavodoxin, they were allowed to oxidize in airsaturated buffers. In initial structural studies of reduced flavodoxins,20 crystals reduced with excess dithionite were sealed in X-ray capillaries in the usual fashion. Reoxidation was monitored visually; when the crystals became purplish, data collection was stopped and the fraction of semiquinone estimated from EPR measurements.64 In contrast, the data used for refinement of the reduced structure were obtained from a crystal mounted in a flow cell, using a diffractometer system at Wayne State University.21 The crystal was bathed with a degassed solution of 2.8 M ammonium sulfate, buffered with Tris at pH 8.0, and made 0.2 M in dithionite.64 Refinements leading to the structural models presented here were carried out with the Hendrickson-Konnert programs,65 starting from coordinates for the oxidized and semiquinone forms, deposited in the Protein Data Bank (3FXN, 4FXN) in 1978. The 1978 models had been obtained by a combination of difference Fourier, real space, and group least squares refinements, and had discrepancy (R) factors of 0.211 and 0.200 for the oxidized and semiquinone states, respectively, for data beyond 5 A with I > 2 cr (Table 2). The models, which had been derived before the era of computer graphics, were displayed for comparison with Fourier or difference Fourier maps in order to inspect the conformations that had been assigned. For each oxidation state the positions of bound solvents were reviewed before beginning refinement. Series of refinements were alternated with interactive display and model-building. Solvents were retained in the models only if their scattering contributions

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at high resolution (a combination of occupancies and thermal factors) wre greater than 30% of an oxygen with full occupancy and a B of 18.0 A2. This criterion corresponds roughly to peak heights that are greater than 3 cr in difference maps. The quality of the structures has been improved significantly by the recent refinements, as measured by the decrease in R for all data (Table 2) and adherence to standard geometry. The overall rms change in main chain coordinates, including the results of manual rebuilding, is 0.32 to 0.38 A, and we estimate the average error in atomic position to be approximately 0.20 A for each oxidation state. Overall B values, given in Table 2, indicate that extension to still higher resolution is feasible; these experiments will be facilitated by the availability of recombinant protein.66 Additional solvent sites should be incorporated in higher resolution models. There are two regions, 39 to 47 and 120 to 125, which have significantly higher thermal factors than the rest of the model. After rebuilding, the 39 to 47 backbone is a good fit to the density; however the side chains in this region remain partly disordered. The only area of the map in which the electron density for backbone atoms is difficult to interpret occurs at peptides 122 to 123 and 123 to 124. As expected, the electron densities for some surface side chain atoms are not well defined. In certain cases, there is little or no density for side chain atoms at the level of 1 a, and in others, there is density which has more than one interpretation. The surface residues in question are exclusively lysines, glutamates, aspartates, and asparagines. Five of 10 lysine residues are poorly defined in one or more of the different oxidation states. Ten of the eighteen glutamate side chains have ill-defined density in all three oxidation states; two have no density at the expected side chain position. Four of the aspartates and three asparagines are poorly defined in all three oxidation states. All of the above side chains are outside the flavin binding site. Only one residue near the flavin is disordered; the side chain of Asp58 has more than one conformation in the oxidized structure (see below). B. THE POLYPEPTIDE FOLD This section briefly reviews the features of the protein, comments on clarifications that have resulted from the refinement, and discusses some of the protein-sol vent interactions. The descriptions and drawings are taken from the structure of fully reduced C. beijerinckii flavodoxin, since major themes of this chapter are the sq/red equilibrium and comparison with the solution structure of reduced M. elsdenii flavodoxin. Revisions of the structural model apply to all three oxidation states unless otherwise noted. The secondary structures in reduced C. beijerinckii flavodoxin, assigned by the algorithm of Kabsch and Sander67 (Figure 4), show the alternating p-sheet and a-helical regions that are typical for an a/p fold. Turns or bends following the C-termini of the first, third, and fourth strands of sheet constitute the binding site for FMN (Figure 1). Each of these chain reversals includes unusual conformations that will be described in more detail below (Section C). The parallel sheet comprising the core of the molecule is connected with the topology - lx,2x,lx,lx, and is the analogue of sheets found in the nucleotide-binding folds of dehydrogenases, except for deletion of the third strand found in dehydrogenases.70 A drawing of the hydrogen bonding within the sheets is shown in Figure 5, where one can see irregularities in the final strand. Residue Val-111 adopts the a-conformation typical for bulges, which are relatively rare features in parallel sheets.71 However, regular hydrogen bonding is resumed at residue 115 rather than at 114, allowing two residues to be inserted in strand 5 to form a small loop that is stabilized by water molecule trapped between N-84 and 0-112. The long insert found in A. nidulans flavodoxin starts after the residue corresponding to Thr-113 of C. beijerinckii flavodoxin (Figure 2). Flavodoxin from C. beijerinckii has been described as including four a-helices;12'19

436

Chemistry and Biochemistry of Flavoenzymes

FIGURE 4. Assignments of secondary structures in reduced C. beijerinckii flavodoxin37 (line 1) and reduced M. elsdenii flavodoxin68'69 (line 4), determined from X-ray and NMR coordinates by the algorithm of Kabsch and Sander.67 For the M. elsdenii structure, the assignments are taken from Table 1 of Reference 24. The sequence alignments, in boldface, were derived from Reference 23, Reference 24, and Figure 13. The phosphate binding region (7 to 12) and the isoalloxazine binding regions (55 to 59 and 89 to 91) are highly conserved; the three residue hairpin turn73 at 56 to 60 appears in both structures. The major secondary structural features are also conserved. Differences in the conformations arise from two inserts in C. beijerinckii flavodoxin and from relatively small changes in coordinates which affect the choice of turn vs. 3helix or bend, van Mierlo et al.24 note the deletion of C. beijerinckii residues Leu-115 and Asp126 in building their model, but inspection of the structures suggests that the residue inserted in sheet strand 5 of the C. beijerinckii chain is actually Pro-114.

however, after refinement and reorientation of the peptide bond between 42 and 43, residues 40 to 43 can also be characterized as a turn of a-helix. Oxygens 39 and 40 accept a-helical hydrogen bonds, and the cj>,v[/, angles of residues 40 to 43 lie in the a-region of conformational space. A corresponding short a-helix is found in flavodoxin from C. crispus,16 and in A. nidulans flavodoxin this region is characterized as a 310 helix.15 The helices in C. beijerinckii flavodoxin have therefore been renumbered. What we now call a-helix 3 begins at Glu-67 and terminates with bifurcated hydrogen bonding from O-70 and 0-71 to N-74. The peptide at 75 to 76 was rebuilt and smaller modifications and refinement have resulted in more regular helical conformations for the sequence 67 to 73. Residues 62 to 66 follow a righthanded helical path and would be considered part of the helix, except for an insertion at Ser-64 and rotations of the peptide planes at 63 to 64 and 64 to 65, all of which accommodate a proline residue at position 66. Helix 4, starting at residue 93, ends with 1 -> 5 and 1 - > 3 hydrogen bonds connecting O-103 and O-104 with N-108 and N-107, respectively. A turn of 310 helix at residues 121 to 124 precedes the final a-helix. Solvents have an important role in determining the stability and conformation of protein structures, and one aim of the refinements has been the examination of solvent-protein

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437

FIGURE 5. A stereoview of the hydrogen bonding in the parallel p-sheet of C. beijerinckii flavodoxin. Distances separating N and O atoms adjoin the dashed lines designating the hydrogen bonds. The trapped solvent which bridges N-84 and O-112 and the two residue loop formed by Glu-112—Thr-113 can be seen at the right side. Atoms beyond Cp have been omitted to simplify the drawing. Three additional hydrogen bonds that are almost eclipsed in this view are: N-31-O-1, 2.86 A; N-55-O-87, 2.94 A; and N-119-O-88, 2.86 A.

interactions. In the present list of well-defined solvent sites, we distinguish several categories. The first includes five solvents that are embedded in the protein. Although they may contact other bound waters, these solvents are regarded as part of the protein structure. Three are near the FMN (Table 3): one adjoins a phosphate oxygen, and the other two are in the vicinity of the isoalloxazine O(2) and Glu-59 (see Section C.I). The fourth structural solvent is trapped between two strands of (3-sheet (Figure 5), and the fifth is the interior end of a cluster of solvents which fills an invagination in the structure lying between Val-60-Leu-61 and Asp-98. Bound solvents situated in the first layer around the protein surface, contacting carbonyl O, peptide NH groups, of polar side chains, constitute a second category. Thirtyfour of the waters identified in the current model for reduced C. beijerinckii flavodoxin are associated with carbonyl oxygens and sixteen with peptide NH groups. Finally the list includes some solvents that interact only with other bound waters. These are located primarily in clusters which form intermolecular bridges. One solvent site (at FMN O(4)) is found only in the semiquinone and reduced structures; other variations of solvent sites with oxidation state have not been fully explored. C. THE FMN BINDING SITE 1. Description of the FMN-Protein Interactions in Reduced Clostridial Flavodoxin FMN is bound noncovalently to clostridial apoflavodoxin by a series of hydrogen bonds and other nonbonded contacts. Three sections of the polypeptide chain contribute to the binding site, forming subsites which interact with different fragments of the prosthetic group. Residues 7 to 12 and 54 bind the FMN phosphate; residues 12, 55, and 87 interact with the ribityl side chain; and residues 55 to 59 and 89 to 92 contact the isoalloxazine ring. The pocket into which the prosthetic group fits is outlined with van der Waals radii in the Plate

438

Chemistry and Biochemistry of Flavoenzymes

FIGURE 6. Schematic drawing of the FMN binding site in reduced C. beijerinckii flavodoxin, showing the polar interactions that are listed in Table 3A. Hydrogen bonds between FMN and protein atoms are dashed; hydrogen bonds between FMN and solvents are drawn as dotted lines.

1*; altogether 28 atoms of the protein are within 3.5 A of some FMN atom. Binding to the protein decreases the accessible surface area72 of FMN by 525 A2, Solvents are found near the remaining exposed surface of the FMN (see Figure 8). Figure 6 is a diagram showing the hydrogen bond contacts between the flavin and protein atoms in reduced clostridial flavodoxin, and Table 3 lists the contacts between the protein or solvent and the FMN for all three oxidation states. The phosphate binding subsite is depicted in Figures 6 and 16. Following the bend at residues 8 and 9, Gly-10, with , i|* angles of 108°, - 10°, redirects the chain into the start of the first helix. The backbone orientations allow five successive NH dipoles, at residues 8, 9, 10, 11, and 12, to point toward the phosphate oxygens. Four side chain hydroxyls at Ser-7, Thr-9, Thr-12 and Ser-54 also interact with phosphate oxygens (Table 3; Figures 6, 16). Analysis of the NH proton chemical shifts in the phosphate binding sequence of M. elsdenii flavodoxin indicates strong hydrogen bonding at the peptides corresponding to residues 8,9, 11, and 12 of C. beijerinckii flavodoxin,23'34 but not at the NH equivalent to Gly-10. In the crystal structure the geometry for this hydrogen bond is poorer than for the other NH-O interactions. In C. beijerinckii flavodoxin, a bound solvent is hydrogen bonded to OP(I) of the dianionic phosphate, and forms a bridge to other solvent sites. Although solvent contacts the phosphate, the 31P resonance of FMN bound to C. beijerinckii flavodoxin is not broadened in the presence of the paramagnetic probe,44 Mn 2 + , an indication that the probe is excluded from contact with the phosphate. The isoalloxazine binding pocket in reduced C. beijerinckii flavodoxin is displayed in Plate 1, and in Figures 8 and 15. The sequence 57 to 59 forms a three residue turn connecting Met-56 and Val~60, which are hydrogen bonded in antiparallel fashion73 (Figure 4). Residues 56 to 59 contact the re face and N(5) edge of the flavin ring. There are several short hydrogen bonds between these residues and FMN. The carbonyl oxygen of Ala-55 is within 2.7 A of 02' of the ribityl side chain, the carboxylate oxygen of Glu-59 is associated with N(3)H, the peptide NH of Glu-59 points toward O(4), and the carbonyl oxygen of 57 is close to N(5)H. Close nonhydrogen bonding contacts are made by the S-CH3 of Met-56, by O 55, *

Plate 1 follows page 454.

Volume III TABLE 3 Interactions Between FMN and Protein Atoms in Clostridium beijerinckii Flavodoxin A. Hydrogen Bonding Contacts Distance in A FMN atom

Contact atom

N(l) 0(2)

Gly-89 N Gly-89 N Trp-90 Nb Gly-91 N Wat-246 O Glu-59 Otl Glu-59 N Wat-278 O Gly-57 O Ala-55 O Wat-248 0 Wat-253 O Asn-11 OS Ser-87 Oy Wat-253 O Gly-8 N Ser-54 Oy Wat-245 O Ser-7 O-y Thr-12 N Thr-12 O-y Thr-9N Thr-9 O-y Gly-10 Nb Asn-11 N

N(3) 0(4) N(5) 0(2') 0(3') 0(4') OP(I) OP(II) OP(III)

Oxidized

Semiquinone

Reduced

3.08 2.97 3.30 2.89 3.28 2.84 2.78

3.09 2.98 3.18 2.84 3.29 2.84 2.87 2.81 2.90 2.69 2.80 2.80 2.74 2.76 3.24 2.80 2.82 2.82 2.80 2.72 2.83 2.70 2.57 3.02 3.05

3.02 2.98 3.11 2.78 3.26 3.01 2.80 2.57 3.03 2.62 2.72 2.74 2.77 2.74 3.30 2.80 2.88 2.89 2.76 2.82 2.92 2.76 2.63 2.94 3.08

— —

2.69 2.73 2.75 2.89 2.65 3.28 2.62 2.80 2.87 2.82 2.80 2,79 2,75 2,61 3.03 3.02

B. Nonhydrogen Bonding Contacts Distance in A FMN atom

Contact atom

C(2) C(4) 0(4)

Gly-89 N Wat-278 0 Glu-59 Ca Gly-57 O Asp-58 O51 Trp-90 C£2 Ala-55 O Trp-90 C£2 Gly-57 N Gly-57 O Met-56 S Wat-299 O Met-56 S Trp-90 On2 Ala-55 O Trp-90 Cn2 Ala-55 O Ala-55 O Ala-55 O

C(4A)

N(5) C(6) C(7) C(7)M C(8) C(9a) N13, an increase of at least 4.5 units from the pK for free FMN semiquinone. This shift, corresponding to a free energy of about 6 kcal/ mol, cannot be attributed solely to the hydrogen bond with the carbonyl group of Gly-57, since electrostatic interactions with Glu-59, whose carboxylate group is 6.3 A from N(5), and the polarity of the environment, also influence the ionization. Instead, it sets an upper limit for the energy associated with the hydrogen bond. As a result of the conformation change, the carbonyl oxygen of Gly-57 is about 3.4 A from O(4) of the FMN ring in the semiquinone and fully reduced structures; these atoms are separated by approximately 6.0 A in the oxidized structure. Charge repulsion between the oxygens will therefore be much greater in the reduced states: partial charges of -.38 e are assigned to peptide oxygens for computation of empirical potential functions,85 and Hall34 calculates a charge of -.545 e for O(4) in oxidized lumiflavin and -.657 e in the anionic form of lumiflavin hydroquinone. Without a hydrogen bond at N(5), the electrostatic repulsion between these atoms is probably sufficient to force the structure into the alternative conformation found in oxidized flavodoxin. The equilibrium between the conformers represents an energetic balance that can be modulated by the effects of the amino acid sequence on conformational energies. B. THE TURN CONFORMATIONS The structural change which is observed on reduction, while localized, is rather intriguing. Rearrangement at this locus appears to be a key feature in the behavior of flavodoxins. However, the relative energies of the different rotamers of the 57 to 58 peptide unit in clostridial flavodoxin have not been carefully studied by computational techniques. Analysis of the transition in C. beijerinckii is currently complicated by uncertainty about the conformation of the turn of oxidized C. beijerinckii flavodoxin in solution. Assuming that the Gly-Asp peptide is the trans conformer, a crude estimate of the relative energies of these residues in the oxidized and semiquinone structures may be obtained from Ramachandran plots.85 87 A simplistic analysis of this sort suggests that the semiquinone conformation may be lower in energy than the oxidized as long as residue 57 is glycine (Table 5): residue 57 in the semiquinone structure adopts a conformation that favors glycine (see below). The problem needs to be addressed by more sophisticated computations. Attractive approaches include molecular dynamics simulations with stochastic boundary conditions88 or energy minimization89 that uses constraints to maintain the ends of the bend. Conformational sampling and minimization has been applied to another segment of flavodoxin,90 and strategies have been proposed for the prediction and analysis of loop conformations.89*91 Experimental estimates of the impact of conformational energies on the redox potentials can be obtained by mutation of Gly-57. The conformation of this residue in the semiquinone structure is well defined, with torsion angles ,i|; = 48°, - 134° (Table 5). This conformation

450

Chemistry and Biochemistry of Flavoenzymes TABLE 5 Backbone Torsion Angles in C. beijerinckii Flavodoxin A. Conformation of the Phosphate-Binding and 90s Loops Residue Ser-7 Gly-8 Thr-9 Gly-10 Asn-11 Tyr-88 GIy-89 Trp-90 GIy-91 Asp-92 Gly-93

*

-141 -118 -105

127 -30 -3 -10 -58 169 -76 -6

-95

-156

-130

22 8

-141

-93 -121

108 -60

106

Area of Ramachandran diagram 3 Region Near position 2 of Type I turn Near position 3 of Type I turn Near Gly bulge region a Helix (3 Region Unusual Bend region Unusual, edge of p region Edge of (3 region Gly bulge region

B. Conformation of the Bend at Residues 56 to 60 Residue Oxidized Met-56 Gly-57 (trans) Asp-58 Glu-59 Val-60 Reduced Met-56 Gly-57 Asp-58 Glu-59 Val-60

*

Area of Ramachandran diagram

177 92 52 53 171

3 Region Near position 2 of Type II turn Near position 3 of Type II turn Near left-handed a helix p Region

-136

175

54

-133

(3 Region Near position 2 of Type II' turn (3 Region Near left-handed a helix p Region

-107

-43 88 59 -132

-115

57 -131

90 50 178

is typical for the second residue in Type II' p-turns, and is relatively high in energy for residues other than glycine. Mutagenesis experiments which swap Gly-57 for other residues, or introduce ^-branched side chains at this position, should yield proteins with decreased ox/sq potentials if the above analysis of the energetics is correct. Replacement of Gly-57 by Ala, Asp, or Asn is in fact observed to decrease the redox potential by 38 to 70 mV; replacement by Thr has a much larger effect: the measured ox/sq potential (pH 7.0) is - 270 mV.92 Initial X-ray studies of these mutants show that the side chains are accommodated without perturbing the remainder of the FMN binding site so that the effects may be attributed to local alterations in the bend sequences. These preliminary results are consistent with substantial contributions of conformational energies to the ox/sq equilibrium. C. THE KINETICS OF THE CONFORMATION CHANGE The rearrangement of the Gly-57-Asp-58 peptide must be associated with a significant activation energy, and may affect the overall rate of interconversion of oxidized and semiquinone forms of the protein. One-electron reduction of the oxidized protein presumably proceeds by electron transfer to form the anionic semiquinone, followed by protonation at N(5). While each of these steps is associated with well-documented changes in the visible spectrum,1 an accompanying (or subsequent) first-order conformation change may be spectrally silent, so that it is not simple to determine its rate.

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451

The kinetics of reduction have been examined using stopped-flow93 and pulse radiolysis methods.94-95 As expected, if the conformation change affects the energy of activation for the ox/sq conversion, the first one-electron step in reduction of M, elsdenii flavodoxin by SO2~ is slower than the second by a factor of 450: k, is 9 x 104 M™ 1 s~ ! and k 2 is 4 x 107 M~* s~ * at low ionic strength.93 To look for evidence of a conformation change occurring on the ms time scale, reduction of oxidized M. elsdenii flavodoxin by methyl viologen and oxidation of the semiquinone by ferricyanide were followed by stopped-flow spectrophotometry,96 monitoring changes in flavin absorbance. A rate-limiting relaxation would be expected to exhibit first order kinetics, but in both directions the reaction remained second order as reactant concentrations were increased to the accessible limit of kobs, about 1000 s~'. These results seem to imply that the structural rearrangement occurs with a rate constant exceeding 103 s ~ l . However, the interpretation is not unequivocal, since the two reactions are not the microscopic reverse of one another, and conformational relaxation might occur after the spectral change in both directions. Pulse radiolysis was used by van Leeuwen et al.94 to study reduction of flavodoxin on the time scale of 105 s" 1 . Reduced methyl viologen served as a reductant for M. elsdenii flavodoxin, and the kinetics were analyzed according to a model in which a postulated protein intermediate relaxed to product at a rate of 105 s"1. In further pulse radiolysis studies, Anderson et al.95 followed the reduction of the oxidized flavodoxin in the presence of formate. Within 2 JULS the spectrum of the anionic semiquinone species could be detected; this transient intermediate was converted to the neutral N(5) protonated species at a rate of about 105 s"1, which was almost independent of pH. No other relevant spectral changes were observed. Thus the relaxation observed by van Leeuwen et al.94 might be the protonation step. It is not certain that protonation must occur after the conformation change. The pK of flavodoxin semiquinone with O-57 pointing away from the flavin is probably high enough for protonation to occur without reorientation of the peptide unit. Thus, while the kinetic analyses seem to constrain the conformation change to a rate between 103 and 105 s"1, none of the results is unambiguous. Rates of electron exchange have been determined by analysis of NMR spectra of equilibrium mixtures containing small fractions of the paramagnetic semiquinone species.97 Exchange rates for M. elsdenii flavodoxin were deduced from the linewidths of several methyl resonances found in the high field region, and assigned to Ala-56 and Leu-62. In mixtures of oxidized and semiquinone species, these resonances show slow exchange behavior, and the exchange rate derived with this assumption is less than 5.7 X 103 M~l s"1. In contrast, mixtures of semiquinone and fully reduced molecules exchange electrons rapidly on the NMR time scale, with estimated rate constants that are dependent on ionic strength, and range from 105 to \(f M~l s ~ l . From these two rates the difference in activation energy is calculated to be approximately 3.4 kcal/mol,97 which might be attributed to the conformation change. The Ala-56 methyl reporter resonance certainly detects electron exchange since it contacts the flavin ring.22-24 However, from the equivalent coordinates for C. bei~ jerinckii flavodoxin, Ala-56C3 would be more than 5 A from any of the backbone atoms of Gly-58 and Ser-59; the closest atom which would move during the conformation change would be O-57 of methionine, 5.2 A away. It would be interesting to repeat these experiments with a 13C label in Gly-58 of M. elsdenii flavodoxin. If the conformation change occurs on the ms time scale, as the data imply, then some special features of the structure probably facilitate the process. As noted in Section III.C, a cis-trans interconversion at these rates would be exceptional. The activation energy of approximately 18 kcal/mol for cis-trans conversions74 corresponds to rates less than I s " 1 , and several measured X-Pro isomerizations in proteins proceed at rates in this range.98-99 Barriers for conversions among turn conformations are not well established. Saddle points on Ramachandran plots, on the order of 4 to 5 kcal/mol above minima for Type I or Type II' turn conformations, are calculated for a models like N-acetyl-W methylalaninamide,

452

Chemistry and Biochemistry of Flavoenzymes TABLE 6 Partial Charges on Atoms in Lumiflavin3 Atom

N(l) C(2) 0(2) N(3) C(4) 0(4) C(4a) C(10a) N(5) N(10) a

b

Oxidized

Reduced

Reduced anionb

-.413 + .734 -.548 -.328 + .641 -.545 -.106 + .292 -.013 -.045

-.220 + .689 -.561 -.296 + .655 -.557 -.240 + .258 -.069 -.109

-.470 -.661

-.657

From Hall, L. H., Orchard, B. J., and Tripathy, S. K., Int. J. Quant. Chem., XXXI, 217, 1987. With permission. Calculated with optimized geometry in MINDO/3. Only the major changes occurring on deprotonation are listed. In the reduced anion, about 0.25 of the charge resulting from deprotonation is distributed in the dimethylbenzene ring. In the anion, all atoms except C(2), C(5a), N(5), and C(10a) are more negatively charged than in the neutral reduced form.

whose ends are not constrained,100 and are not likely to be applicable to a dipeptide in situ in a protein. The flavodoxin system offers a good opportunity to study these phenomena.

V. THE SEMIQUINONE-REDUCED EQUILIBRIUM The equilibrium constant for reduction of the semiquinone is larger for free FMN than for M. elsdenii or C. beijerinckii flavodoxin by about 5 kcal/mol (cf. Scheme 1), employing the FMN potentials of Draper and Ingraham,29 or by about 7 kcal/mol using the potentials of Anderson.32 The calculated Ka for association of reduced FMN is thus smaller than for the semiquinone by a factor of at least 103. This considerable decrease in the apparent affinity for FMN occurs in the absence of significant structural changes in the holoprotein. The possibility that a high energy conformer of the reduced isoalloxazine ring is bound is discussed in Section C for historical perspective. However, current attempts to explain the effects of the protein environment on Ka for binding of reduced FMN and on the sq/red equilibrium have focused on electrostatic interactions between the reduced anionic flavin and its protein milieu. A. REDUCED FLAVODOXINS ARE ANIONIC SPECIES Evidence that bound reduced FMN is the anionic species in flavodoxins comes from measurements of the chemical shifts of 15N(1). In flavodoxins from M. elsdenii,63'™ C. beijerinckii A. vinelandii,63 D. vulgaris,21 and Anabaena 7120,25 the chemical shifts of 15 N(1) in the reduced state vary from 182 to 187 ppm at pH 8.0. Comparison with free reduced FMN, which displays characteristic 15N(1) chemical shifts783 of 128 ppm at pH 5.5 and 181.3 at pH 8.5, with a pK78 at 6.7, establishes that bound reduced FMN is anionic in all these flavodoxins. No N(l) pKs have been observed in flavodoxins by 15N NMR spectroscopy; even at pH 5.5 the chemical shift in reduced M. elsdenii flavodoxin is 182 ppm79 (see below). The most important consequence of the association of reduced FMN with the protein may be the immersion of the reduced flavin ring, with its distributed net negative charge (Table 6), in a medium whose average dielectric constant is less than that of water (Plate

Volume III

453

TABLE 7 Acidic Residues in C. beijerinckii and A. nidulans Flavodoxin C. beijerinckii* Residue Asp-58 Glu-59 Glu-62 Glu-63 Glu-67 Asp-92 Asp-98 3 b

Distance (A)b

5.0 3.6 9.3 9.3 13.7 8.9 10.7

A. nidulans FMN atom

0(4) N(3) C(7)M C(6) 0(4) 0(2) 0(4)

Residue Glu-61 Asp-90 Asp-96 Asp- 100 Asp- 144 Glu-145 Asp- 146

Distance (A)

FMN atom

8.0 5.2 9.3 6.0 7.3 12.5 7.1

0(4) N(l) 0(4) N(3) 0(2) N(l) N(l)

Coordinates from the structure of the reduced form. Measured from the C^ or C5 atoms of Asp or Glu.

1*, Figure 15). The accessible surface72 of the 18 atoms of the isoalloxazine ring decreases from 356 A2 in free FMN to 113 A2 in the holoprotein. The apolar and polar protein contacts with the negatively charged isoalloxazine heteroatoms, N(l), O(2), O(4), N(3), N(5), and N(10) can be seen in Figure 15 and Table 3. Small structural changes that occur at O-55, Glu-59, and water 278 when the semiquinone form is reduced are consistent with the postulated electrostatic effects (Section III. C.4). The protein offers no compensating positive charges in the immediate vicinity of the prosthetic group; instead the surroundings include seven aspartate or glutamate residues within a 15 A radius of the flavin ring. Acidic groups are highlighted in Plate 2* and distances to their isoalloxazine neighbors are given in Table 7. If charging of the flavin ring as a result of reduction of the semiquinone dominates the energetics of interactions with the protein, and results in unfavorable (repulsive) interactions, it is important to understand why reduced FMN is not bound as the neutral dihydroquinone species, protonated at N(l). Examination of the structure suggests that protonation is resisted by steric hindrance. In reduced C. beijerinckii flavodoxin, N(l) is 3.0 A from the peptide nitrogen of Gly-89, which is oriented to hydrogen bond to N(l); protonation of N(l) appears to be impossible unless the structure is perturbed. To test the premise that steric hindrance prevents protonation, the structure of 1-deaza-FMN-flavodoxin, with a stable CH(1) replacing N(l), was determined.61 In oxidized 1-deaza-FMN flavodoxin, atoms of residues 88 to 90 are displaced to make room for the hydrogen at C(l) (Figure 14) and the peptide unit connecting Tyr-88 and Gly-89 rotates to point NH toward O(2). The association constants for binding oxidized 1 -deaza-FMN to apoflavodoxins from M. elsdenii or from C. beijerinckii are smaller, by factors of about 100, than for binding of FMN. This difference includes not only perturbation of the structure, but also loss of hydrogen bond interactions at N(l) and differences in free energy of desolvation of 1-deaza-FMN vs. FMN. Thus the change in Ka, amounting to about 2.5 kcal/mol, cannot be equated to the cost of the structural perturbation, but it is reasonable to assume that deformation of the structure to accommodate a proton at position (1) is energetically unfavorable. B. ESTIMATES OF THE MAGNITUDE OF ELECTROSTATIC INTERACTIONS Measurements of the redox potentials of 1-deaza-FMN flavodoxin from M. elsdenii, at a series of pH values, afford an estimate of the magnitude of electrostatic interactions between the reduced flavin analogue and the protein.61 In reduced 1-deaza-FMN flavodoxin, a redoxlinked pK of 7.6 is determined from measurements of the sq/red equilibrium and large pH *

Plate 1 and 2 follow page 454.

454

Chemistry and Biochemistry of Flavoenzymes

FIGURE 14. Structural changes resulting from the binding of 1-deaza-FMN.61 Positive densities in the (|FdF|V|-|FFlv|) map are drawn with solid contours; negative densities are dashed. Coordinates for residues 89 to 92 of 1-deaza-FMN flavodoxin (labeled 1) are compared with coordinates for FMN flavodoxin (labeled 2). The view is approximately along the peptide plane of residues 88 to 89, displaying the shift of this peptide toward O(2), which accommodates the hydrogen at C(1)H of the FMN analogue by moving the peptide hydrogen approximately 0.6 A to 0.7 A. The structural perturbation is propagated to residue 90. Atoms of FMN flavodoxin are labeled; the flavin coordinates are those for 1-deaza-FMN.

dependent changes in the optical spectrum of the bound flavin analog are consistent with titration of a group with a pK of 7.5. The redox-linked group is thus assumed to be 1-deazaflavin. The corresponding ionization of free 1-deaza-FMN occurs at pH 5.6, and from the pK shift one estimates a value of approximately 2.6 kcal/mol for unfavorable interactions between the protein and a negatively charged flavin. Although 1-deaza-FMN is not a perfect mimic of the electronic structure of FMN, the charge on reduced 1-deaza-FMN is similarly distributed in the pyrimidine ring.82 In comparison, the behavior of reduced FMN flavodoxin from M. elsdenii is anomalous: bound reduced FMN is anionic at low pH, despite the negative and partly apolar environment (Figure 15). Combined optical and NMR measurement establish that the pK is shifted well below the value of 6.5 for free FMN; the optical spectrum of the reduced anionic FMN in M. elsdenii flavodoxin remains unchanged down to pH 5.0. Hence the pK of 5.8 which is observed in measurements of the redox potential cannot be assigned to the flavin but must represent the ionization of some other group. These observations have been incorporated in a model that invokes repulsive electrostatic interactions between individual residues and the reduced flavin ring as contributors to modulation of the sq/red potential. Glu-60 in M. elsdenii flavodoxin or Glu-59 in C. beijerinckii flavodoxin, the ionizable residue closest to the flavin ring, is proposed to be the critical redox-linked group (Scheme 2). It is estimated that the pK of this glutamate increases by two to three pH units when flavodoxin semiquinone is reduced,61 corresponding to an interaction that is 2.6 to 3.9 kcal in magnitude. Free energy effects of this size represent a significant fraction of the total perturbation of the sq/red potential. The properties of a Glu-59—>Gln mutant of C. beijerinckii flavodoxin will provide a test of the roles of this glutamate that are postulated in Scheme 2. The contributions of more distant acidic residues (Table 7) can also be examined by mutagenesis techniques.

PLATE 1. Stereo photograph of the FMN binding site in flavodoxin from C. beijerinckii. Surfaces have been drawn around the protein atoms with the atoms color coded as follows: carbon and sulfur, green; oxygen, red; nitrogen, dark blue. Solvents are not represented in this drawing. The sequences comprising the binding pocket are 7S-G-T-G-N-T, 54S-A-M-G-D-E, 87S-G-Y-G-W-G, and 119N-E.

PLATE 2. Stereo photograph showing the distribution of acidic residues surrounding the flavin in flavodoxin from C. beijerinckii. Each glutamate or aspartate is highlighted in cyan against the blue of the remainder of the model.

Volume HI

SCHEME 2. A model for redox-linked proton binding by fully reduced M. elsdenii flavodoxin, in which the proton acceptor, with a pK of 5.8, is assumed to be Glu-60 (Glu-59 in C. b. flavodoxin). Free flavin species are on the left and flavodoxin species on the right. NMR31 and optical spectra61 show that the pK of 5.8 is not associated with formation of protein-bound FMNH2. At neutral pH, FMNH~ is bound to the Glu~ species, resulting in charge repulsion. At lower pH, the potential for the sq/red equilibrium in flavodoxin depends on pH because of the linked protonation of Glu-60. For Glu-60 to be the proton acceptor, its pK must increase significantly when flavodoxin semiquinone is reduced.

FIGURE 15. A stereo drawing of the isoalloxazine binding site in the crystal structure of reduced C. beijerinckii flavodoxin. The view is of the re face of the flavin ring, looking from the interior of the protein. The ribityl side chain has been truncated at O2'. The drawing includes protein sequences 55A-M-G-D-E and 88Y-G-W-G. Calculations34-35 have shown that much of the net charge of - 1 is distributed on the heteroatoms of the isoalloxazine ring (Table 6). The contacts of some of these atoms with the protein are illustrated here: N(l) and O(2) interact with the dipoles of NH-89 and NH-91; O(4) hydrogen bonds to NH-59 but is only 3.4 A from O-57; 59-COO" adjoins N(3)H and solvents (large open circles). (Reproduced from Ludwig, M. L., Schopfer, L., Metzger, A. L., Pattridge, K. A., and Massey, V., Biochemistry, 29, 10364, 1990. With permission.)

455

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Chemistry and Biochemistry of ¥ lav o enzymes

Electrostatic interactions between the FMN phosphate and the flavin ring have been considered to be critical for control of the sq/red potential.101 Linkage of the phosphate ionization and the sq/red potential should be reflected in shifts of the phosphate pKs, but the pKs of the bound FMN phosphate have not been determined. 31P NMR spectroscopy has established that the bound phosphate group is the dianion above pH 6.0 in oxidized flavodoxin from M. elsdenii,102 and is dianionic at pH 8.0 in C. beijerinckii, and D. vulgaris flavodoxins.63 The only measurements on reduced flavodoxins are at the relatively high pH of 8.0, where the phosphate is the dianion.63 Interactions between the flavin ring and the phosphate, which in C. beijerinckii flavodoxin are separated by a minimum distance of 6.56 A (OP to C(9)), deserve further experimental study. Other experiments have explored the effects of phosphate groups on the redox potentials. One was a study of the interactions of the 3'-5' bisphosphate analogue of FMN with flavodoxins fromM. elsdenii and from D. vulgaris.*4 The bound 3' phosphate is monoanionic at pH 8.0 in the M. elsdenii complex, indicating significant electrostatic interaction between the 3' phosphate and the remainder of the flavoprotein. However, the redox potential for the sq/red equilibrium at pH 8.0 decreases by only 20 mV in the presence of the partly ionized 3' substituent. In A. vinelandii flavodoxins with covalently-linked phosphodiester substituents,103 the sq/red potentials were found to be lower by about 50 mV than the potentials of their dephospho analogues.53 C. A REVIEW OF THE ENERGETICS OF RING BENDING IN REDUCED FLAVINS In the 1970s, folded or bent conformations were thought to be preferred by reduced isoalloxazines. The crystal structures of ring-substituted reduced flavins were taken as evidence that a folded conformation is favored for unsubstituted reduced flavins. Model compounds, such as 9-Br-l,3,7,8,10-pentamethyl 1,5-dihydroisoalloxazine or its 5-acetyl analogue, are bent around the N(10)-N(5) axis,104 with interplanar angles as large as 35°. The current view is that steric (periplanar) crowding by the 1,5, and 9 substituents in these model compounds is responsible for ring folding, and that unsubstituted reduced flavins are nearly planar. The key experiments which determined the conformation of reduced flavins in solution were measurements of the chemical shifts of 15N and 13C in a series of model compounds, including four whose crystal structures had been determined.33 The chemical shifts depend on Tr-electron densities, which are influenced not only by bond hybridization, but also by hydrogen bonding, and in the case of reduced flavins, deprotonation. However, comparisons of spectra and chemical shifts measured in apolar and aqueous solvents, and correlation of the data from both 15N and 13C nuclei, were consistent with a nearly planar conformation for reduced flavins in polar solvents, as determined from estimates of the endocyclic angles at N(5) and N(10). This conclusion relies in part on interpretation of the chemical shifts of 15 N(10) in the FMNH~ species as reflecting the sp2 character of bonding at this atom. Moonen and Muller105 also reexamined the mobility of reduced flavins, both free and bound to M. elsdenii flavodoxin. Earlier NMR studies106 had suggested a large barrier to butterfly motion in reduced flavins, implying that the planar form was high in energy. Moonen and Muller105 concluded that the barrier to ring inversion in free flavins is in fact small. Recent calculations estimate the barrier to formation of the planar conformation in reduced flavins to be approximately 2 kcal/mol,82 in agreement with the spectroscopic studies of Muller and co-workers. There is no evidence for internal motion of the flavin in its protein site on the subnanosecond or nanosecond time scale: the NMR relaxation parameters correspond to those for the protein itself.105 Fluorescence measurements11-107 offer an alternative approach to study the motion of the protein-bound flavin on very short time scales. Thus the conformation of the reduced flavin ring is not altered in major ways by

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FIGURE 16. The phosphate binding site in C. beijernickii and A. nidutans flavodoxins, illustrating the conservation of the structure at residues 7-12 (C. beijerinckii numbering). The two structures were aligned by matching Ca atoms in the sheet and the first helix. The phosphate is at the center, with its terminal oxygen atoms enlarged. Residues shown for C. beijerinckii flavodoxin (open bonds) are 7S-G-T-G-N-T and Ser-55; residues for A. nidulans flavodoxin (labeled with a trailing A) are T-Q-T-G-V-T, Pro-55, and Trp57. Backbone nitrogens of the sequence 7 to 12 in clostridial flavodoxin are filled to emphasize the series of NH dipoles encircling the phosphate. The indole NH of Trp-57A occupies a position similar to that of the water (•) bound at OP(I) in clostridial flavodoxin.

attachment to apoflavodoxins. Although some contributions from ring strain and restricted mobility may appear in the net free energy of binding of reduced FMN to the apoproteins, these parameters are probably not the dominant factors that determine the decreased affinity of the protein for reduced FMN, relative to the affinity for FMN semiquinone.

VI. SOME COMPARISONS OF C. beijerinckii WITH A. nidulans AND OTHER FLAVODOXINS In Section II we presented alignments of C. beijerinckii, A. nidulans, and D.vulgaris flavodoxins, based on three-dimensional superpositions of the Ca coordinates (Figure 2). These three flavodoxins were chosen for comparison because they exhibit intriguing structural variations in the regions which bind the isoalloxazine moiety of FMN. However, we will not attempt to correlate the oxidation-reduction behavior of all three flavodoxins with their structures, but instead will concentrate almost exclusively on a comparison of the flavinbinding sites of C. beijerinckii and A. nidulans flavodoxins, both studied in our laboratory. This comparison alone leads us to suggest that there will be qualitative and well as quantitative differences in the mechanisms deployed by various flavodoxins to control oxidation-reduction potentials. Each of the major flavin-binding subsites is considered in turn: the phosphate-binding loop, the turn which is at residues 56 to 60 in C. beijerinckii flavodoxin, and the chain following residue 88. A. THE PHOSPHATE BINDING LOOP The phosphate binding sites are very similar in conformation (Figure 16), and appear to be a constant feature that achors the FMN to the protein. In particular, the interactions of phosphate oxygens with main-chain peptides and side-chain hydroxyls in the residues preceding the first helix are remarkably well conserved. However, in A. nidulans and other flavodoxins, including M. elsdenii flavodoxin, a proline replaces the serine 54 that contacts

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FIGURE 17. The turn at residue 56T-W-N-V-G in A. nidulans flavodoxin. The view is from the N(10) edge of the flavin ring, with the same perspective as Figure 18. When the structure is aligned with C. beijerinckii flavodoxin by matching the parallel sheets and first helices, the turns do not superimpose because they are attached somewhat differently to the core of the molecule (see Figure 1). However, the local folding and the orientation of the backbone with respect to the flavin are similar in both structures.

the phosphate in C. beijerinckii flavodoxin (Figure 19). In A. nidulans as in several other species, tryptophan is the residue touching the re or inner face of the flavin ring. Trp-57 hydrogen bonds via its indole NH to the phosphate OP(I), replacing the interaction of this phosphate oxygen with solvent. These substitutions provide a less polar environment for the charged FMN phosphate in A. nidulans flavodoxin, and may affect the contribution of phosphate interactions to the free energy of binding. B. THE ISOALLOXAZINE BINDING SITE: THE BEND NEAR RESIDUE 60 We have postulated that the conformational equilibrium involving residues 57 to 58 is important in control of the redox potentials of C. beijerinckii flavodoxin (Section IV). This is not the only flavodoxin in which the conformation of the bend or loop region is linked to the redox state of the flavin. X-ray structures of A. nidulans113 and D. vulgaris17 flavodoxins provide evidence for changes in conformation in each of these molecules; preliminary NMR data for oxidized M. elsdenii flavodoxin47 indicate some, as yet undefined, alterations in the turn corresponding to residues 56 to 60 in C. beijerinckii flavodoxin. The turn at residues 57 to 61 in A. nidulans flavodoxin is shown in Figure 17. The Asn58-Val-59 peptide is trans in this crystal structure, but otherwise the main chain conformation resembles the bend in C. beijerinckii flavodoxin. Studies at 3.0 A resolution indicate changes that are consistent with the formation of a hydrogen bond between O-58 and N(5)H in the semiquinone form, 113 but in the fully reduced A. nidulans flavodoxin the structure reverts to a conformation resembling that found in the oxidized form.114 Thus although the backbone fold in this region is similar to C. beijerinckii flavodoxin, the stability of the bend conformers differs, as does the ox/sq potential, which is 129 mV lower than that of C. beijerinckii flavodoxin. These properties seem to be influenced by the amino acid sequence of the bend, G-D-E in C. beijerinckii flavodoxin vs. N-V-G in A. nidulans flavodoxin (Figure 19), although differences in the main chain-flavin contacts may also affect the energetics. The structure of D. vulgaris flavodoxin presents interesting contrasts to the structures of C. beijerinckii and A. nidulans flavodoxins. In particular, the chain reversal near position 60 is elongated by two residues (Figures 2 and 19) and adopts a different conformation than found in clostridial flavodoxin. However, the ox/sq potential of —103 mV is very similar to the potential observed in C. beijerinckii flavodoxin, and the loop sustains the same kind of peptide inversion observed in clostridial flavodoxin. The Gly-61-Asp-62 peptide rotates

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FIGURE 18. Comparison of the sequence 88Y-G-W-G-D in C. beijerinckii flavodoxin (open bonds with a trailing C) with the sequence 89G-D-Q-V-G-Y-S-D-N-F in A. nidulans flavodoxin (filled bonds with a trailing A). A total of 5 residues is inserted in A. nidulans flavodoxin. The viewpoint and conventions are as in Figure 17. The side chains have been removed except for the aromatic residues covering the si face of the flavin and the aromatic residues at 88C and 98A. The stacking of Tyr-94A is seen to be more perfect than that of Trp-90C. This drawing provides a good view of the unusual conformation of residues 88 to 91 in C. beijerinckii flavodoxin.

to permit hydrogen bonding with N(5)H in both the semiquinone and reduced forms of D. vulgaris flavodoxin.17 Mutations in the 50s bend or loop in all three flavodoxins alter the ox/sq potentials: substitutions G57A, G57D, or G57N in C. beijerinckii flavodoxin,92 N58G in A. nidulans flavodoxin,114'116 and G61N in D. vulgaris flavodoxin115 all change the potentials by 40 to 100 mV. The higher potential is associated with the presence of glycine in every case. These initial observations with mutants lend further support to the idea that the amino acid sequence can affect the potentials by influencing conformational energies. C. THE ISOALLOXAZINE BINDING SITE: THE STRUCTURE NEAR RESIDUE 90 The flavin-binding subsite which begins at residue 87 in C. beijerinckii flavodoxin differentiates C. beijerinckii flavodoxin and its cousin, M. elsdenii flavodoxin, from A. nidulans, D. vulgaris, and other flavodoxins. Only two features of this region are preserved in both C. beijerinckii and A. nidulans flavodoxins: the interaction of a backbone NH with the N(l) and O(2) isoalloxazine atoms, and the contact between an aromatic residue and the si face of the flavin ring. These interactions are not quite identical in the two structures; the aromatic ring is better stacked against the flavin in A. nidulans flavodoxin, and the 90 to 91 peptide plane points more toward O(2). However, the remainder of the region is highly divergent (Figure 18). The first three of five residues inserted in A. nidulans flavodoxin participate in an a-like turn at positions 91 to 94 and the remaining inserts at 95 to 96 are part of a p-turn that precedes main chain contacts with isoalloxazine atoms N(3) and O(2) (Figure 18). One effect of the insertions and the backbone-flavin hydrogen bonds is to bury the pyrimidine end of the flavin more deeply in the protein in the A. nidulans structure. No bound water is found near O(2), nor do solvents penetrate to the vicinity of N(3) and O(4) as they do in C. beijerinckii flavodoxin (Figure 8). The elimination of water from the

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pyrimidine end of the bound flavin may be a crucial structural factor that affects the properties of flavodoxins with inserts in the 90s region. Concomitant with displacement of solvent is the replacement of an acidic side chain at N(3), Glu-59 in C. beijerinckii flavodoxin, by backbone atoms. In summary, species differentiation in this part of the chain preserves the geometry at N(1)/O(2) that is thought to be responsible for the low pK of the reduced isoalloxazine ring in flavodoxins, but varies the polarity and charge distribution of the flavin environment. The presence of extra residues and different substructures in the 90s region is probably responsible for some of the spectral differences which distinguish clostridial and M. elsdenii flavodoxins from A. nidulans, D. vulgaris and other flavodoxins.' -42 The ability of the latter flavodoxins to bind riboflavin also correlates with the insertion of residues in the 90s subsite. Replacement of the N(3): glutamate interaction has implications for the electrostatic effects which have been a focus of this chapter. Glu-59 of C. beijerinckii flavodoxin, postulated to be a redox-linked ionizable group (Scheme 2), has no structural counterpart in A. nidulans or related flavodoxins. To generate specific electrostatic interactions, flavodoxins for which A. nidulans is the prototype must therefore utilize some alternative to the mechanism proposed in Scheme 2. Acidic residues in the neighborhood of the isoalloxazine ring in A. nidulas and C. beijerinckii flavodoxins are compared in Table 7, which shows that no acidic side chains contact FMN in A. nidulans flavodoxin. The acidic groups closest to the flavin ring are Asp-90, a residue which is conserved in long-chain flavodoxins, and Glu-146 (Figure 19). Asp-144 is buried in the protein; Asp-90 and Asp-100 are only partly accessible to solvent. The spatial distribution of these acidic groups, shown in Table 7 and Figure 1, supports the idea the several residues not in direct contact with the flavin may cooperate to provide electrostatic interaction with the reduced flavin anion in A. nidulans flavodoxin and its structural relatives. The decreased polarity of the N(1)-O(2) environment would enhance the effects of these charges. Preliminary studies of the A. nidulans mutant Asp-9CM>Asn show that the charge at this position does lower the redox potential, but by only about 50 mV:114'116 this single interaction between Asp-90 and FMN cannot by itself account for the very low sq/red potential of A. nidulans flavodoxin. Analogous mutations at other acidic residues114 of A. nidulans flavodoxin, singly and in combination, will assess the functional roles of the acidic residues listed in Table 7.

VII. PERSPECTIVES The differences between the flavin binding sites of C. beijerinckii and D. vulgaris flavodoxins were an early but surprising finding that has been reinforced by structural data from additional flavodoxins.15'16 Comparisons of natural variants have suggested that the redox potentials of flavodoxins are controlled by somewhat different structural devices in different species. NMR spectroscopy is detecting differences among flavodoxins at the level of the electronic structure of the cofactor,63 and other techniques like Raman and fluorescence spectroscopy, which we have not attempted to review in detail (see Reference 11) are sensitive to variations in flavin-protein interactions. Quantitative description of the effects of the protein environment on the properties of the flavin cofactor remains a challenge. Understanding why full reduction of flavodoxins is thermodynamically unfavorable has been complicated by the difficulties in accurate computation of electrostatic free energies in proteins,117 and by the complexity of the electronic structure of the flavin nucleus itself. 82 - 118 - 119 Fortunately, site-directed mutagenesis is beginning to provide some estimates of the roles of conformational equilibria and electrostatic interactions in determining the redox potentials of flavodoxins. The wealth of experimental data from studies of mutants and species variants of flavodoxins may in turn be used to test the application of computational techniques to the study of flavin-protein interactions in these model flavoproteins.

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FIGURE 19. Alignments of segments of sequences from several flavodoxins. This figure predicts the structural equivalences of residues from a series of flavodoxins, some of whose structures are not known. Numbering is from A. nidulans flavodox'm. Key features in the alignment of the 50s region are the hydrophobic side chain (Trp57, Met or Leu) contacting the re face of the flavin, and glycine residues either one or three positions beyond the hydrophobic residue. Flavodoxin from D. vulgaris has two extra residues near position 60, and in C. crispus flavodoxin a few residues appear to be inserted, not at the same site as in D. vulgaris flavodoxin, but instead, just beyond the bend. In the 90s region, glycine in clostridial and M, elsdenii flavodoxin is aligned with Asp in the other sequences, because these are the positions which hydrogen bond to N( 1) of the flavin ring. The aromatic residues which contact the si face of the flavin (Tyr-94 in A. nidulans) are also aligned, as are the aromatic residues corresponding to position 98 of A. nidulans flavodoxin. The third group of sequences is from long chain flavodoxins which share a cluster of acidic residues at turns just beyond the final strand of (3-sheet.

ACKNOWLEDGMENTS Structural studies of C. beijerinckii and A. nidulans flavodoxins have been supported by NIH Grants GM 16429 and GM 08270, and by the San Diego Supercomputer Center. We are pleased to acknowledge continuing collaborations with Drs. V. Massey, D. B. Ballou, and R. P. Swenson, and thank Drs. W. Watt, K. D. Watenpaugh, and J. Vervoort for communication of data prior to publication. The comments and assistance of K. A. Pattridge and A. L. Metzger are much appreciated. The coordinates from refinement of C. beijerinckii flavodoxin are being deposited in the Brookhaven Data Bank.

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REFERENCES 1. Mayhew, S.G. and Ludwig, M. L., Flavodoxins and electron-transferring flavoproteins, The Enzymes, 57, 1975. 2. Knight, E., Jr. and Hardy, R. W. F., Isolation and characteristics of flavodoxin from nitrogen-fixing Clostridium pasteranium, J, Bioi. Chem., 241, 2752, 1966. 3. Smillie, R. M., Isolation of two proteins with chloroplast ferredoxin activity from a blue-green alga, Biochem. Biophys. Res. Comm.t 20, 621, 1965. 4. Nieva-Gomez, D., Roberts, G. P., Klevickis, S., and Brill, W. J., Electron transport to nitrogenase in Klebsiellapneumoniae, Proc. Nat. Acad, Sci. U.S.A., 77, 25555, 1980. 5. Deistung, J., Cannon, F. C., Cannon, M. C., Hill, S., and Thorneley, R. N., Electron transfer to nitrogenase in Klebsiella pneumoniae, ni/F gene cloned, and the gene product, a flavodoxin, purified, Biochem. J.t 231, 743, 1985. 6. Banerjee, R. V. and Matthews, R. G., Cobalamin-dependent methionine synthase, FASEB J., 4, 1450, 1990. 7. Fujii, K. and Huennekens, F. M., Activation of methionine synthetase by a reduced triphosphopyridine nucleotide-dependent flavoprotein system, J. BioL Chem., 249, 6745, 1974. 8. Knappe, J. and Blasckowski, H. P., Pyruvate formate-lyase from Escherichia coli and its activation system, Methods Enzymot,, 61, 508, 1975. 9. Blasckowski, H. P., Neuer, G., Ludwig-Festl, M., and Knappe, J., Routes of flavodoxin and ferredoxin reduction in Escherichia coli. CoA-acylating pyruvate:flavodoxin and NADPH: flavodoxin oxidoreductases participating in the activation of pyruvate formate-lyase, Eur. J, Biochem., 123, 563, 1982. 10. Fitzgerald, M. P., Husain, A., and Rogers, L. J., A constitutive flavodoxin from a eukaryotic alga, Biochem. Biophys. Res. Comm., 81, 630, 1978. 11. Mayhew, S. G. and Tollin, G., General properties of flavodoxins, in Chemistry and Biochemistry of Flavoenzymes, Vol. 3, Muller, R, Ed., CRC Press, Boca Raton, FL, chap. 14, this volume. 12. Burnett, R. M., Darling, G. D., Kendall, D. S., LeQuesne, M. E., Mayhew, S. G., Smith, W. W., and Ludwig, M. L., The structure of the oxidized form of clostridial flavodoxin at 1,9 A resolution. Description of the flavin mononucleotide binding site, J. Bioi. Chem,, 249, 4383, 1974. 13. Watenpaugh, K. D., Sicker, L. C., and Jensen, L. H., The binding of riboflavin-5'-phosphate in a flavoprotein; flavodoxin at 2.0 A resolution, Proc. Nat. Acad. Sci. U.S.A., 70, 3857, 1973. 14. Watenpaugh, K. D., Sieker, L. C., and Jensen, L. H., A crystallographic structural study of the oxidation states of Desulfovibrio vulgaris flavodoxin, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 405. 15. Smith, W. W., Pattridge, K. A., Ludwig, M. L., Petsko, G. A., Tsernoglou, D., Tanaka, M., and Yasanobu, K. T., Structure of oxidized flavodoxin from Anacystis nidulans, J. Mol. Bioi., 165, 737, 1983. 16. Fukuyama, K., Wakabayashi, S., Matsubara, H., and Rogers, L. J., Tertiary structure of oxidized flavodoxin from an eukaryotic red alga Chondrus crispus at 2.35 A resolution, J. Bioi. Chem., 265, 26, 1990, 17. Watt, W., Tulinsky, A., Swenson, R. P., and Watenpaugh, K. D., Comparison of the crystal structures of a flavodoxin in its three oxidation states at cryogenic temperatures, J. Mol. Bioi., 218, 195, 1991. 18. Andersen, R. D., Apgar, P. A., Burnett, R. M., Darling, G. D., LeQuesne, M. E., Mayhew, S. G., and Ludwig, M. L., Structure of the radical form of clostridial flavodoxin: a new molecular mode!, Proc. Natl. Acad. Sci. U.S.A., 69, 3189, 1972, 19. Smith, W. W., Burnett, R. M., Darling, G. D., and Ludwig, M. L., Structure of the semiquinone form of flavodoxin from Clostridium MP, J. Mol. BioL, 117, 195, 1977. 20. Ludwig, M. L., Burnett, R. M., Darling, G. D., Jordan, S. R., Kendall, D. S., and Smith, W. W., The structure of Clostridium MP flavodoxin as a function of oxidation state: some comparisons of the FMNbinding sites in oxidized, semiquinone and reduced forms, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 393. 21. Smith, W. W., Ludwig, M. L., Pattridge, K. A., Tsernoglou, D., and Petsko, G. A., Crystallographic studies of flavodoxins: some correlations between structure and redox potential, in Frontiers of Biological Energetics: From Electron to Tissues, Vol. 2, Dutton, P. L., Leigh, J. S., and Scarpa, A., Eds., Academic Press, New York, 1978, 957. 22. van Mierlo, C. P. M., Vervoort, J., Muller, F., and Bacher, A., A two-dimensional 'H NMR study on Megasphaera elsdenii flavodoxin in the reduced state, Eur. J. Biochem., 187, 521, 1990. 23. van Mierlo, C. P. M., Muller, F., and Vervoort, J., Secondary and tertiary structure characteristics of Megasphaera elsdenii flavodoxin in the reduced state as determined by two-dimensional 'H NMR, Eur. J. Biochem., 189, 589, 1990.

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24. van Mierlo, C. P. M., Lijnzaad, P., Vervoort, J., Muller, F., Berendsen, H. J. C., and de Vlieg, J., Tertiary structure of two-electron reduced Megasphaera elsdenii flavodoxin and some implications, as determined by two-dimensional 'H NMR and restrained molecular dynamics, Eur. J. Biochem., 194, 185, 1990. 25. Stockman, B. J., Westler, W. M., Mooberry, E. S., and Markley, J. L., Flavodoxin from Anabaena 7120: uniform nitrogen-15 enrichment and phosphorus-31 NMR investigations of the flavin mononucleotide binding site in the reduced and oxidized states, Biochemistry, 27, 136, 1988. 26. Stockman, Brian J., Krezel, A. M., and Markley, J. L., Hydrogen-1, carbon-13, and nitrogen-15 NMR spectroscopy of Anabaena 7120 flavodoxin: assignment of p-sheet and flavin binding site resonances and analysis of protein-flavin interactions, Biochemistry, 29, 9600, 1990. 27. Vervoort, J., Muller, F., Le Gall, J., Bacher, A., and Sedlmaier, H., Carbon-13 and nitrogen-15 nuclear-magnetic-resonance investigation onDesulfovibrio vulgaris flavodoxin, Eur, J. Biochem., 151, 49, 1985. 28. Clubb, R. and Wagner, G., private communication. 29. Draper, R. D. and Ingraham, L. L., A potentiometric study of the flavin semiquinone equilibrium, Arch. Biochem. Biophys., 125, 802, 1968. 30. Mayhew, S. G., Studies on flavin binding in flavodoxins, Biochim. Biophys. Acta, 235, 289, 1971. 31. Mayhew, S. G., Foust, G. P., and Massey, V., Oxidation-reduction properties of flavodoxin from Peptostreptococcus elsdenii, J. Biol. Chem., 244, 803, 1969. 32. Anderson, R. F., Energetics of the one-electron reduction steps of riboflavin, FMN and FAD to their fully reduced forms, Biochim. Biophys. Acta, 772, 158, 1983. 33. Moonen, C. T. W., Vervoort, J., and Muller, F., Reinvestigation of the structure of oxidized and reduced flavin: carbon-13 and nitrogen-15 nuclear magnetic resonance study, Biochemistry, 23, 4859, 1984. 34. Hall, L. H., Orchard, B. J., and Tripathy, S. K., The structure and properties of flavins: molecular orbital study based on totally optimized geometries. II. Molecular orbital structure and electron distribution, Int. J. Quant. Chem,, XXXI, 217, 1987. 35. Teitell, M. F. and Fox, J. L., MO study of flavin reduction, Int. J. Quant, Chem., XXII, 583, 1982. 36. Laudenbach, D. EM Reith, M. E., and Straus, N. A., Isolation, sequence analysis, and transcriptional studies of the flavodoxin gene from Anacystis nidulans R2, J. Bacteriol., 170, 258, 1988. 37. Tanaka, M., Mitsuru, H., and Yasunobu, K. T., The amino acid sequence of the Clostridium MP flavodoxin, /. Biol. Chem., 249, 4393, 1974. 38. Dubourdieu, M., Le Gall, J., and Fox, J. L., The amino acid sequence of Desulfovibrio vulgaris flavodoxin, Biochem. Biophys. Res. Commun., 52, 1428, 1973. 39. Krey, G. D., Vanin, E. F., and Swenson, R. P., Cloning, nucleotide sequence, and expression of the flavodoxin from Desulfovibrio vulgaris (Hildenborough), /. Biol. Chem,, 263, 15436, 1988. 40. Drummond, M. H., The base sequence of the ni/F gene of Klebsiella pneumoniae and homology of the predicted amino acid sequence of its protein product to other flavodoxins, Biochem. J., 232, 891, 1985. 41. Laudenbach, D. E., Straus, N. A., Pattridge, K. A., and Ludwig, M. L., Sequence and structure of Anacystis nidulans flavodoxin: comparisons with flavodoxins from other species, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 249. 42. Edmondson, D. E. and Tollin, G., Circular dichroism studies of the flavin chromphore and of the relation between redox properties and flavin environment in oxidases and dehydrogenases, Biochemistry, 10, 113, 1971. 43. I)"Anna, J. A. and Tollin, G., Studies of flavin-protein interaction in flavoproteins using protein fluorescence and circular dichroism, Biochemistry, 11, 1073, 1972. 44. Vervoort, J., van Berkel, W. J. H., Mayhew, S. G., Miiller, F., Bacher, A., Nielsen, P., and Le Gall, J., Properties of the complexes of riboflavin 3',5'-bisphosphate and the apoflavodoxins from Megasphaera elsdenii and Desulfovibrio vulgaris, Eur. J. Biochem., 161, 749, 1986. 45. Ballou, D. B. and Ludwig, M. L., unpublished. 46. Smith, W. W., Crespi, H. L., Entsch, B., Ludwig, M. L., and Nordman, C. E., Crystallographic characterization of flavodoxin from Anacystis nidulans, J. Mol. Biol., 94, 123, 1975. 47. van Mierlo, C. P. M., A Two-Dimensional 'H NMR Study on the Apoflavodoxin of Megasphaera elsdenii, Ph.D. dissertation, Department of Biochemistry, Agricultural University, Wageningen, The Netherlands, 1990. 48. Paulsen, K. E., Stankovich, M. T., Stockman, B. J., and Markley, J. L., Redox and spectral properties of flavodoxin from Anabaena 7120, Arch. Biochem. Biophys., 280, 68, 1990. 49. Entsch, B. and Smillie, R. M., Oxidation-reduction properties of phytoflavin, a flavoprotein from bluegreen algae, Arch. Biochem. Biophys., 151, 378, 1972. 50. Sykes, G. A. and Rogers, L. J., Redox potentials of algal and cyanobacterial flavodoxins, Biochem. J., 217, 845, 1984.

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51. Deistung, J. and Thorneley, R. N., Electron transfer to nitrogenase, characterization of flavodoxin from Azotobacter chroococcum and comparison of its redox potential with those of flavodoxins from Azotobacter vinelandii and Klebsiella pneumoniae (rc(/F-gene products), Biochem. J., 239, 69, 1986. 52. Yoch, D. C., The electron transport system in nitrogen fixation by Azotobacter. IV. Some oxidationreduction properties of azotoflavin, Biochem, Biophys. Res. Commun., 49, 335, 1972. 53. Taylor, M. F., Boylan, W. H., and Edmondson, D. E., Azotobacter vinelandii flavodoxin: purification and properties of the recombinant, dephospho form expressed in Escherichia coli, Biochemistry, 29, 6911, 1990. 54. Barman, B. G. and Tollin, G., Ravine-protein interactions in flavoenzymes, thermodynamics and kinetics of reduction of Azotobacter flavodoxin, Biochemistry, 11,4755, 1972. 55. Klugkist, J., Voorberg, J., Haaker, H., and Veeger, C., Characterization of three different flavodoxins from Azotobacter vinelandii, Eur. J. Biochem., 115, 33, 1986. 56. Mayhew, S. G., Properties of two clostridial flavodoxins, Biochim. Biophys. Acta, 235, 276, 1971. 57. Dubourdieu, M., LeGall, J., and Favaudon, V., Physicochemical properties of flavodoxin from Desulfovibrio vulgaris, Biochim. Biophys. Acta, 376, 519, 1975. 58. Vetter, H., Jr. and Knappe, J., Flavodoxin and ferredoxin of Escherichia coli, Hoppe-Seyler'sZ. Physiol. Chem., 352, 433, 1971. 59. Crespi, H. L., Morris, J. R., Bays, J. P., and Katz, J. J., ESR and NMR studies with deuterated flavodoxin, Ann. N.Y. Acad. Sci., 222, 800, 1973. 60. Schopfer, L. M., Ludwig, M. L., and Massey, V., A working proposal for the role of the apoprotein in determining the redox potential of the flavin in flavoproteins: correlations between potential and flavin pKs, in Flavins andFlavoproteins 1990, Curti, B., Zanetti, G., andRonchi, S., Eds., Walter de Gruyter, Berlin, 1991, 399. 61. Ludwig, M. L., Schopfer, L. M., Metzger, A. L., Pattridge, K. A., and Massey, V., Structure and oxidation-reduction behavior of 1-deaza-FMN flavodoxins: modulation of redox potentials in flavodoxins, Biochemistry, 29, 10364, 1990. 62. Stankovich, M. T., An anaerobic cell for studying the spectral and redox properties of flavoproteins, Anal. Biochem., 109, 295, 1980. 63. Vervoort, J., Miiller, F., Mayhew, S. G., van den Berg, W. A. M., Moonen, C. T. W., and Bacher, A., A comparative carbon-13, nitrogen-15, and phosphorus-31 nuclear magnetic resonance study on the flavodoxins from Clostridium MP, Megasphaera elsdenii, and Azotobacter vinelandii, Biochemistry, 25, 6789, 1986. 64. Smith, W. W., The Crystal Structure and Refinement of Clostridium MP Flavodoxin in the Semiquinone State and Some Comparisons of the Protein in the Oxidized, Semiquinone, and Fully Reduced States, Ph.D. dissertation, University of Michigan, 1977. 65. Hendrickson, W. A., Stereochemically restrained refinement of macromolecular structures, Methods Enzymol, 115, 252, 1985. 66. Eren, M. and Swenson, R. P., Chemical synthesis and expression of a synthetic gene for the flavodoxin from Clostridium MP, J. Biol. Chem., 264, 14874, 1989. 67. Kabsch, W. and Sander, C., Dictionary of protein secondary structure: pattern recognition of hydrogenbonded and geometrical features, Biopolymers, 22, 2577, 1983. 68. Tanaka, M., Mitsuru, H., and Yasunobu, K. T., The primary structure of Peptostreptococcus elsdenii flavodoxin, /. Biol. Chem., 248, 4354, 1973. 69. Tanaka, M., Mitsuru, H., and Yasunobu, K. T., Correction of the amino acid sequence of Peptostreptococcus elsdenii flavodoxin, J. Biol. Chem., 249, 4397, 1974. 70. Rossmann, M. G., Liljas, A., Branden, C.-L, and Banaszak, L. J., Evolutionary and structural relationships among dehydrogenases, The Enzymes, 11A, 62, 1975. 71. Richardson, J. S., The anatomy and taxonomy of proteins, Adv. Protein Chemistry, 34, 167, 1981. 72. Richards, F. M., Calculation of molecular volumes and areas for structures of known geometry, Methods Enzymol, 115, 440, 1985. 73. Milner-White, E. J. and Poet, R., Four classes of (5-hairpins in proteins, Biochem. J., 240, 289, 1986. 74. Jorgensen, W. L. and Gao, J., Cis-trans energy difference for the peptide bond in the gas phase and in aqueous solution, J. Am. Chem. Soc., 110, 4212, 1988. 75. Schnur, D. M., Yuh, Y. H., and Dal ton, D. R., A molecular mechanics study of amide conformations, J. Org. Chem., 54, 3779, 1989. 76. Stewart, D. E,, Sarkar, A., and Wampler, J. E., Occurrence and role of cis peptide bonds in protein structures, J. MoL Biol, 214, 253, 1990. 77. Miiller, F., Hemmerich, P., Ehrenberg, A., Palmer, G., and Massey, V., The chemical and electronic structure of the neutral flavin radical as revealed by electron spin resonance spectroscopy of chemically and isotopically substituted derivatives, Eur. J. Biochem., 14, 185, 1970. 78a. Vervoort, J., Muller, F., O'Kane, D. J., Lee, J., and Bacher, A., Bacterial luciferase: a carbon-13, nitrogen-15,, and phosphorus-31 nuclear magnetic resonance investigation, Biochemistry, 25, 8067, 1986.

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78. Franken, H.-D., Riiterjans, H., and Muller, F., Nuclear-magnetic-resonance investigation of ls N-labeled flavins, free and bound to Megasphaera elsdenii apoflavodoxin, Eur. J. Biochem., 138, 481, 1984. 79. van Schagen, C. G. and Muller, F., A 13C nuclear-magnetic-resonance study of free flavins and Megasphaera elsdenii and Azotobacter vinelandii flavodoxin. 13C-Enriched flavins as probes for the study of flavoprotein active sites, Eur. J. Biochem., 120, 33, 1981. 80. Macheroux, P., Kojiro, C. L., Schopfer, L. M., Chakraborty, S., and Massey, V., 19F NMR studies on 8-fluoroflavins and 8-fluoro flavoproteins, Biochemistry, 29, 2670, 1990. 81. Hall, L. H., Orchard, B. J., and Tripathy, S. K., The structure and properties of flavins: molecular orbital study based on totally optimized geometries, I. Molecular geometry investigations, Int. J. Quant. Chem., XXXI, 217, 195, 1987. 82. Hall, L. H., Bowers, M. L., and Durfor, C. N., Further consideration of flavin coenzyme biochemistry afforded by geometry-optimized molecular orbital calculations, Biochemistry, 26, 7401, 1987. 83. Moonen, C. T. W., Vervoort, J M and Muller, F., Carbon-13 nuclear magnetic resonance study on the dynamics of the conformation of reduced flavin, Biochemistry, 23, 4868, 1984. 84. Wang, M. and Fritchie, C. J., Jr., Geometry of the unperturbed flavin nucleus. The crystal structure of 10-methylisoalloxazine, Acta Cryst., B29, 2040, 1973. 85 Weiner, S. J., Kollman, P. A., Case, D. A., Singh, U. C., Ghio, C., Alagona, G., Profeta, S., Jr., and Weiner, P., A new force field for molecular mechanical simulation of nucleic acids and proteins, /. Am. Chem. Soc., 106, 765, 1984. 86. Roterman, I. K., Lambert, M., Gibson, K. D., and Scheraga, H. A., A comparison of the CHARMM, AMBER, and ECEPP potentials for peptides. II. Maps for AT-acetyl alanine TV-methyl amide: comparisons, contrasts, and simple experimental tests, /. Biomol. Struct. Dyn., 7, 421, 1989. 87. Nicholson, H., Soderlind, E., Tronrud, D. E., and Matthews, B. WM Contributions of left-handed helical residues to the structure and stability of bacteriophage T4 lysozyme, J. Mol. BioL, 210, 181, 1989. 88. Brooks, C. L., Briinger, A., and Karplus, M., Active site dynamics in protein molecules: a stochastic boundary molecular dynamics approach, Biopolymers, 24, 843, 1985. 89. Dudek, M. J. and Scheraga, H. A., Protein structure prediction using a combination of sequence homology and global energy minimization. I. Global energy minimization of surface loops, J. Comp. Chem., 11, 121, 1990. 90. Bruccoleri, R. E. and Karplus, M., Prediction of the folding of short segments by uniform conformational sampling, Biopolymers, 26, 137, 1987. 91. Joseph, D., Petsko, G. A., and Karplus, M,, Anatomy of a conformational change: hinged "lid" motion of the triose phosphate isomerase loop, Science, 249, 1425, 1990. 92. Ludwig, M. L M Pattridge, K. A., Eren, M., and Swenson, R. P., Structural characterization of site mutants of clostridial flavodoxin, in Flavins and Flavoproteins 1990, Curti, B., Zanetti, G., and Ronchi, S., Eds., Walter de Gruyter, Berlin, 1991, 423. 93. Mayhew, S. G. and Massey, V., Studies on the kinetics and mechanism of reduction of flavodoxin from Peptostreptococcus elsdenii by sodium dithionate, Biochim. Biophys. Acta, 315, 181, 1973. 94. van Leeuwen, J. W., vanDijk, C., and Veeger, C., A pulse-radiolysis study or the reduction of flavodoxin from Megasphaera elsdenii by viologen radicals. A conformation change as a possible regulating mechanism, Eur. J. Biochem., 135, 601, 1983. 95. Anderson, R. F., Massey, V., and Schopfer, L. M., Pulse radiolysis studies on flavodoxin, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 279. 96. Ballou, D. B., Ludwig, M. I., and Massey, VM unpublished. 97. Moonen, C. T. W. and Muller, F., On the intermolecular electron transfer between different redox states of flavodoxin from Megasphaera elsdenii, Eur. J. Biochem., 140, 303, 1984. 98 Kordel, J., Forsen, S., Drakenberg, T., and Chazin, W. J M The rate and structural consequences of proline cis-trans isomerization in Calbindin D9k: NMR studies of the minor (c/s-Pro43) isoform and the Pro43Gly mutant, Biochemistry, 29, 4400, 1990. 99. Evans, P. A., Dobson, C. M., Kautz, R. A., Hatful, G., and Fox, R. O., Proline isomerism in staphylococcal nuclease characterized by NMR and site-directed mutagenesis, Nature, 329, 266, 1987. 100. MacKay, D. H. J., Cross, A. J., and Hagler, A. T., The role of energy minimization in simulation strategies of biomolecular systems, in Prediction of Protein Structure and the Principles of Protein Conformation, Fasman, G. D., Ed., Plenum Press, New York, 1989, 317. 101. Moonen, C. T. W., Vervoort, J., and Muller, FM Some new ideas about the possible regulation of redox potentials in flavoproteins, with special reference to flavodoxins, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 494. 102. Moonen, C. T. W. and Muller, F., Structural and dynamic information of the complex of Megasphaera elsdenii apoflavodoxin and riboflavin 5'-phosphate. A phosphorus-31 nuclear rrfagentic resonance study, Biochemistry, 21, 408, 1982.

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103. Edmondson, D. E. and James, T. L., Covalently bound non-coenzyme phosphorus residues in flavoproteins. M P nuclear magnetic resonance studies of Azotobacter flavodoxin, Proc. Nat. Acad. Sci. U.S.A., 76, 3786, 1979. 104. Kierkegaard, P., Norrestam, R., Werner, P., Csoregh, I., Von Glehn, M., Karlsson, R., Leijonmarck, M., Ronnquist, O., Stensland, B., Tillberg, O., and Torbjornsson, L., X-ray structure investigation of flavin derivatives, in Flavins and Flavoproteins, Kamin, H., Ed., University Park Press, Baltimore, 1971, 1. 105. Moonen, C. T. W. and Miiller, F., On the mobility of riboflavin 5'-phosphate in Megasphaera elsdenii by 13C-nuclear magnetic resonance relaxation, Eur. J. Biochem., 133, 463, 1983. 106. Tauscher, L., Ghisla, S., and Hemmerich, P., NMR.-Study of nitrogen inversion and conformation of 1,5 dihydro-isoalloxazines ("reduced flavin"). Studies in the flavin series. XIX., Helv. Chim. Acta, 56, 630, 1973. 107. Visser, A. J. W. G., van Hoek, A., Kulinski, T., and LeGall, J., Time-resolved fluorescence studies of flavodoxin. Demonstration of picosecond fluorescence lifetime of FMN in Desulfovibrio flavodoxins, FEES Lett., 224, 406, 1987. 108. Leonhardt, K. G. and Straus, N. A., Sequence of the flavodoxin gene from Anabaena variabilis, Nucleic Acids Res., 17, 11, 1989. 109. Bennett, L. T., Jacobson, M. R., and Dean, D. R., Isolation, sequencing, and mutagenesis of the nifp gene encoding flavodoxin from Azotobacter vinelandii, J. Biol. Chem., 263, 1364, 1988. 110. Wakabayashi, S., Kimura, T., Fukuyama, K M Matsubara, H., and Rogers, L. J., The amino acid sequence of a flavodoxin from the eukaryotic red alga Chondrus crispus, Biochem. J., 263, 981, 1989. 111. Osborne, C M Chen, L., and Matthews, R. G., Isolation, cloning, mapping, and nucleotide sequencing of the gene encoding flavodoxin in Escherichia coli, J. Bact., 173, 1729, 1991. 112. Curley, G. P. and Voordouw, G., Cloning and sequencing of the gene encoding flavodoxin from Desulfovibrio vulgaris Hildenborough, FEMS J., 49, 295, 1988. 113. Ludwig, M. L., Pattridge, K. A., and Tarr, G., FMN: protein interactions in flavodoxin from A. nidulans, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 253. 114. Luschinsky, C. L., Dunham, W. R., Osborne, C., Pattridge, K. A., and Ludwig, M. L., Structural analysis of fully reduced A. nidulans flavodoxin, in Flavins and Flavoproteins 1990, Curti, B., Zanetti, G. and Ronchi, S., Eds., Walter de Gruyter, Berlin, 1991, 409. 115. Curley, G. P., Carr, M. C., O'Farrell, P. A., Mayhew, S. G., and Voordouw, G., Redox properties of wild-type and mutant flavodoxins from Desulfovibrio vulgaris (Hildenborough), in Flavins and Flavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 429. 116. Hoover, D., Webb, H. and Ludwig, M. L., unpublished. 117. Sharp, K. A. and Honig, B., Electrostatic interactions in macromolecules: theory and applications, in Annual Review Biophysics and Biophysical Chemistry, Vol. 19, Engelman, D. M., Cantor, C. R., and Pollard, T. D., Annual Reviews, Palo Alto, 1990, 301. 118. Dixon, D. A., Branchaud, B., and Lipscomb, W. N., Conformations and electronic structures of oxidized and reduced isoalloxazine, Biochemistry, 18, 5770, 1979. 119. Nishimoto, K., Ab initio MO study of the redox function of flavin, in Biomolecules, Nagata et al., Eds., Japan Sci. Soc. Press, Tokyo/Elsevier 1985, 9.

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Chapter 16

THE BIOCHEMISTRY AND MOLECULAR BIOLOGY OF BACTERIAL BIOLUMINESCENCE Thomas O. Baldwin and Miriam M. Ziegler

TABLE OF CONTENTS I.

Overview A. Bacterial Bioluminescence in Vitro B. Cloning and Sequencing of the Iwc Genes C. Control of lux Gene Expression D. Applications of Bioluminescence

468 469 471 471 475

II.

Enzymology of Bacterial Luciferase A. Substrates, Products and Reaction Stoichiometry B. Reaction Scheme 1. Binding of Reduced Flavin Mononucleotide 2. Binding of Aldehyde 3. Kinetic Mechanism of the Reaction C. Stable and Transient Intermediates D. Postulated Reaction Mechanisms

478 478 479 480 481 483 483 485

III.

Structural Studies of Bacterial Luciferase A. Architecture of the Enzyme B. Amino Acid Sequences of Bacterial Luciferases 1. Interaction Between the a and £ Subunits 2. Regions of Sequence Conservation 3. An Internal Deletion Within the (3 Subunit, or an Imprecise Gene Duplication? 4. Taxonomic Relationships Among the LuciferaseProducing Bacteria C. Residues Contributing to the Active Center 1. The Reactive Thiol Residue 2. Reactive Histidinyl Residue and a-Amino Groups 3. Studies of the Active Center by Limited Proteolysis 4. Location of the Protease Labile Regions Relative to the Active Center 5. Investigation of the Properties of Mutant Forms of Luciferase 6. Location of the Active Center Relative to the Subunit Interface D. Properties of Bacterial Luciferases Having Genetically Fused Subunits E. Folding of the Subunits and Assembly of the Heterodimer 1. Refolding of Luciferase Subunits and Formation of Active Enzyme 2. Formation of Hybrid Luciferases 3. Subunit Dissociation and Reassociation

487 488 488 492 492 493 494 494 495 499 499 500 502 503 506 506 506 507 507

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F. IV.

4. Association of Subunits Folded Independently in Vivo Temperature Sensitive Folding Mutants

507 508

Bacterial Bioluminescence in Vivo A. Accessory Enzymes B. Yellow Fluorescent Protein C. Lumazine Protein D. Nonfluorescent Flavoprotein Regulation of lux Gene Expression A. Structure of the lux Regulon of Vibrio fischeri B. The LuxR Protein: Autoinducer and the Autoinduction Mechanism C. Regulation of Expression of the lux Genes: A Model

510 510 511 512 513 515 515

Addendum

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Acknowledgments

522

References

523

V.

VI.

516 520

I. OVERVIEW The decade of the 1980s has seen a dramatic change in the techniques and strategies available for the study of proteins and enzymes through the development and dissemination of recombinant DNA technology. During the latter half of the 1970s, recombinant DNA technology was utilized in the laboratories of molecular biologists who were primarily responsible for the development of the technology, but the 1980s saw the technology spread from the laboratories of the molecular biologists into the laboratories of protein chemists and enzymologists. It was early in the decade that the first clones of the bacterial luciferase genes were isolated,1"5 and much of the time since then has been devoted to manipulation of the lux genes. Our last review on the subject of bacterial luciferase6 was written in 1980 and covered literature primarily of the 1970s. In the past 10 years, several other thorough reviews have been written, discussing the general biology and physiology of bacterial bioluminescence,7-8 the mechanism of the reaction,9'10 and the genetics and accessory enzymes for bacterial bioluminescence in vivo.11 This review will endeavor to summarize approximately the last 10 years of research on bacterial luciferase, drawing on the older literature only to place the more recent research in proper historical perspective. The cloning of the lux genes and the demonstration that these genes function in a variety of organisms has resulted in a proliferation of literature on the applications of bioluminescence. Many of these studies have added to our understanding of the fundamental properties of the enzyme and the reactions that it catalyzes. However, the applications literature is too broad to permit adequate treatment here. This review, therefore, will concentrate on the fundamental literature with reference to specific applications only when those investigations reveal fundamental features of bacterial luciferase. Since it has been impossible to cover the literature completely, we apologize to the readers of the review for the inevitable (though inadvertent) omissions and shortcomings.

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FIGURE 1. Time course for light emission from the bacterial luciferasecatalyzed reaction. In this assay, a vial containing V. harveyi luciferase in 1 ml of 0.2% bovine serum albumin in phosphate buffer, pH 7.0, with dodecanal (15 jxl of a 0.1% sonicated suspension in water) is placed in a housing above a photomultiplier tube, and at zero time, 1 ml of 50 u>A/ FMNH2 is injected.12 The initial maximum light intensity reached is a measure of the initial velocity of the reaction and is proportional to the luciferase concentration. The inset shows the time course of light emission with different aldehydes on a logarithmic scale, demonstrating the first-order decay of the long-lived intermediate.

A. BACTERIAL BIOLUMINESCENCE IN VITRO Bacterial luciferase is an uncommon flavoprotein in that it employs reduced flavin as a substrate rather than as a tightly bound cofactor. The enzyme, a monooxygenase, catalyzes the reaction of FMNH2, O2, and a long-chain aliphatic aldehyde to yield FMN, the carboxylic acid, and blue-green light. The time course of the bioluminescence reaction in vitro in which FMNH2 is mixed with a solution of enzyme, aldehyde and O2 is shown in Figure 1. The fundamental features of the reaction sequence were first described by Hastings and Gibson in 1963.13 The enzyme binds reduced flavin mononucleotide to form a noncovalent enzyme:reduced flavin complex, intermediate I. Intermediate I reacts with O2 to form the 4a,5dihydro-4a-hydroperoxyflavin, intermediate II. The aldehyde binds to intermediate II and reacts to form the final products through a series of chemical steps which are presently ill defined. A network of binding equilibria involving the enzyme, FMNH2, O2, and aldehyde is presented in Figure 2;14'15 this scheme will serve as the basis for organization of the discussion below of recent advances in our understanding of the luciferase-catalyzed reaction. The chemical mechanism of the reaction remains the subject of much debate, which presumably will continue until more data regarding the precise chemical events preceding formation of the excited state of the flavin are available. Many methods are employed to assay the luciferase,12 and it appears that some of the differences in specific activity reported by various laboratories might be the result of different

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FIGURE 2. Network of binding equilibria involved in the bioluminescence reaction.14-15 Luciferase can bind FMNH2 to form intermediate I, the enzyme • FMNH2 complex, or aldehyde to form the deadend complex, enzyme-RCHO. The intermediate I appears to undergo a conformational change to yield E' • FMNH 2 which reacts with molecular oxygen to form the 4a,5dihydro-4a-hydroperoxyflavin, intermediate II. If the intermediate I binds aldehyde prior to reaction with oxygen, another complex, E • FMNH2 • RCHO, is formed, which reacts very slowly with oxygen. In the absence of aldehyde, intermediate II decays to yield FMN and H2O2,6 and the enzyme in an altered conformational state (see Section III.C.3). Binding of aldehyde to intermediate II yields the ternary complex, E • FMNHOOH • RCHO, which forms the tetrahedral intermediate shown in Figures 10 and 11. (Adapted from Reference 15.)

assay methods. The basic assay involves rapid mixing (manually, with the aid of a syringe) of enzyme, FMNH2, aldehyde, and O2. In one format, FMNH2, either catalytically reduced (using H2 with Pt catalyst) or photoreduced, is rapidly injected into a vial containing enzyme, aldehyde, and O2. In another, a solution of enzyme and FMN is reduced and rendered anaerobic by addition of dithionite, and the bioluminescence reaction initiated by injection of an air-equilibrated sonicated suspension of aldehyde. In a third assay format, the reduced flavin is supplied by the reduced pyridine nucleotide dependent flavin oxidoreductase to a solution of luciferase, O2, and aldehyde. The assay involves a single turnover of the luciferase in the first two methods, but the third method permits continuous turnover of the enzyme. For the enzyme from Vibrio harveyi, the first method, which involves incubation of the enzyme with the aldehyde substrate prior to injection of FMNH2, results in a reduced light yield relative to the second method, in which the aldehyde (with dissolved O2) is injected into a solution of enzyme and reduced flavin.16'17 This appears to be the result of aldehyde substrate inhibition of the enzyme, which is more apparent with the enzyme from V. harveyi than with the luciferases from the other species of luminous bacteria.17'19 Regardless of the mixing method, in the single turnover assay, the light intensity rises to a maximum and then decays exponentially (Figure 1). The peak intensity, which is proportional to the amount of luciferase, is reached within about 200 ms. The rate of decay of the light is dependent upon the species source of the enzyme and the alkyl chain length of the aldehyde.18 The light intensity in the coupled reductase assay is a function of the species source of the luciferase, the alkyl chain length of the aldehyde, and the concentrations of the reductase and the substrates. The coupled assay system can be established to supply a glow rather than a flash of light. The reaction catalyzed by luciferase will be discussed in more detail in Section II.

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FIGURE 3. Physical and restriction map of the V. harveyi lux AS genes. The first clone of lux DNA that was isolated was the fragment from the EcoRI site in the luxD gene to the EcoRI site in /laB.1 A 4 kb Hindlll fragment extending from the Hindlll site in luxD to a Hindlll site downstream of luxE cloned into pBR322 conferred aldehyde stimulatable bioluminescence upon E. coli carrying the recombinant plasmid. 2 ' 4 ' 21

B. CLONING AND SEQUENCING OF THE lux GENES The first clone of the genes for bacterial luciferase was obtained as the result of a collaborative effort between our group and that of John Abelson and his colleagues at the Agouron Institute in LaJolla. In a study of the limited proteolysis of luciferase, S. K. Rausch determined an amino acid sequence, Met-Asp-Cys-Trp-Tyr-Asp,20 within the a subunit whose utility he recognized for construction of a mixed-sequence oligonucleotide probe with minimal ambiguity. The 17-base oligomer, comprising only 8 unique sequences, was synthesized and used to probe a V. harveyi genomic clone bank in bacteriophage X; the 1.8 kb EcoRI fragment (see Figure 3) detected by the probe was isolated and shown to encode the luciferase a subunit (luxA gene) and the amino-terminal portion of the 3 subunit (luxB gene).1 The same mixed-sequence probe hybridized to a 4 kb Hindlll fragment, and we found that several clones of a genomic Hindlll digest of V. harveyi in pBR322 (12 of approximately 6000 colonies) were capable of light emission in E. coli when exposed to ndecanal vapor.2-4 (This result was surprising in view of the failure of earlier attempts to obtain glowing E. coli and the previous reports that the levels of flavin oxidoreductase in E. coli were too low to support high levels of bioluminescence. The finding that E. coli can support high level bioluminescence demonstrates that either the reduction of the flavin is not accomplished in vivo by the flavin oxidoreductase or the turnover of the E. coli enzyme and the redox potential within £. coli are capable of supplying adequate levels of reduced FMN for the luciferase to function.) Indeed, this experiment, in which transcription of the /wjcAB genes was from a promoter which has become known as the "anti-Tet" promoter, was the first demonstration of the utility of the lux genes to monitor gene expression in other organisms.21 This application of the lux genes is now widely recognized and exploited in numerous laboratories worldwide. Prior to the cloning of the luxA and luxR genes, we had been working to determine the amino acid sequences of both subunits of the enzyme. The partial amino acid sequences were very useful in obtaining what appear to be error-free nucleotide sequences for the luxA and luxB genes, and amino acid sequences of the a and 3 subunits22'23 (see Figures 4 and 5). The cloning and sequence determination of the luxA and luxB genes from two strains of V. fischeri25'2* Photobacterium leiognathi25*29'32 and Xenorhabdus luminescens26^ as well as V. harveyi, have now been completed. These sequences will be discussed and compared below (Section III.B). C. CONTROL OF lux GENE EXPRESSION A decade ago, the lack of a genetic system in luminous bacteria seriously impeded progress in understanding the molecular mechanisms that regulate the intensity of bioluminescence from these cells. Since the cloning of the lux gene systems, progress in the area of regulation of expression has been rapid. The regulatory systems for V. fischeri bioluminescence appear to function with fidelity in Escherichia coli, so that much of our current understanding comes from studies in that well-developed genetic background. The autoinduction system was originally described by Kempner and Hanson,34 who mistakenly ascribed

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 4. Nucleotide sequence of the /taA and lux& genes and the encoded amino acid sequences of the a and p subunits of V, harveyi luciferase. The sequence is shown for the region outlined in Figure 3. The nucleotide sequence and the encoded amino acid sequence demonstrate that the four open reading frames in this region are transcribed from left to right.22-23 Downstream of luxB, a region of hyphenated dyad symmetry is shown underlined. This region, noted by Johnston et al. >23 was shown by Sugihara and Baldwin24 to be necessary for high level expression of luciferase, apparently because of mRNA stability. Similar sequences in the same location have been noted in V. fischeri25 and in X. luminescent,™ but appear to be absent from P. leiognathi and P. phosphoreum (see text).

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FIGURE 4

(continued).

473

474

Chemistry and Biochemistry of Flavoenzymes

FIGURE 4 (continued).

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FIGURE 4 (continued).

the lag in production of luminescence to an inhibitor in the growth medium. The elucidation of the molecular basis of autoinduction was begun by Nealson et al.,35 who showed that the increase in luminescence in midlog phase cultures is due to production of a compound which they called the autoinducer. The basis of the catabolite repression of bioluminescence36 is now becoming better understood at the molecular level.37 40 The physical map of the genes encoded in the lux regulon of Vibrio fischeri is shown in Figure 6. References for the information provided in the following brief summary are in Section V. The regulon consists of two divergently transcribed operons, operonR (rightward operon) and operonL (leftward operon). The twcR (the only known gene of operonj and luxl genes together comprise the basic functions required for autoinduction. The Luxl protein is required for synthesis of the autoinducer, a diffusible molecule which passes the cell membrane to equilibrate with the environment. When the cells are free-living in the ocean, the autoinducer concentration remains very low. The LuxR protein is a positive regulatory protein which, together with the autoinducer, stimulates transcription of operonR. Transcription of operonL, and therefore the production of LuxR protein, is under control of the cyclic AMP receptor protein. If the cells are in a rich environment and grow to high cell densities, the local concentration of autoinducer will increase. If LuxR protein is available, a complex can form with the autoinducer, which binds to a specific palindromic sequence immediately upstream of the promoter of operonR to stimulate transcription of that operon. Since the first gene of operonR is luxl, enhanced transcription of operonR will lead to enhanced levels of Luxl protein and therefore, of autoinducer. The resulting positive feedback loop causes a strong response of the system to small initial levels of autoinducer. The discussion of genetic regulation, which will be reviewed in detail in Section V, is included here, not because of involvement of any flavoproteins, but because significant current interest in bacterial luciferase is due to potential applications of the system for genetic studies in other organisms. D. APPLICATIONS OF BIOLUMINESCENCE During the decade of the seventies, there was much interest in using luciferase as a nonradioactive alternative to radioimmunoassay procedures that are commonly performed in clinical laboratories. However, even though luciferase is stable and the assay is sensitive, the enzyme linked immunosorbant assay (ELISA) procedures based on other enzymes have become well established. One luciferase-based enzyme linked immunoassay, which is distinct from other ELISA assays, deserves some comment. The luciferase from Vibrio harveyi has a highly reactive thiol at position 106 of the a subunit.22-41 Stoichiometric reaction of this thiol with a variety of alkyl alkanethiolsulfonates renders the enzyme inactive, but the activity is readily recoverable by reduction of the mixed disulfide that results from the initial reaction.42-43 Derivatives of a variety of compounds, such as estradiol, having a methyl

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 5. Amino acid sequences and predicted secondary structures of the a and p subunits of luciferase from V. harveyi. Amino acid residues within predicted a-helical regions are indicated as such, residues within predicted (3 sheet regions are indicated within broad arrow-like forms, and random coil regions are indicated within parallel lines. Amino acid residues found more often on the surfaces of proteins are indicated by white backgrounds, and amino acid residues found more often within globular proteins are indicated by black backgrounds. Secondary structure prediction was by the method of Chou and Fasman.27 (Figures adapted from References 22 and 23.)

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FIGURE 6. Physical map of the lux regulon of Vibrio fischeri. The locations of the "leftward" and "rightward" operons of the regulon are shown at the bottom. The 7 genes, luxR and /«*ICDABE, and the direction of transcription of each are indicated. The nucleotide sequence of these 7 genes, which are necessary and sufficient to confer regulated bioluminescence on E. coli carrying the recombinant genes,5 has been reported.25 Regulatory functions, /wjcR and luxl, reside in both operons, giving rise to strong interaction between the two (see Section V). The structural genes required for bioluminescence, /wixCDABE, encode the fatty acid reductase complex (LuxC, LuxD, and LuxE) and the subunits of luciferase, a (LuxA) and p (LuxB). The SDS molecular weight of each of the gene products is included in the figure, as are many of the restriction sites. Downstream of luxE is another extended open reading frame that was described by Baldwin et al.25 The sequence of the downstream open reading frame has recently been completed and designated luxG (see text, Section IV.D), but no protein product or direct involvement in bioluminescence has been demonstrated.

thiolsulfonate ligand were synthesized and shown to stoichiometrically inactivate the luciferase.43'44 The luciferase-ligand complex could then be used in a conventional immunoassay format, and the amount of luciferase-ligand-antibody complex formed determined by reduction of the mixed disulfide, which would release the luciferase into solution for assay by standard methods. Numerous other methods have been developed that incorporate immobilized enzyme technology into a coupled bioluminescence assay.45"49 In these methods, a specific metabolite is detected through an enzyme-catalyzed reaction that yields, ultimately, reduced pyridine nucleotide, which is used by one of the pyridine nucleotide-dependent FMN oxidoreductases to supply FMNH2 for luciferase.50 Many versions of these assays have been described, with some of the more recent advances being in the format of the immobilized enzymes.45"47 Thin film technology and fiber optic systems have been incorporated that allow exquisite sensitivity and selectivity.48'49 It has recently been reported that the enzyme from Xenorhabdus luminescens has a very high thermal stability, and thus may be the enzyme of choice for applications of bioluminescence.51 Today there is much interest in the luciferase genes for diagnostic systems in vivo.

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Chemistry and Biochemistry of Flavoenzymes

Availability of the cloned genes has allowed studies employing light emission to monitor gene activity in a wide spectrum of organisms. For example, in the cloning of the luxAE genes from V. harveyi, the "antiTet" promoter of the plasmid pBR322 was clearly demonstrated.4 Several laboratories5256 have reported use of the livcAE gene cassette to monitor regulated gene expression in procaryotes. By expressing luminescence in pathogenic bacteria, it has been possible to monitor the movement of pathogens through tissues.57 Transposons carrying the lux genes have been developed which are capable of random insertion into the chromosomes of specific organisms.58 Insertion into a regulated gene results in substitution of bioluminescence for the original function of the interrupted gene. Schauer and Losick and their colleagues have made elegant use of this system to study developmental control of gene expression in the filamentous bacterium Streptomyces coeiicolor.59'61 Another exciting recent development is the discovery62-63 that the cloned tuxA and luxB genes from V. harveyi can be fused into a single open reading frame by deletion of the translational stop codon from luxA and altering the intergenic sequence such that the two reading frames are in phase. The resulting construction encodes a single polypeptide chain comprising the original a and (3 subunit amino acid sequences with a short linker polypeptide connecting the two; the single polypeptide is active in the bioluminescence reaction. This finding is of fundamental importance for studies of the structure and folding of the luciferase enzyme, and is of practical importance as a vehicle for expression of bioluminescence in eucaryotic cells. It is probable that in the near future, there will be a large number of reports of the use of the fused livcAB construction to monitor gene expression in eucaryotic cells.

II. ENZYMOLOGY OF BACTERIAL LUCIFERASE The literature describing the enzymology of bacterial luciferase is difficult for nonspecialists (and perhaps also for devotees!) due to conflicting reports concerning such fundamental features as reaction stoichiometry and the involvement of thiols on the enzyme. In recent years, a consensus has been developing within the field regarding such basic features of the system. Without discussing in detail the demise of conflicting hypotheses, we will endeavor here to present the current ideas that are being investigated by workers in the field. A. SUBSTRATES, PRODUCTS, AND REACTION STOICHIOMETRY The reaction catalyzed by bacterial luciferase is presented below.6 The enzyme binds a single molecule of reduced flavin mononucleotide, forming intermediate I, which in FMNH2 + O2 + RCHO -> FMN + RCOOH + H2O + hv turn reacts with molecular oxygen to form an activated hydroperoxy flavin species, intermediate II64-66 (see Figure 7). The enzyme-bound C4a-hydroperoxyflavin intermediate then binds a long-chain aldehyde. The chemical products of the luminescent reaction are FMN, the long-chain carboxylic acid, and presumably water. In the reaction, the molecular oxygen is cleaved, with one atom of oxygen being delivered to the aldehyde substrate to form the acid product;67 the second atom of oxygen is presumably reduced to water. In the older literature, there were suggestions of the mechanistic involvement of two flavins per turnover of the enzyme, but we believe that the preponderance of the existing data are not consistent with such proposals. Meighen and Hastings16 developed a kinetic assay to measure the number of binding sites on luciferase for reduced flavin, and found a stoichiometry of 1:1; subsequently, equilibrium binding of FMNH2 to luciferase was measured by Becvar and Hastings68 using circular dicnroism spectroscopy, again demonstrating a single binding site for FMNH2 per luciferase a($ dimer. Perhaps the most crucial experiment in this regard was reported in the same publication of Becvar and Hastings,68 who measured

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FIGURE 7. Formation of intermediate II from FMNH2 and O 2 . The identity of intermediate II was postulated by Hastings et al.64 on the basis of spectroscopic analysis, and the structure was demonstrated by Vervoort et al.66 based on I3CNMR.

the quantum yield of the reaction relative to the limiting reactant, enzyme or flavin, under conditions in which the ratio of enzyme to flavin was varied over a range of approximately 106. Over this range, the quantum yield relative to flavin under conditions of enzyme excess was the same as the quantum yield relative to enzyme under conditions of flavin excess. That is, the quantum yield relative to flavin when the ratio of enzyme to flavin was 1000 to 1 was the same as the quantum yield relative to enzyme when the ratio of enzyme to flavin was 1 to 1000. This experiment demonstrated clearly that in the light emitting reaction, flavin and enzyme are involved with a stoichiometry of 1 to 1. It is worth noting, however, that there is some indication from NMR studies69 that more than one flavin can bind to luciferase, though the second molecule is bound very weakly, perhaps "nonspecifically", and also that in a kinetic study, Matheson and Lee70 observed a greater than first order dependence of bioluminescence intensity on FMNH2 concentration at high luciferase concentrations (200 |xAf). The stoichiometry of the reaction with aldehyde in the light-emitting reaction is also 1:1.19 The behavior of the enzyme in response to aldehyde is, however, more complex. It has been shown that the luciferase from V. harveyi is sensitive to aldehyde substrate inhibition. 1719 This reversible inhibition is seen when the FMNH2 injection assay is used with elevated concentrations of aldehyde; inhibition is not observed in the dithionite assay, in which the enzyme is first incubated with FMNH2 under anaerobic conditions (in the presence of dithionite) and then the reaction is initiated by the injection of a solution containing aldehyde and oxygen. The data could be fit to a model in which the inhibition resulted from binding of two molecules of aldehyde, one with a higher affinity that was chemically competent and the second with a lower binding affinity that resulted in inhibition of the enzyme.19 An alternative explanation for the observed aldehyde substrate inhibition, implicit in the reaction scheme presented by Raushel and Baldwin14 and Raushel et al.15 and discussed below, is that the binding of aldehyde to free enzyme blocks binding of FMNH2, such that aldehyde must dissociate prior to FMNH2 binding. It should be stressed here that the components of the reaction, luciferase, FMNH2, O2, and aldehyde, are necessary and sufficient for the bioluminescence reaction. While the involvement of accessory emitter proteins in the light-emitting reaction catalyzed by bacterial luciferases has been clearly demonstrated by numerous experiments, primarily from the laboratories of John Lee and his colleagues (see References 9 and 10 for recent reviews), these accessory proteins are not essential for the reaction catalyzed by the luciferase, and appear to function as secondary emitters without being required for the chemistry of the reaction. B. REACTION SCHEME The reaction scheme first proposed in 1963 by Hastings and Gibson13 remains today the

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Chemistry and Biochemistry of Flavoenzymes

central core of what is a rather complex reaction. Reports from other investigators during the intervening years have resulted in modification of specific details, but the basic reaction scheme remains essentially as it was originally proposed. Many investigations of the kinetic mechanism of the luciferase-catalyzed reaction have relied on measurements of light emission rather than on spectroscopic or other methods of detection of intermediates or products. As can be seen from the scheme presented in Figure 2, 14 - 15 there are numerous steps between the mixing of substrates and the emission of light, so that from measurement of light emission alone, little can be learned about the precise details of the kinetic mechanism. To dissect the kinetic mechanism of bacterial luciferase, detailed measurements of the rate of each individual step must be made. Such measurements have been reported (see, e.g., earlier studies reviewed in Reference 6, and more recent studies in References 15, 70, and 71), but a complete set of rate constants determined under comparable conditions is not yet available. The following discussion reflects the current understanding of the kinetic mechanism as it appears in the literature. 1. Binding of Reduced Flavin Mononucleotide Earlier studies of binding of reduced flavin and also of oxidized flavin to luciferase have been reviewed previously6-10 and will not be discussed extensively here. It has been known since the early 1970s that the binding affinity of luciferase for reduced flavin (Kd about 1 (jiM16) is about 100 times its affinity for oxidized flavin (Kd > 0.1 mA/72). In fact, a recent report from Paquatte and Tu73 points out, with data on binding affinities for luciferase with a series of flavin analogues in their oxidized and reduced forms, that while luciferase is indeed quite fastidious in its structural requirements for tight binding of reduced flavins,74 it is far more tolerant of variations in structure of the flavin in its oxidized form. In the 1970s, Meighen and MacKenzie74 reported an extensive study on the structural requirements of the flavin side chain for good reduced flavin binding and high quantum yield with the V. harveyi enzyme. The luciferase appears to be quite specific for the reduced riboflavin-5'-phosphate; the anionic group seems to function not only in binding to the enzyme but also (probably indirectly) in enhancing the quantum yield of the reaction. Meighen and MacKenzie74 found that the N(3)H group of the flavin isoalloxazine moiety is essential for the reaction (alkylation at this position resulted in near-abolition of activity), and that a negatively charged group on the flavin side chain at least 8.4 A from position N(10) is required both for tight binding and for a high quantum yield. Chen and Baldwin,75 using 8-substituted reduced flavin analogues with the V. harveyi enzyme, showed that 8substitution had little effect on the binding of reduced flavins, suggesting that the 8 position of the FMNH2 is oriented away from the protein (exposed to solvent) in the complex. Vervoort and co-workers have recently reported a thorough investigation by NMR spectroscopy of the binding of oxidized and reduced FMN69 and also of intermediate II.66 The NMR data suggest that the bound reduced flavin is in the form of the N(l) anion (FMNH~). 69 Ziegler-Nicoli et al.41 studied the pH dependence of FMNH2 binding and reported the existence of two ionizable groups (flavin or protein), one with an apparent pKa of about 6.8 which must be protonated for good flavin binding, and one with an apparent pKa of about 6.2 which must be deprotonated for tight binding. Since the protein stabilizes the N(l) anion of the flavin,69 the N(l) pKa when bound to luciferase might correspond to the pKa of the more acidic group reported by Ziegler-Nicoli et al.41 The NMR studies69 indicated that the N(5) atom of bound reduced flavin possesses a high degree of sp2 hybridization, suggesting that the isoalloxazine moiety is highly coplanar around the N(5) atom, whereas the N(10) atom has a somewhat decreased sp2 character and thus might be somewhat out of the molecular plane. The phosphate group of bound flavin, both oxidized and reduced, appears to be in the dianionic state.69 FMNH2 binds to luciferase under equilibrium conditions to form a 1:1 complex.16-68 The

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FIGURE 8. Competition between the enzymatic and nonenzymatic reactions of FMNH2 with O2. In the dithionite assay described by Meighen and Hastings,16 an equilibrium mixture of luciferase and FMNH2 is established under anaerobic conditions (in the presence of excess dithionite). The bioluminescence reaction is initiated by rapid injection of O2-equilibrated aldehyde suspension. The assay is based on the assumption that the initial maximum light emission observed is proportional to the amount of E • FMNH2 present at the time of oxygen injection (see Figure 2 and text).

stoichiometry of the interaction has been determined by numerous methods, but the equilibrium dissociation constant has been measured only by the kinetic method described by Meighen and Hastings.16 The method is based on several assumptions of kinetic rate constants that have not been rigorously tested, particularly with the various strains and mutants from which luciferases have been purified. In the Meighen and Hastings method, the enzyme is incubated with a particular concentration of FMNH2 prepared by reduction with dithionite, and the reaction is initiated by injection of O2 and aldehyde. It is assumed that the enzyme-FMNH2 complex is quantitatively converted to intermediate II by a reaction which is fast compared with the rate of dissociation of the enzyme*FMNH2 complex. It is further assumed that any free FMNH2 present at the time of injection of O2 would be quantitatively removed from the reaction by the nonenzymatic autooxidation pathway.76 The yield of light would then be proportional to the concentration of E*FMNH2 complex present at the time of injection of oxygen. This series of reactions is depicted in Figure 8. The first assumption appears to be reasonable. Recent measurements of the reaction of the enzyme-FMNH2 complex with oxygen (k3) give a pseudo-first order rate constant of about 350 s'1 at 25°C with ambient oxygen (120 (xM),15 while the value of k4 (which is a complex reaction) would be about 10 s"1.76 The second order rate of formation of the enzyme*FMNH2 complex is not known, but it is reasonable to assume that it is fast, approaching the diffusion limit of perhaps 107 M~l • s ~ l . It would appear reasonable then that rapid mixing of oxygen with a preexisting equilibrium mixture of enzyme and reduced flavin would lead to a rapid and quantitative conversion of the enzyme-FMNH2 complex to intermediate II. However, with FMNH2 at 1 to 10 \LM, it would be expected that not all of the free FMNH2 would be removed by nonenzymatic reaction with O2, but that some additional enzyme-FMNH2 complex could form as a result of the shifted equilibrium condition. This second assumption of the dithionite assay (no formation of additional E-FMNH2 after O2 injection) probably does not lead to large errors with the wild-type enzyme, but with the isolation and construction of numerous mutant forms of luciferase having greatly altered kinetic parameters, and with the use of flavin analogues, the validity of the assumptions underlying the kinetic determination of Kd for FMNH2 must be reevaluated. 2. Binding of Aldehyde The organic substrate of the luciferase in vivo appears to be the 14-carbon aliphatic aldehyde, myristic aldehyde,77 while in vitro, many aliphatic aldehydes have been employed.6 For standard laboratory work, n-decanal is convenient because of its lower melting point

482

Chemistry and Biochemistry of Flavoenzymes TABLE 1 Binding of Aliphatic Aldehydes to Luciferase

a

b

c

Aldehyde

K Ma

Kd"

K/

M-octanal n-decanal n-dodecanal

5.0 (xM

47 JJLM 21 jxA/ 11 ^M

43 (AM 28 jxM 18 \LM

1.1 (JLM

0.2 V.M

The values presented are the concentrations required for half-maximal bioluminescence activity for the luciferase from Vibrio harveyi in 20 n\M Bis-Tris buffer, pH 7.1, at room temperature (22 to 25°C) as described by Holzman and Baldwin. 19 The apparent dissociation constants were calculated from the data of Ziegler-Nicoli, Meighen and Hastings41 who demonstrated that aliphatic aldehydes protect the reactive thiol of the luciferase from reaction with Nethylmaleimide. In calculating the apparent equilibrium dissociation constants, the assumption was made that the reactive thiol is not reactive when the enzyme is complexed with the aldehyde. Reaction conditions were 20 mM phosphate, pH 7.0, 25°C. The apparent dissociation constants were calculated from the data of Welches and Baldwin42 who showed that aliphatic aldehydes protect the enzyme from inactivation by 2,4-dinitrofluorobenzene. Reaction conditions were 50 mAf phosphate, pH 7.0, 25°C.

and greater solubility. The question of aldehyde binding to luciferase and its possible significance in the kinetic mechanism has not been carefully discussed in the literature. The first demonstration of binding of aldehyde to the free luciferase was published by ZieglerNicoli et al., 41 who demonstrated that long-chain length aldehydes protect the reactive thiol in the active center of the luciferase from reaction with A^-ethylmaleimide. They used the aldehydes n-octanal, n-decanal, and n-dodecanal at 50 jxM, the concentration used in the normal flavin injection assay, and observed significant but not quantitative protection. Using their data and assuming that /V-ethylmaleimide does not react with the enzyme:aldehyde complex, approximate equilibrium dissociation constants can be calculated for the aldehydes bound to the free enzyme. These values are given in Table 1. As can be seen from the table, the calculated equilibrium dissociation constants are too high to explain saturation by aldehyde in the flavin injection assay, suggesting that either the thiol is not completely protected by bound aldehyde or that the affinity of binding of aldehyde to luciferase is enhanced by binding of reduced flavin mononucleotide or formation of the flavin hydroperoxide, intermediate II. The observation that aldehyde binding to free enzyme can protect functional groups in the active center of the enzyme from modification has been repeated several times, but the significance of this observation was not discussed until Holzman and Baldwin investigated the mechanism of aldehyde substrate inhibition.19 The inhibitory effects of high concentrations of aldehyde had been reported in previous studies17'18-74'78 and clearly demonstrated both for the V. harveyi enzyme and for a mixed subunit hybrid luciferase consisting of the a subunit from V. harveyi luciferase and the p subunit from P. phosphoreum luciferase.17 In the report from Holzman and Baldwin,19 the data obtained showing inhibition of the luciferase by high aldehyde were fit to a mathematical model involving formally the binding of two molecules of aldehyde, one associated with the active center and the second associated with the inhibition of the enzyme. Alternatively, the data could have been fit to a more complex model in which binding of aldehyde to the enzyme precluded binding of the flavin substrate, but there were at that time no data to allow distinction between the two models. However, recent stopped-flow measurements of the effects of aldehyde on the formation of intermediate II suggest that O2 cannot react (or reacts with a greatly reduced rate) with the flavin in the ternary complex of enzyme*FMNH2-aldehyde and suggest that the binding of flavin to the binary complex of enzyme-aldehyde does not occur.15 We therefore wish to withdraw the previous model involving binding of two molecules of aldehyde in favor of the alternative model presented in References 14 and 15, and discussed in this review.

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FIGURE 9. Catalytically significant substrate binding pathway. While substrates can bind to the enzyme in random order (see Figure 2), nonproductive complexes form which are apparent from the aldehyde substrate inhibition described by Holzman and Baldwin.19

3. Kinetic Mechanism of the Reaction In the current model (Figure 2), Raushel et al.14'15 propose that binding of aldehyde to free enzyme or to enzyme-FMNH2 results in formation of deadend complexes from which aldehyde must dissociate before the enzyme may proceed successfully through the reaction. This kinetic model would explain the aldehyde substrate inhibition as well as the two binding site model advanced by Holzman and Baldwin;19 furthermore, it accommodates the apparently lower binding affinity of the free enzyme for the aldehyde that is observed in the bioluminescence reaction (Table 1). It should be stressed here that the binding of substrates (FMNH2 and aldehyde) to the luciferase clearly can occur in random order, but the catalytically significant binding appears to be ordered, as shown in Figure 9. This kinetic mechanism explains why the activities and kinetics reported can vary as a consequence of the method that is used to perform the assays. C. STABLE AND TRANSIENT INTERMEDIATES The stable complexes formed during the reaction of luciferase with its substrates to yield bioluminescence include the enzyme*FMNH2 stoichiometric complex (intermediate I), the product of reaction of intermediate I with O2 [the C(4a) hydroperoxydihydroflavin, intermediate II (Figure 7)], and the ternary complex of intermediate II with aldehyde, intermediate HA. Intermediate I has been demonstrated under equilibrium conditions using fluorescence spectroscopy79 and circular dichroism spectroscopy.68 The stoichiometry of the complex was shown to be 1:1 by both methods. Intermediate II was first isolated and characterized by Hastings and colleagues,64 who suggested, based on spectral comparisons with model compounds, that it was a 4a,5-dihydro4a-hydroperoxyflavin. Definitive proof of its chemical structure was provided by Vervoort et al.66 who used 13C NMR spectroscopy with intermediate formed by reaction of luciferasebound FMNH2 with O2. The intermediate was isolated by low temperature (0°C) chromatography in buffers containing n-dodecanol, which stabilizes the intermediate.80 The NMR results suggested that the 4a-hydroperoxyflavin bound to luciferase is in an almost planar configuration.66 Intermediate IIA has not been questioned in the literature, but its existence has not been rigorously demonstrated. Hastings and Balny81 demonstrated a slight shift in the absorbance spectrum of intermediate II in the presence of aldehyde, but no other equilibrium spectral measurements have been reported. Holzman and Baldwin,19 using a kinetic method, demonstrated that the aldehyde:luciferase stoichiometry in the light emitting reaction is 1:1. Several investigators have shown that the formation of intermediate IIA is reversible.78'82 In the normal flavin injection assay, secondary injection (following the injection of reduced flavin by several seconds) of aldehyde of a different chain length will result in a shift of

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 10. Eberhard and Hastings mechanism. In 1972, Eberhard and Hastings83 postulated the formation of a 4a,5-dihydro-4a-peroxyflavin, intermediate II, from reaction of luciferasebound FMNH 2 with O 2 . Intermediate II reacted with the aldehyde to form a tetrahedral intermediate which decayed by a Baeyer-Villiger type shift to yield the acid, hydroxide, and excited FMN. Their proposal predicted several important aspects of the bioluminescence reaction, which have been demonstrated in the intervening years (see Section II. D), even though the BaeyerVilliger rearrangement is not supported by current data.84

the decay of bioluminescence to a rate determined by the relative concentrations and respective decay rates of the two aldehydes present after the second injection. This observation demonstrates that formation of intermediate IIA is reversible, but it does not offer any information about the steps or potential intermediates involved in formation of the intermediate that we refer to collectively as intermediate IIA. That is, the initial binding of aldehyde is certainly a noncovalent process, but formation of the covalent bond which yields the proposed tetrahedral intermediate (see Figures 10 and 11) could likewise be reversible. Until we know the rates of the various steps involved in reaction of aldehyde with the intermediate II, we will continue to describe the reaction in macroscopic terms. With regard to the complexity of the kinetic mechanism of luciferase depicted in Figures 2 and 9, it should be stressed that it is dangerous to draw conclusions regarding the mechanistic effects of mutations on the luciferase-catalyzed reaction without performing a thorough investigation of the kinetics of the reactions catalyzed. A recent example of these dangers comes from the work of Xi et al.85 with a mutant enzyme, aCys-106 —> Val, that had previously been reported by Baldwin et al.86-87 Xi et al.85 attributed the very rapid appearance of the product oxidized FMN (by monitoring absorbance at 450 nm) in the reaction catalyzed by the aCys-106 —> Val luciferase to a fundamental change in reaction mechanism for the pathway of flavin oxidation, from that of a flavin monooxygenase (via the C(4a)-peroxydihydroflavin, intermediate II) to a flavin-dependent oxidase activity. However, Raushel et al. 15 - 87 determined the rate constants for the reaction of oxygen with the enzyme-bound FMNH2 to form intermediate II and for the breakdown of intermediate II to form FMN for both the wild-type and the mutant enzymes. By direct observation of intermediate II, they found that the rate and extent of formation of intermediate II for the mutant were essentially the same as with the wild type, ruling out any fundamental change in chemical mechanism of flavin-oxygen reaction. The rapid appearance of the product FMN in the reaction with the valine mutant enzyme is due to the instability of the enzyme-bound peroxyflavin, whose rate of breakdown is about 150 times faster than that of the wild type,87 but not to failure to form the intermediate. In the 1980s, a series of papers from the laboratories of Lee,70'88-90-91 and of Ghisla and Hastings71-84-92 reported the kinetic and spectroscopic properties of a fluorescent transient formed in the luciferase reaction. The possibility that this transient represents the luciferasebound flavin emitter [perhaps the C(4a) hydroxyflavin?] has been discussed in the primary references and also in a recent review,10 and will not be discussed further here.

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FIGURE 11. Formation of the proposed dioxirane. Raushel and Baldwin14 have proposed an alternative to the Baeyer-Villiger rearrangement, also shown here for comparison. The Raushel-Baldwin proposal involves decomposition of the tetrahedral intermediate to form an oxygen-oxygen-carbon three-membered ring, a dioxirane, and the C(4a) hydroxyflavin. Decay of the dioxirane (Figures 12 and 13) could lead to formation of the emitting species in the bioluminescence reaction (see text).

D. POSTULATED REACTION MECHANISMS A central concern for any chemiluminescence mechanism is formation of the primary excited state. Eberhard and Hastings83 proposed the first chemical mechanism for the bacterial bioluminescence reaction (Figure 10). In their proposed mechanism, the tetrahedral intermediate IIA broke down by a Baeyer-Villiger reaction to yield the carboxylic acid and the luciferase-bound flavin in the first singlet excited state. This mechanism has stood for many years as the best available to explain the many observations regarding this system. Many other mechanisms have been proposed,6-84 but can be discounted due to lack of consideration of critical aspects of the bioluminescence reaction. The Eberhard and Hastings mechanism83 anticipated several features of the bioluminescence reaction which have since been demonstrated by other workers. First, the mechanism as drawn shows the reaction of oxygen with the reduced flavin to yield the C(4a) peroxydihydroflavin. This site of reaction was consistent with model chemistry of the day, but the definitive demonstration of the site of oxygen addition awaited the NMR studies of Vervoort et al.66 Second, the Eberhard and Hastings mechanism required that the molecular oxygen be split, one atom being incorporated into the carboxylic acid and the other being incorporated into water. By utilizing 18O2, Suzuki et al.67 demonstrated incorporation of one atom of the oxygen into the carboxylic acid product. However, there is one critical aspect of bacterial bioluminescence which the mechanism

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FIGURE 12. Decay of dioxirane to form the excited carboxylic acid. Homolytic cleavage of the oxygen-oxygen bond of the dioxirane would yield the diradical, which should rearrange to yield the excited carboxylic acid in either the singlet or triplet state. In the presence of the lumazine protein or the yellow fluorescent protein (see Sections IV.B and IV.C), the primary excited state (carboxylic acid) could excite the chromophore on the secondary protein, while in the absence of the secondary protein, the C(4a) hydroxyflavin could receive the excitation of the excited carboxylic acid. 14

of Eberhard and Hastings83 does not accommodate (several other difficulties with the BaeyerVilliger pathway have been discussed in a recent review84). Lee and his co-workers have demonstrated that the reaction that is catalyzed by the luciferase enzyme from some species can result in formation of the excited state of the lumazine chromophore of the lumazine protein if the accessory protein is included in the reaction mixture (reviewed in Reference 10). The bioluminescence emission spectrum is thereby effectively blue-shifted relative to the emission spectrum from the luciferase alone. This observation raises serious questions concerning the nature of the primary excited state in the bioluminescence reaction. If the primary excited state is some form of luciferase-bound flavin, which emits at about 490 nm, it is difficult to explain how it could also populate the excited state of the lumazine protein, with a peak emission at about 478 nm.10 There appear to be two possible resolutions for this difficulty. The first would be to propose a change in mechanism resulting from the presence of the lumazine protein such that the primary excited state in the reaction would be formed on the lumazine protein. The second would be to propose that the primary excited state formed on the luciferase is not the excited state of the flavin, but of some other species of sufficient energy to populate (by transfer) the excited state of either the lumazine protein or the luciferase-bound flavin. It should be stressed here that these two possibilities are not mutually exclusive (see below). We pointed out in an earlier review6 the problem discussed here, and suggested that an excited carbonyl group would be an excellent candidate for the primary excited state. Matheson et al.88 have also proposed the production of an energy-rich primary excited species ("EX") which could excite either a luciferase-bound flavin species or the chromophore of an accessory protein. Raushel and Baldwin14 have recently proposed a mechanism which appears to accommodate all currently known aspects of bacterial bioluminescence and suggests an avenue to an excited carbonyl. Central to this mechanism is the formation of a dioxirane from the tetrahedral intermediate (Figure 11). The idea that a dioxirane might be a "high energy intermediate" has been proposed by Cho and Lee.89 The field of dioxirane chemistry is comparatively new and various potential pathways for breakdown of the dioxirane cannot at this time be eliminated based on model compound chemistry. Based on current understanding of these compounds, we can propose two potentially chemiluminescent pathways for the breakdown of the dioxirane, both sufficiently energetic to populate an excited state yielding a photon in the blue. Adam and his co-workers93 suggest that the dioxirane could undergo homolytic cleavage of the oxygen-oxygen bond to yield the diradical, which should rearrange to form the carboxylic acid in either the triplet or singlet state (Figure 12), a mechanism analogous to the mechanism of degradation of dioxetanes.94 Even the

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FIGURE 13. CIEEL mechanism for decay of dioxirane. An alternative to the decay of the dioxirane described in Figure 12 is the CIEEL pathway, in which the dioxirane receives an electron from a donor/fluorophore [C(4a) hydroxyflavin or the lumazine protein/yellow fluorescent protein] to yield a radical ion pair. Conversion of the dioxirane radical anion to the carboxyl radical and back transfer of an electron would yield the excited state of the fluorophore.14

triplet state should have sufficient energy for detection by means of enhanced chemiluminescence in the presence of a suitable fluorophore.93 The postulated C(4a) hydroxyflavin is such a fluorophore, which could become excited by interaction with either the triplet or the singlet product of the decay of the dioxirane. If the primary excited state in the reaction were the carboxylic acid, then the most likely emitter in the reaction catalyzed by pure luciferase would be the luciferase-bound flavin (Figure 12). Addition of either the lumazine protein or the yellow fluorescent protein to the reaction mixture would supply an alternative fluorophore. The second pathway for bioluminescent decay of the dioxirane involves a CIEEL mechanism. In the chemically initiated electron exchange luminescence (CIEEL) mechanism, postulated by Koo and Schuster95 for the process leading to chemiluminescence of dioxetanes, the dioxirane could receive an electron from a donor/fluorophore to form a radical ion pair. Rearrangement of the radical anion of the dioxirane would yield the radical anion of the carboxylic acid. Backtransfer of the electron to the donor would then yield the excited state of the fluorophore (Figure 13). In the case of luciferase, Raushel and Baldwin14 have proposed that in a CIEEL mechanism, the C(4a) hydroxyflavin could play the role of the electron donor/fluorophore. In the presence of the lumazine protein or the yellow fluorescent protein, direct protein:protein interaction between the luciferase and the secondary emitter protein could result in participation of the chromophore of the secondary emitter protein in the CIEEL process with the luciferase-bound dioxirane. This proposed mechanism incorporates a solution to the long-standing paradox regarding the apparent blue-shift in bioluminescence mediated through the lumazine protein. In fact, it was this enigma that led to the proposal6 10 years ago that the flavin might not be the primary excited state, but that there might be some other higher energy state that could excite either the flavin or the lumazine protein. Both the lumazine protein10 and the yellow fluorescent protein96-97 appear to interact directly with the luciferase and to cause an acceleration in the luciferase-catalyzed reaction. This observation does not permit discrimination between the CIEEL mechanism and the direct transfer mechanism, since in the former, the secondary emitter protein would supply an alternative donor and therefore would be expected to effect a kinetic change, and in the latter, the direct protein:protein interaction could well contribute to the observed kinetic alterations. Furthermore, both mechanisms involve a comparable shift of an H" atom, so distinguishing on the basis of isotope effects would be difficult.

III. STRUCTURAL STUDIES OF BACTERIAL LUCIFERASE Determination of the structure of bacterial luciferase by single crystal X-ray diffraction methods has been slow due in part to the instability of the crystals in the X-ray beam and in part to the size of the unit cell of the crystal. Diffraction quality crystals have been reported and substantial progress has been made, but a high resolution structure is not yet available.98 In the absence of a high resolution structure, the classical methods of chemical modification,

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limited proteolysis and mutant enzyme analyses have been employed to probe the structure. The discussion which follows will present our current state of understanding of the structure of bacterial luciferase. A. ARCHITECTURE OF THE ENZYME Bacterial luciferases from all species so far studied are heterodimeric, consisting of two subunits, a and p, with molecular weights of about 40,000 and 35,000, respectively. The luciferase from Vibrio harveyi, the enzyme which has been investigated most extensively, has a single active center that appears to reside primarily, if not exclusively, on the a subunit. It has been suggested that the single active center resides at the subunit interface, but the data supporting this hypothesis are not very strong, a point discussed below (Section III.C.6). The active center of the enzyme appears to be a hydrophobic cleft or pocket which accommodates the reduced flavin, oxygen and the aldehyde substrate. Several lines of evidence suggest that the enzyme undergoes a significant conformational rearrangement during the catalytic cycle. The amino acid residues surrounding the active center apparently undergo a major structural rearrangement which slowly relaxes to the original structure following the catalytic cycle if reduced flavin is no longer available. These points will be discussed in detail in the following sections. B. AMINO ACID SEQUENCES OF BACTERIAL LUCIFERASES The amino acid sequence of the luciferase from Vibrio harveyi was the first to be determined, and it was determined in its entirety by DNA sequencing methods.22'23 In addition, the amino acid sequence of about 2/3 of each subunit was determined by protein chemical methods, so there is little doubt that the sequence is correct and the encoded amino acid sequences are not further processed posttranslationally. Since the cloning and sequencing of the luxAE genes from V. harveyi, the nucleotide sequences of the luxAB genes of two strains of V. fischeri25'2* two strains of Photobacteriwn leiognathi25-31*32 and the nonmarine luminous bacterium Xenorhabdus Iuminescens2(> have been reported. An alignment of the amino acid sequences of these polypeptides is shown in Figure 14. The amino acid sequence of the luciferase subunits from only one strain of each species is included in the alignment. Inspection of these sequences reveals several very interesting features. First, it is immediately apparent that the a and (3 subunits are homologous, as suggested earlier from N-terminal amino acid sequence data.100'101 Second, within the a subunit, there are several regions that are strongly conserved, and several regions that are remarkable in the apparent lack of sequence conservation. Third, it is apparent that a major difference between the a and (3 subunits resides in the apparent deletion from p of a region of about 30 amino acid residues. This region is strongly conserved within the a subunit, and appears to comprise a portion of the active center of the enzyme (discussed below, Section III.C). Finally, it is of interest that the amino acid sequences of the luciferases are as different as they are. The differences would suggest that either the strains carrying the lux genes have been genetically isolated for a very long time or that the force of natural selection maintaining the constancy of the amino acid sequence of the luciferase protein is not as strong as for such proteins as the cytochromes c. The strategy used in establishing the alignment of the amino acid sequences shown in Figure 14 was to minimize the number of gaps (insertions or deletions) in the sequences and to maximize the number of conserved positions. The definition of a conserved residue was based on the considerations of Swanson,102 who has tabulated the frequencies of specific residues occurring in solvent contact and buried in globular proteins. A substitution was considered to be conservative if a particular amino acid residue was replaced by a residue having the same (or similar) positional propensity. This alignment is similar to others that have relied on a smaller data base."

FIGURE 14. Alignment of the amino acid sequences of the a and p subunits of the luciferases from V. harveyi22-2* (V.h.), V. fischeri25-2* (V.f.), P. leiognathi25-™*2 (P.I.), and X. luminescent2* (X.I.). The alignment is similar to that presented earlier for a less extensive sequence database," and to an alignment recently presented by Johnston et al.26 A minimum of gaps were introduced to maximize the identity between the sequences, and in the case of the apparent deletion in the p subunits, to accommodate the apparent conservation of sequence within the C-termini of the a and (3 subunits.

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FIGURE 14 (continued).

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1. Interaction Between the a and P Subunits From a comparison of the sequences of the 8 polypeptides, it is apparent that 46 positions, or 14% of the possible comparisons, are identical for all 8. These residues, which are invariant between the a and (3 subunits of the luciferases of all 4 species, reside in several extended regions: between residues 36 and 53, residues 80 and 104, and residues 174 and 194. In addition, there are several other smaller regions of strong sequence similarity between the a and 3 subunits. These include the amino terminal 4 residues, residues 113 to 117, residues 153 to 163, and residues 218 to 224, as well as several other isolated or very short regions of sequence similarity. If one were to include the conservative substitutions in the comparison of the set of a subunits with 3 subunits, about 80% of the residues in 3 are either identical to or of the same positional propensity as the homologous residue in the a subunit. These observations argue compellingly that the a and 3 subunits of luciferase are homologous (of a common evolutionary origin) and that the subunits have a comparable overall three-dimensional folding pattern, similar to other homologous proteins such as the globin family. There are few heterodimeric proteins with which to compare the luciferase subunits in establishing the significance of the regions of sequence similarity. However, in comparing luciferase with oligomeric proteins, it is possible to make several predictions. First, it is common for oligomeric proteins to assemble in one of a variety of well-defined patterns of symmetry.103 That is, homodimers usually assemble in a fashion that establishes a clear twofold rotation axis of symmetry. While the luciferase a and 3 subunits are distinctly different, they are clearly homologous and therefore likely to assemble in the same motif as for a homodimer, that is, with a pseudo-twofold rotation axis of symmetry, not unlike the pseudo-twofold axes of the hemoglobin tetramer that arise from the nonidentity of the a and 3 subunits of that protein and their tetrahedral packing array (see Reference 103 for a review). It therefore appears reasonable to suggest that the subunits of luciferase associate to form a dimer with a pseudo-twofold rotation axis, with the symmetry-related regions of the two subunits being those regions that are evolutionarily related. The regions of strong sequence conservation between the two subunits, then, would be primarily those regions that are involved directly with the structure of the active center or the interface between the two subunits. This hypothesis is based on the well-documented fact that the most strongly conserved regions of proteins are those which are involved directly in the biological activity and, with multimeric proteins, those regions that are involved in intersubunit contacts. We therefore suggest that the amino acid residues that are conserved between the a and the 3 subunits should be likely candidates for the intersubunit interface. Specifically, major components of the interaction interface between the two subunits may be contributed by the extended regions of sequence invariability, namely residues 36 to 53, residues 80 to 104, and/or residues 174 to 194. 2. Regions of Sequence Conservation A direct comparison of the a subunits of the four luciferases (Figure 14) shows that at 194 positions (55%), the four polypeptides are identical. A similar comparison of the 3 subunit sequences reveals that 118 positions (36%) are identical. The more demanding conservation of the sequence of the luciferase a subunit is consistent with the proposed participation of the a subunit both in the formation of the active center and in the direct interactions with the 3 subunit discussed above. In making the comparisons between the a subunits, and between the 3 subunits, we employed a simple similarity index scale in which a score of 1 was assigned at a given position if the four a (or four 3) subunits were identical at that position, a score of 0 if one or more subunits differed but the substitution(s) were conservative, and a score of — 1 if there were one or more nonconservative substitutions at that position. The score was then summed for a 5-residue window and that score assigned

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FIGURE 15. Sequence similarity index plot. The calculation of the "sequence similarity index", as described in the text, is based on a sliding window of 5 amino acid residues, and considers three conditions: identity, conservative substitutions, and substitutions with chemically dissimilar residues. The locations of the reactive thiol of the V. harveyi a subunit, the protease labile regions (PLR), and the lesions in AK-6 and AK-20 are indicated. The a subunit is symbolized by a solid line and the fi subunit by a dashed line. The apparent deletion from the p subunit is evidenced by the discontinuity of the dashed line in the region around residues 260 to 290.

to the first (N-terminal) residue within the window. The window was then displaced one residue toward the C-terminus and the process repeated, until the C-terminus of the subunit was reached. This process was carried out for both the a subunits and the p subunits, and the results are displayed graphically in Figure 15. In this presentation, a similarity index of 5 means that the residues at that position and at the following four positions are identical for the 4 proteins. There are several regions of the a subunit that appear, based on this analysis and other data, to be of special interest. First, the regions discussed above (positions 36 to 53, 80 to 104, and 174 to 194) that were highly conserved between the a and (3 subunits are, of course, conserved among the a subunits. In addition, there is a region spanning residues 109 to 120 that is highly conserved in the a subunits but is not conserved in the P subunits. On the basis of mutant enzyme analysis, chemical modification, and limited proteolysis, this area appears to comprise a critical portion of the active center, a point which will be discussed in detail in Section III.C. The region from residues 125 to 140 is of interest because this region of the a subunit is the least well conserved. In comparison, the same region in the p subunits is better conserved. Residues 190 through 230 of the a subunit appear to be strongly conserved, while the homologous positions in the p subunits do not appear to be so critical. The only residue in this region of the a subunit that is known to affect the structure of the active center is at position 227, and that position is not thought to be within the active center.104 3. An Internal Deletion Within the P Subunit, or an Imprecise Gene Duplication? Residues 230 to 257 appear to be relatively unimportant for both subunits, but residues 258 through 340 of the a subunit comprise a prolonged region of strong sequence conservation. Residues 258 to 286 of the p subunit are lacking, apparently due either to a deletion or to unequal crossing over at the time that the p subunit was originally formed.99'100 It is interesting to note that the region that is "missing" from the p subunit is one of the more

494

Chemistry and Biochemistry of Flavoenzymes TABLE 2 Percent Identity Between Pairwise Comparisons of Subunits of Luciferases from V. harveyi, V. fischeri, P. leiognathi, and X. luminescens

V. harveyi a V. fischeri a P, leiognathi a X. luminescens a V. harveyi p V. fischeri p P. leiognathi (3 X. luminescens 3

V.h. a

V.f. a

/>./. a

*./. a

V.h. (5

V/- P

/>./. P

100 64 61 86 32 31 30 29

100 76 66 31 31 26 25

100 62 28 30 25 25

100 30 33 28 30

100 51 49 60

100 63 53

100 49

*./. P

100

/Vote: The sequence data from which this table was derived is contained in Figure 14. The original data for the sequence of the V. harveyi luxAB genes was from References 22 and 23, the V. fischeri ATCC7744 luxAB sequence was from Reference 25, and the X. luminescens sequence was from Reference 26. Column headings indicate V. harveyi (V.h.); V. fischeri (V.f.); P. leiognathi (P.1.); and X. luminescens (X.I.).

highly conserved regions in the sequence of the a subunit, and it comprises a conserved structural feature of the luciferase a subunit, the protease labile region.20-105 no This region, which is thought to be a feature of the active center, will be discussed below. Finally, it should be pointed out that the region of greatest sequence divergence is at the C-terminus (about 25 residues) of the P subunits, a point of interest in the folding of the subunits and assembly of the heterodimer.24 4. Taxonomic Relationships Among the Luciferase-Producing Bacteria One of the common uses of amino acid sequence data is for taxonomic studies of the organisms that carry the proteins, and it was with great interest that we awaited the determination of the sequence of the luxAB genes from the terrestrial bacterium Xenorhabdus luminescens. The genes were cloned by Dr. Kenneth Nealson and his collaborators33 and the sequence of the DNA has recently been determined by Dr. Timothy Johnston and his students.26 Perhaps surprisingly, the "terrestrial" (X. luminescens) luciferase subunits are not very different from any of the luciferases from marine bacteria, and in fact are very similar to those of the well-studied V. harveyi (Figure 14). To aid in quantitative evaluations of the relatedness of the various luciferase subunits, the percent identity for each pairwise comparison has been determined and tabulated (Table 2). Fully 86% of the residues in the a subunit of the luciferase from X. luminescens are identical to the residues in the a subunit of the luciferase from V. harveyi, while a comparison between V. harveyi a subunit and V. fischeri a subunit revealed that only 64% of the positions are identical. V. harveyi and V. fischeri, two species of the Vibrio genus, are less similar than V. fischeri is to P. leiognathi (76% identity). It is clear that, based on the sequence of the lux genes, V. harveyi and X. luminescens belong in one genus while V. fischeri and P. leiognathi belong in another genus. A graphical representation of the results of this comparison is shown in Figure 16. The conclusions are supported by comparisons of the P subunit sequences as well (Table 2). C. RESIDUES CONTRIBUTING TO THE ACTIVE CENTER There are two regions of the V. harveyi luciferase a subunit that are believed, on the basis of chemical modification, limited proteolysis, spectroscopic studies, and mutant enzyme analyses (discussed below), to contribute to the structure of the active center. These include roughly residues 105 to 120 and residues 180 to 190. Three specific sites at which amino acid substitutions are known to cause dramatic alterations in the activity of the enzyme are positions 106, 113, and 227.

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FIGURE 16. Phylogenetic relationship based on luxAB sequence similarity. The degree of sequence identity between the luciferases from V. harveyi, X, luminescens, V. fischeri and P. leiognathi (Table 2) indicates that V. harveyi and X, luminescens are more closely related than either is to V. fischeri or P. leiognathi, suggesting that the lineage of present-day V. harveyi and X. luminescens diverged more recently that did the organisms that gave rise to present-day V. fischeri and P. leiognathi.

I . The Reactive Thiol Residue Chemical modification studies nearly 20 years ago showed that there is a highly reactive thiol (one of 14 in the heterodimer) on the a subunit of V. harveyi luciferase, in or near the active center.41-111 We isolated the tryptic peptide carrying the reactive thiol and determined the amino acid sequence to be Phe-Gly-Ile-Cys-Arg;112 when the entire a subunit sequence was determined, the peptide sequence permitted identification of the reactive residue as cysteine a!06.22 Its position is indicated by an arrow in Figure 15. Every thiol-modifying reagent so far utilized has resulted in inactivation of the enzyme. A partial list of the reagents and the apparent bimolecular rate constants for the reactions are given in Table 3. The observation that the rate of reaction of a family of A^n-alkylmaleimides with the reactive thiol is strongly dependent on the chainlength of the alkyl substituent led to the conclusion that the thiol residues in or near a highly hydrophobic cleft in the surface of the enzyme. 111 This observation was confirmed by Merritt and Baldwin, 113 who used nitroxide spin-labeled maleimides to probe the hydrophobicity and size of the cleft associated with the reactive thiol, as well as by Dougherty et al., 116 who used 7V-[/?-2(benzoxazolyl)phenyl]maleimide. The pK a of the reactive thiol, 9.4, is too high to allow correlation with any of the activity-associated protonation/deprotonation processes,41 and no functional role has ever been demonstrated for the group. In fact, we have always been careful to point out41 that inactivation by chemical modification does not by any means demonstrate "essentiality" of a specific group for substrate binding or catalysis. Definitive proof that the reactive thiol is not required for the activity of luciferase finally came in the 1980s from the sequences of the luxA genes from V. fischeri and P. leiognathi and from site-directed mutagenesis of the V. harveyi luxA gene. Although the enzyme from X. luminescens, like that of V. harveyi, has a cysteinyl residue at position a!06,26 both the V. fischeri15 and P. leiognathi25'31^2 luciferases have valyl residues at that position. It is interesting to note that, not surprisingly, the luciferase from V. fischeri is far less sensitive to thiol-directed reagents than is the enzyme from V. harveyi,"-ns apparently as a result of the Cys/Val difference at position a!06 between the two strains. An additional demonstration that the thiol is not "essential" for the activity of luciferase came from site-directed mutagenesis: Baldwin and colleagues86'87'99 replaced the codon for the cysteine at position a 106 with codons for serine, alanine, and valine. All three mutant enzymes are active. The serine and alanine mutants are excellent enzymes, having near wild-type quantum yield, albeit somewhat reduced FMNH2 affinity,86'99 but the enzyme with valine at position a!06 has a reduced affinity for aldehyde and

TABLE 3 Structures of ThioI-Directed Reagents Used to Modify Luciferase and the Second-Order Rate Constants for the Reactions

The apparent second order rate constants for each of the compounds listed above were determined at pH 7.0 and 23° to 25°C. The original literature references are as follows: lodoacetamide;41 2-bromo-n-decanal;114 MMTS43 (note that Reference 43 also reports the rate constants for a series of alkylalkanethiolsulfonates); 1,5IAEDANS and 1,8-IAEDANS, Ziegler and Baldwin (unpublished); MNBS;73 NEM, NBM, NHM, NOM and DDPM;111 ANM, Ziegler and Baldwin (unpublished); NBPM — use of this reagent with luciferase was first reported by Tu et al., 117 but the rate constant for the reaction was determined by Dougherty et al.;116 3(maleimidomethyI)-2,2,5,5-tetramethyl-l-pyrrolidinyloxyI and 3-(3-maleimidopropylcarbamoyl)-2,2>5,5-tetramethyl-l-pyrrolidinyloxyl;113N-(l-pyrene)maleimide — this reagent was reported by Tu et al. 117 to react with luciferase stoichiometrically, but the rate constant was not reported.

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Chemistry and Biochemistry of Flavoenzymes

an approximately 50-fold reduced quantum yield.86 As mentioned above (Section II.C), the peroxyflavin intermediate (intermediate II) of the valine mutant enzyme breaks down very rapidly,87 effectively uncoupling the flavin oxidation from aldehyde hydroxylation/light emission. Paquatte and Tu,73 studying luciferase chemically modified (inactivated) by introduction of a methyl group onto the thiol at position a 106, were unable to isolate any peroxyflavin intermediate by low-temperature molecular sieve chromatography, but they were unable to distinguish between a defect in formation of the intermediate and a defect in its stability. It is tempting to speculate that perhaps the defect resulting in activity loss for the S-methylated enzyme is similar to that of the a 106 valine mutant: extreme instability of the peroxyflavin intermediate. The a!06 alanine and valine mutants reported by Baldwin's laboratory86'87-99 were subsequently constructed by Xi et al.,85 who reported flavin and aldehyde binding properties and quantum yields similar to those presented in the earlier publications. The relatively high quantum yields of the enzymes from V. fischeri and the Photobacterium species, all of which have valine rather than cysteine at a 106, and also of the mutant V. Harveyi enzymes with serine or alanine at a 106, have demonstrated unequivocally that the V. harveyi a 106 cysteine is not required for catalytic activity. The activity of the three mutant V. harveyi luciferases (Ala, Ser, and Val at a 106) is much less sensitive to alkylating reagents than is the wild-type enzyme, confirming that the thiol at position a 106 of the wild type is the most reactive target of the alkylation reaction. Like the enzyme from V. fischeri,"Jls the a!06 mutant luciferases were found to be inactivated by thiol directed reagents, but at a greatly reduced rate.99 This finding suggests that alkylation of at least one other thiol on the enzyme can cause inactivation, but to our knowledge, no experiments to delineate the location of the other putative "essential" cysteinyl residue(s) have been reported. The effects of ligand binding on the rate of thiol modification with different modifying reagents and the effects of modification on ligand binding affinities are interesting, but not simple. Long-chain aldehydes have been shown to offer some protection against modification by A^-ethylmaleimide,41 2-bromo-n-decanal,114 l-diazo-2-oxoundecane,119 and methyl p-nitrobenzene sulfonate.73 However, after methylation with the latter reagent, the enzyme still binds decanal without a marked change in binding affinity. Oxidized flavin mononucleotide, a product of the enzymatic reaction, at concentrations much greater than its dissociation constant for luciferase, offers virtually complete protection against modification by iodoacetamide,120 and has also been shown to protect the enzyme against jV-ethylmaleimide modification.41 The active center of luciferase must be composed of contiguous or overlapping binding sites both for FMNH2 (and presumably for the product (FMN) and for aldehyde. Fried and Tu114 have reported the use of 2-bromo-n-decanal (Table 3) as an affinity label to modify the luciferase. They found that n-decanal protects the enzyme from inactivation by the reagent. While they reported that the enzyme is not protected from 2-bromo-n-decanal by FMN, in fact they used FMN at 0.2 mM, which was only about 0.5 Kd for the enzyme:FMN complex72 under the conditions of their experiment, so it is not surprising that they did not observe significant protection of the reactive thiol by FMN. However, even at concentrations well in excess of Kd, FMN does not protect the enzyme against modification by methylmethanethiolsulfonate115 or by methyl p-nitrobenzene sulfonate;73 indeed, enzyme modified even by A^-n-octylmaleimide, which introduces the bulky A^-octylsuccinimido group onto the thiol, still binds FMN with nearly the same affinity as the native enzyme, suggesting that there is no direct interaction between FMN and the thiol group.121 Paquatte and Tu73 have recently provided the first direct demonstration of protection of the thiol by FMNH2, in this case against inactivation by the methylating reagent methyl/7-nitrobenzene sulfonate. Ziegler-Nicoli et al.41 demonstrated that enzyme modified by Af-ethylmaleimide is greatly impaired in ability to bind FMNH2 (Kd increased by > tenfold). It has been possible to demonstrate weak FMNH2 binding to enzyme modified by Af-rc-octylmaleimide, by pertur-

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bation of the circular dichroism spectrum of the bound flavin, but neither the precise affinity nor the stoichiometry were measurable.121 It is possible that the weak FMNH2 binding to the modified enzyme detected by circular dichroism is related to the weak, apparently less specific binding of a second reduced flavin detected by 13C NMR69 (see Section II.B.I above). Paquatte and Tu73 have shown that luciferase inactivated by methylation with methyl /^-nitrobenzene sulfonate can still bind FMNH2 in 1:1 stoichiometry with an affinity similar to that of native enzyme. The reactive thiol has been modified with a variety of reagents and used as a molecular marker to probe structural changes in the vicinity of the active center and to attach probes for fluorescent energy transfer measurements of distances between the thiol and ligand binding sites. Tu et al.117 attached ^-[p-CZ-benzoxazolyOphenylJmaleimide and A^-(l-pyrene)maleimide to the luciferase through the reactive thiol and used energy transfer measurements from the covalently attached probes to bound 8-anilino-l-naphthalenesulfonate (ANS) to calculate the distance from the reactive thiol to the ANS binding site. The calculated distance was in the range of 21 to 37 A. With bound ANS as donor and bound FMN as acceptor, the distance from the ANS site to the FMN binding site was calculated from fluorescence lifetime measurements to be in the range of 30 to 58 A. These measurements are consistent with the model in which the reactive thiol is located in or near the active center, which is removed some significant distance from the ANS binding site. 2. Reactive Histidinyl Residue and ot-Amino Groups The luciferase from V. harveyi has been shown by Cousineau and Meighen122'123 to be inactivated by ethoxyformic anhydride modification of a histidinyl residue on the a subunit; the residue is protected from modification by both long chain aldehydes and FMN, and is thought to reside in or near the active center. No functional role has yet been demonstrated for the histidinyl residue, and in fact no one has yet deduced which of the 11 histidinyl residues on the a subunit22 is the modified residue. However, the pKa of the reactive imidazole, 6.8,122 is in the same range as the two pKa values manifested in the pH dependence of FMNH2 binding, about 6.2 and 6.8,41 and the histidinyl residue might in fact provide one of the hydrogen bonding or positively charged functional groups postulated by Vervoort et al.69 to interact with the isoalloxazine moiety of the bound reduced flavin anion; alternatively, it might be involved in binding the 5' phosphate of the ribityl chain. Modification of V. harveyi luciferase by 2,4-dinitrofluorobenzene inactivates the enzyme, apparently by reaction not with the thiol, but with the a-amino groups of both subunits; the enzyme is protected from inactivation by long-chain aldehydes and by FMN.42 Modification of the a-amino group of either a or (3 inactivates the enzyme, and no FMNH2 binding to the modified enzyme could be detected, suggesting that either the a-amino group of the (3 subunit lies close to the active center, or its modification causes a conformational perturbation in the active center.42 3. Studies of the Active Center by Limited Proteolysis It has been known for many years that bacterial luciferase is highly sensitive to inactivation by proteases.105 In fact, luciferases comprise an excellent, highly sensitive assay for a broad range of proteolytic enzyme activities.124 Limited proteolysis of the luciferase from V. harveyi has been the most thoroughly investigated,20-105-108 but comparative studies suggest that the phenomena observed for the V. harveyi enzyme are generally applicable to all bacterial luciferases.106-107 Treatment of luciferase in dilute buffers with low concentrations of proteases results in a rapid loss of activity. Concomitant with this inactivation is conversion of the a subunit from a circa 42-kDa polypeptide (SDS mol wt) to a family of polypeptides (referred to collectively as the y fragments) of about 28 kDa, and to another set of fragments of about 14 kDa known as the 8 fragments.20-115 The y fragments generated by the limited

500

Chemistry and Biochemistry of Flavoenzymes

action of trypsin range in molecular weight from approximately 27,000 to 28,500 and are about 5 in number, while there are only 2 'y fragments resulting from chymotrypsin treatment, differing in molecular weight by only 200 to 400.20-108 If the luciferase is labeled at the reactive thiol with radioactive yV-ethylmaleimide and then treated with trypsin or chymotrypsin, the radiolabel appears in the y family of fragments, and by Edman degradation of y fragments eluted from SDS gels, Rausch 20 ' 115 has demonstrated that the y family of fragments has a unique N-terminal amino acid sequence that is the same as that of the a subunit. These observations demonstrated that the sites for the initial proteolysis events, generating the y fragments, reside near residues 250 to 300 in the primary structure of the a subunit. Inspection of the amino acid sequence in that region shows that there are many potential trypsin sensitive sites, consistent with the observed multiplicity of tryptic fragments (Figure 14). Rausch eluted several of the 8 fragments generated by the action of chymotrypsin from SDS gels and determined N-terminal amino acid sequences which were consistent with the primary cleavage sites for that enzyme being the phenylalanyl-valyl bond between residues 280 and 281 and the leucyl-lysyl bond between residues 282 and 28320'22-115 (see Figure 14). Continued incubation of the inactive luciferase with chymotrypsin results in conversion of the y fragments to a fragment of about 10,000 mol wt (8L), which was shown by radiolabeling to contain the reactive thiol, and a second fragment of about 17,000 mol wt which Rausch designated 8 H . 20JI5 The N-terminal amino acid sequence of the 8L fragment was the same as that of the a subunit (and of the y family), and the N-terminal amino acid sequence of the 8H fragment showed that it had resulted from chymotryptic cleavage of the phenylalanyl-glycyl bond separating residues 117 and 118.20'22 Near the N-terminus of the yH fragment, Rausch discovered the sequence Met-Asp-Cys-Trp-Tyr-Asp (residues 128 to 133 of the a subunit). This sequence, containing Met and Trp, which have unique codons, and Asp, Cys, and Tyr, which have two base redundancy, allowed us to construct a probe for the cloning of the first luciferase gene.1 Treatment of the luciferase from V. harveyi with a variety of proteolytic enzymes yields families of fragments of similar size to those described in detail for trypsin and chymotrypsin. 124-125 It is therefore assumed that all proteases act at the same region of the luciferase structure. With all proteases studied, the rate of inactivation is indistinguishable from the rate of hydrolysis of the first bond in the a subunit which results in the loss of the intact a polypeptide.105 The inactivation of luciferase is therefore due directly to the hydrolysis of bonds within the region of residues 280 to 285 or to a fast but subtle conformational change that follows the hydrolysis reaction. Analysis of the products of the limited action of trypsin and of chymotrypsin in an analytical ultracentrifuge under nondenaturing conditions showed that the proteolytic fragments do not dissociate, but remain associated in a complex of about 75,000 mol wt.105 Extended incubation of the inactive luciferase with proteases does not lead to gross degradation, indicating that the fragments that result from the initial proteolytic cleavages do not undergo major rearrangements. Circular dichroism measurements, sedimentation velocity measurements, and reactivity of protein thiols with DTNB, all suggest that the inactive luciferase has a slightly more open structure than the native enzyme, but that the changes in structure are not major.108 4. Location of the Protease Labile Regions Relative to the Active Center The issue of the location of the protease labile regions of the luciferase relative to the active center has been addressed through several experimental approaches. First, the binding of FMN to the luciferase stabilizes a conformation that has reduced sensitivity to the action of proteases.108-121'125 Second, phosphate and a series of other multivalent anions stabilize the luciferase to inactivation by proteases. 107>125-126 Phosphate also has a profound effect on the quantum efficiency of the bioluminescence reaction when reduced riboflavin, rather than

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FMNH 2 , is used as a substrate.74 Meighen and MacKenzie74 have demonstrated using flavin analogues with modifications in the ribityl chain that luciferase requires a negative charge approximately 5 carbons removed from the isoalloxazine ring both for good binding to luciferase and for a high quantum yield. They demonstrated that addition of orthophosphate to the reaction of luciferase with reduced riboflavin results in an enhanced quantum yield but does not alter the binding of the reduced riboflavin, which has an apparent binding affinity about 103-fold weaker than that of FMNH2. Their observations suggest that the binding of the anion stabilizes a conformational rearrangement to a high quantum yield form of the enzyme. The effect of phosphate and the other multivalent anions on the sensitivity of the luciferase to proteolytic inactivation is consistent with this interpretation and suggests that the regions of the enzyme which constitute the protease labile regions become less exposed in the phosphate complex.107-125'126 Another series of observations consistent with the suggestion that the protease labile regions constitute a portion of the active center came from limited proteolysis of mutant forms of luciferase which have altered active centers as reflected in altered enzymatic activity.104*108 With these mutants, analysis of the protease sensitivity suggested that subtle changes in the active center result in changes in the protease sensitivity. For example, with the luciferase from AK-20, a substitution of Phe for Ser at position a227 results in an increased sensitivity of the "slow" protease labile region around residues 115 to 120, such that the y family of polypeptides does not accumulate as it does with the wild-type enzyme.104 Ziegler-Nicoli et al.41 demonstrated that following a single catalytic cycle the luciferase was in an altered conformational state. The altered conformational state was observed at temperatures near 0°C by injection of FMNH2 into an aerobic solution of enzyme. Secondary addition of A^-ethylmaleimide did not yield the biphasic inactivation of the enzyme that was expected based on the known rate of turnover of the enzyme at that temperature. Instead, an apparent first order inactivation was observed that was much slower than the rate required for the flavin to oxidize on the luciferase to yield FMN, which binds weakly to the enzyme and offered only marginal protection under the conditions of the experiment.41 AbouKhair et al. 127 - 128 have extended this observation to demonstrate a slow return of the enzyme to its initial state of sensitivity to inactivation by Af-ethylmaleimide. In their experiments, FMNH2 was rapidly mixed with the luciferase at 0°C and the mixture then applied to a column of Sephadex G-25. The luciferase which eluted from the column was largely free of flavin, but retained the reduced sensitivity to inactivation by Af-ethylmaleimide. This form of the enzyme was also insensitive to trypsin and chymotrypsin. If the addition of Af-ethylmaleimide or protease to the luciferase was delayed, a biphasic inactivation was observed. In the initial rapid phase, the rate of inactivation was the same as for the original enzyme. The rate of the slow phase stayed the same regardless of the time between mixing of FMNH2 with luciferase and the addition of the inactivating agent, but decreased in amplitude. These observations demonstrated that the luciferase undergoes a significant conformational change associated with the binding of FMNH2, conversion of the flavin to intermediate II and the release of FMN. At temperatures near 0°C, this altered conformational state relaxes slowly to the original state of the enzyme, significantly slower than the dissociation of the FMN. The altered conformational state is characterized by insensitivity to inactivation by A^-ethylmaleimide and to the limited action of proteases.41'127-128 These observations strongly support the suggestion that the protease labile regions actually comprise a portion of the active center of the luciferase and that these regions undergo structural rearrangements during catalysis. 127 - 128 The proximity of the "slow" protease labile region (residues al!5 to 120) and the reactive thiol (position a 106) support this interpretation (see Figures 14 and 15). The "fast" protease labile region (residues a280 to 285) apparently approaches the "slow" protease labile region in three-dimensional space such that the two regions together form a cationic surface that could facilitate flavin binding and/or docking of one of the accessory

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emitter proteins.1(X96'97'129'130 It is possible that catalytic turnover-associated conformational rearrangements within this region of the luciferase are critical to the functioning of the enzyme in vivo, where interactions with other proteins are required for the biological function. Using spectroscopic methods, Tu et al. 117 have calculated the rotational correlation time of the luciferase-ANS complex and the derivatizedvV-(l-pyrene)succinimidyl-luciferase and found it to be 47 ± 2 ns at 23 ± 0.5°C, significantly longer than the value of 28.6 ns calculated assuming a spherical structure. These authors concluded that the luciferase-dye complex, and probably the native luciferase, has an asymmetric structure, not unreasonable for a molecule composed of two interacting subunits. For the native luciferase, Lee et al.90 have measured rotational correlation times of 62 ns or 74 ns at 2°C, using the anisotropy of decay of the intrinsic fluorescence or the fluorescence of luciferase-bound ANS, respectively. However, analysis of the decay of anisotropy of fluorescent species that form at 2°C upon mixing of FMNH2, O2 and luciferase (presumably intermediate II; see Figure 7) yielded a rotational correlation time of about 100 ns. These observations led Lee and his colleagues to propose that the luciferase undergoes a major conformational change during the catalytic cycle, such that it has an increase in axial ratio of 50%.w The proposal is consistent with the earlier observations of Ziegler-Nicoli et al.41 and AbouKhair et al. 127 - 128 and suggests that determination of the rotational correlation time using the intrinsic fluorescence of the form of the enzyme studied by AbouKhair et al.127'128 should yield a value of about 100 nsec. Based on the foregoing considerations, it has been proposed108'110 that the active center of the luciferase is composed in part of residues within the two protease labile regions indicated in Figures Hand 15. It is clear that during the catalytic cycle, the enzyme undergoes a structural rearrangement that involves, at least in part, residues within the protease labile regions. One consequence of the conformational change is a marked reduction in the reactivity of the thiol residue at a 106, a residue that also appears to be in or very near the active center of the enzyme. 5. Investigation of the Properties of Mutant Forms of Luciferase By random mutagenesis, Cline and Hastings78 generated and characterized a very large collection of mutant forms of luciferase from V. harveyi having altered reaction kinetics. Based on the large body of data generated by Cline and Hastings,78 the suggestion has emerged that the active center of the enzyme resides primarily, and perhaps exclusively, on the a subunit, an issue discussed in more detail below. In an effort to obtain more information concerning which portions of the enzyme contribute to the structure of the active center, luxA genes of two of the mutants characterized by Cline and Hastings78 have been cloned. A mutant that is defective in flavin binding, AK-6, was cloned by Lawrence Chlumsky,131 and a mutant defective in aldehyde binding, AK-20, was cloned by Lorenzo Chen.104 The mutant AK-6 results from the substitution of Asn for Asp at position al!3.131 The luciferase from AK-6 has a greatly reduced affinity for reduced flavin mononucleotide, but a normal aldehyde binding affinity. 78 The spectrum of bioluminescence from AK-6 luciferase is red-shifted approximately 12 nm, and the pH-activity profile is strongly acid shifted; at pH values above 7.0, the lifetime of the intermediate II on the AK-6 luciferase is dramatically lengthened, reaching a half-life of tens of seconds at pH 8.O.132-133 The charge change in or near the active center suggested that the reactive thiol at position a 106 might have altered reactivity or ionization properties in this enzyme, but it was found to have the same pKa as the thiol of the wild-type enzyme, about 9.441 (Baldwin, unpublished). Chlumsky has shown that substitution of the al 13 Asp by either Glu or Cys results in an enzyme with near-normal catalytic properties, suggesting that the ability of a residue in that position to form an anion might be crucial.87'99 He has also incorporated numerous other substitutions at position al!3, and all but Asp (the wild type), Glu and Cys have greatly reduced activity in the bioluminescence reaction.87-99

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Based on the studies of Chlumsky, 87 ' 89 it appeared likely that chemical modification of the thiol at position al!3 of the al 13Cys mutant with reagents such as iodoacetamide or AT-ethylmaleimide would render the enzyme inactive, especially since the thiol at a 106 is highly reactive, and the nature of the residues surrounding al 13 (wild-type sequence Lys U2 Asp-Phe-Arg-) suggests that the residue should be near the surface of the enzyme. However, when Chlumsky introduced two mutations into the luciferase, al!3Cys and cd06Ala, and attempted to modify the new cysteinyl residue, he found, surprisingly, that the cysteinyl residue at position al!3 is not reactive with alkylating reagents.87 In a similar series of experiments, Lorenzo Chen cloned the lux A gene of the mutant AK-20 described by Cline and Hastings78 as an aldehyde binding mutant. The lesion in the luciferase from AK-20 was shown to be a227 Ser -» Phe.I04 The effect of the mutation was to reduce the quantum yield of the reaction, enhance slightly the affinity for reduced flavin mononucleotide, reduce the sensitivity to aldehyde substrate inhibition, and greatly alter the aldehyde chain length dependence of the bioluminescence decay.78-104 Unlike the wild-type luciferase, which exhibits a more rapid turnover with n-decanal as substrate than with either n-octanal or n-dodecanal, the enzyme from AK-20 has a faster decay of bioluminescence with both n-octanal and n-dodecanal than with n-decanal.78 In contrast with the effects found by Chlumsky for substitutions at position a 113 (see above), Chen found that the a227-Ser could be replaced by a variety of amino acid residues with minimal effect on the catalytic efficiency of the enzyme.104 Based on the properties of the collection of variants at position a227 generated and characterized by Chen, it appears that this position is not an integral part of the active center, but instead is in a region of the protein that is very forgiving of large changes in the size and chemical properties of the amino acid side chain. The most likely structural element for such flexibility would be a surface loop, such that the protein could fold properly regardless of the amino acid residue at that site. The observed effects of the amino acid substitutions at position a227 could be the consequence of subtle changes in structure at the active center that result from changes in the chemical properties of residues at that position. Chen and Baldwin104 therefore proposed tentatively that position a227 is close in the three-dimensional structure to the active center, but not a component of the active center, and probably resides in a surface loop that has significant structural flexibility. 6. Location of the Active Center Relative to the Subunit Interface Relatively few experiments have been reported that bear on the question of the location of the active center relative to the subunit interface of the luciferase. In the early 1970s, Meighen et al. 134 - 135 demonstrated, with hybrid luciferases consisting of native subunits and succinylated subunits, that modification of the a subunit virtually inactivated the enzyme, significantly altering several kinetic parameters, while succinylation of the p subunit had little effect on the activity or kinetic parameters. These results were interpreted as a demonstration that the active center of the enzyme resides on the a subunit. Of course, it could be argued from the viewpoint of 1991 that the only residues of the p subunit directly involved in the active center might be at the subunit interface, and that the subpopulation of p chains with those residues modified would have failed to form heterodimers, and thus would not have been studied with regard to kinetic parameters. Cline and Hastings generated numerous kinetic mutants by random mutagenesis, and all 20 of the mutants isolated in the first kinetic screen were a subunit mutants.78 Cline was very concerned that he did not find any mutants with lesions in the p subunit having altered enzymatic activity, even though it was believed that both subunits were required for activity (T. W. Cline, personal communication). Cline and Hastings did report numerous mutants having lesions in the p subunit that had temperature sensitive luciferases,78'136 but no p defective mutants were found on the basis of altered enzymatic activity, until an altered

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screen was devised, by which mutants defective in binding of reduced flavin mononucleotide were isolated.133 Through this rigorous screen, Cline was successful in isolating a single (3-defective flavin binding mutant, FB-1, as well as two more a-defective flavin-binding mutants. 133 Unfortunately, FB-1 is a strikingly thermally sensitive protein, suggesting that the structure of the protein has been perturbed significantly by the mutation, and thus that the effect of the p mutation on the flavin-binding site might be through the subunit interface.133 The failure to obtain any stable p mutant luciferase with altered kinetic properties, despite expenditure of significant effort to search for such mutants, argues very strongly that the p subunit contributes few if any residues to the structure of the active center. Meighen and Bartlet17 formed a mixed species hybrid in vitro using the a subunit of V. harveyi luciferase and the p subunit of P. phosphoreum luciferase. The kinetic properties of the hybrid luciferase reflecting the decay of intermediate II and the aldehyde chain length dependence of the reaction were essentially the same as for the luciferase from V. harveyi, the source of the a subunit, but the affinity for the reduced flavin was lower than that of V. harveyi, closer to that of the P. phosphoreum luciferase.17 As with the FB-1 mutant described by Cline,133 the hybrid luciferase described by Meighen and Bartlet provided direct evidence that modification or alteration of the p subunit can affect the kinetic properties of the enzyme. Based on the properties of the hybrid luciferase, Meighen and Bartlet proposed that both subunits might be involved with the binding of FMNH2, whereas subsequent steps in the mechanism would be the province exclusively of the a subunit and would be unaffected by alterations in the p subunit.17 The FB-1 luciferase was generated by nitrosoguanidine treatment of V. harveyi and was therefore probably a point mutant, whereas the substitution of the P. phosphoreum p subunit for the V. harveyi p subunit resulted effectively in substitution of at least 40% of the amino acid residues in the P subunit (see Table 2). The two subunits obviously interact, and the high quantum yield activity of luciferase requires both subunits, so it is expected that structural changes in one subunit should have effects on the other subunit. It thus is possible, and perhaps indeed most reasonable, to interpret the behavior of both the p subunit-defective flavin-binding mutant FB-1, which was also markedly temperature sensitive, and the hybrid luciferase as indicating that alterations in the p subunit can alter the structure of the active center through interaction with the a subunit, rather than suggesting that the p subunit contributes residues directly to the flavin-binding pocket. In a related series of experiments, Welches and Baldwin42 showed that 2,4-dinitrofluorobenzene inactivates luciferase at a 1:1 stoichiometry. The incorporated label was distributed between the two subunits, about 60% on a, indicating that reaction of the reagent with one subunit blocked reaction with the other subunit.42 It appeared that the reagent reacted not with the reactive thiol, but with the a amino groups of both subunits. The enzyme was protected from the inactivation reaction by binding of FMN, both subunits being protected, suggesting that the a amino groups of both subunits are in or near the active center. This result indicated that the N-terminal methionyl residue of the p subunit should reside near the active center.42 However, recent reports from several laboratories of formation of a biologically active fusion polypeptide between the two subunits clearly calls into question such an interpretation.62-63 Rather, it now appears more reasonable to propose that the Ntermini of the two subunits reside in or near the subunit interface and that formation of the a-dinitrophenyl derivative perturbs the active center through perturbations in the interactions between the two subunits. The only direct experiments regarding the location of the active center relative to the subunit interface that have been published are chemical modification experiments by Tu and his co-workers,114'119-137 and cross-linking experiments reported only in the dissertation of S. K. Rausch.115 Tu and Henkin119 have reported an elegant experiment using a radiolabeled photoaffinity label, l-diazo-2-oxoundecane, which they found at low concentration to be competitive with respect to n-decanal with Kt = 0.7 ± 0.1 p-M. In their labeling experiments,

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they employed reagent at 65 fxM, or about 100 times the K i; and a sixfold molar excess over luciferase. Photoactivation of the diazo group resulted in inactivation of the luciferase. Including the substrate n-decanal at 500 jjiM in the photoinactivation reaction slowed the rate of inactivation by only about twofold. Following inactivation (in the absence of aldehyde), the label was found on both subunits, with a slight excess on the a subunit. There was good correspondence between the total incorporation of label and the fractional inactivation of the luciferase. These very difficult experiments were ably performed and the data clearly demonstrate incorporation of label into both subunits. However, interpretation of these observations is complicated by the fact that the highly reactive carbene species generated in the photoactivation reaction can be inactivated by reaction with water, and the efficiency of labeling of a specific site would therefore be affected by the efficiency with which water is excluded from the site. Furthermore, the binding of aldehyde to the free enzyme appears to be different from binding of aldehyde to the enzyme:flavin complex (see discussion in Sections II.A and II.B, above, and Table 1) which occurs in the competitive binding assays involving l-diazo-2-oxoundecane and n-decanal as described by Tu and Henkin.119 The finding that 500 |xA/ n-decanal (see Table 1) did not reduce the rate of inactivation by more than a factor of 2 raises questions about the relative disposition of the aldehyde-binding site and the binding site for the reagent. Finally, with such a high concentration of reagent relative to the K i? the possibility of nonspecific binding becomes problematic. As a result of all of the uncertainties discussed here, the conclusion of the authors that the aldehyde binding site is at or near the subunit interface appears to be only weakly supported by the data. Paquatte et al.137 have employed three isomeric bifunctional reagents, 0-(2'-pyridyldithio)benzyl[l-14C]diazoacetate and the meta andpara isomers, to form mixed disulfides with the luciferase through the reactive thiol. Photoactivation of the diazo group following covalent attachment of the reagent to the a!06 thiol, and subsequent reduction of the mixed disulfide with dithiothreitol, resulted in the vast majority of the label remaining with the a subunit, showing that the photochemical cross-linking occurred with relative efficiency (about 25 to 33%) with residues on the a subunit when the reagent was attached to the reactive thiol at position a 106. Reduction of the mixed disulfide prior to photoactivation of the diazo moiety and gel filtration resulted in removal of most of the reagent, but about 8 to 14% of the reagent remained associated with the enzyme. The stoichiometry of attachment to the a subunit via the mixed disulfide was 1.02, 0.98, and 0.80 mol/mol subunit for the ortho, meta and para isomers, respectively. The labeling of the p subunit by the mixed disulfide was 0.04, 0.05, and 0.04 mol/mol subunit for the same reagents. Photoactivation, reduction of the disulfide and subunit separation resulted in a labeling stoichiometry of the a subunit of 0.25, 0.27, and 0.26 mol/mol subunit and for the p subunit, 0.05, 0.05, and 0.04 mol/ mol for the ortho, meta and para isomers, respectively.137 Given the nonspecificity of the chemical labeling reactions and the fact that between 0.08 and 0.14 mol of reagent remained associated with the enzyme following reduction but in the absence of photochemical crosslinking, this experiment could be interpreted as a demonstration that the closest approach of the 3 subunit to the reactive thiol is more than about 10 A; however, the authors chose to interpret the low-level labeling of the P subunit as proof that the P subunit is close to the reactive thiol.137 In a different attempt to demonstrate proximity of the reactive thiol to the subunit interface, Rausch115 employed the bifunctional cross-linking reagent p-azidophenacylbromide. Attachment of the reagent to the reactive thiol followed by photochemical activation of the azido functionality should have led to cross-linking of the subunits if the thiol were close to the subunit interface. Rausch was unable to demonstrate any cross-linking and concluded that the closest approach of the p subunit to the thiol must be greater than 13 A. It is clearly difficult to build a strong case on negative results, so the conclusions of the Rausch experiment

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have appeared only in his dissertation.115 While it is possible that the reactive thiol is in the aldehyde binding site, that the aldehyde binding site is at the subunit interface, and/or that the thiol is located near the subunit interface, we regard these issues as open questions, not yet rigorously demonstrated by experimental results. In our view, at the present, the most definitive statement justified by existing data is that the active center resides primarily, if not exclusively, on the a subunit. The direct participation of residues of the (3 subunit in the active center remains to be demonstrated. D. PROPERTIES OF BACTERIAL LUCIFERASES HAVING GENETICALLY FUSED SUBUNITS Recently, several laboratories have succeeded in fusing the coding regions of the lux A and luxE genes such that a single polypeptide is produced with the sequences of both the a and P subunits.62'63 The motivation of both laboratories was to create a single polypeptide with the activity of the bacterial luciferase enzyme for expression in eucaryotic cells. The requirement of producing two polypeptides with separate ribosome binding sites and other features has resulted in little interest in using the bacterial lux genes as reporters for eucaryotic gene expression. However, the availability of a single gene encoding both subunits which fold into a biologically active form should result in greater interest in luciferase gene marker studies for eucaryotic systems. In the process of generating a novel form of luciferase for eucaryotic expression, these investigators have also generated an interesting protein for use in studies of the folding and assembly of the luciferase. The fact that the single polypeptide folds into a biologically active form indicates that in the normal dimeric luciferase, the C-terminus of the a subunit must reside in close proximity to the N-terminus of the (3 subunit. Escher et al.62 have demonstrated that the monomeric form is active, and have reported a dimeric structure with about 150,000 mol wt, which is also active. These authors report that the folding of the fused luciferase is much more thermally sensitive than is the wild type. The mechanism of the greatly enhanced temperature sensitivity of the folding of the fusion polypeptide constitutes an interesting problem. Unfortunately, in the approximately 2 years since the activity of the fusion proteins was reported, no reports have appeared of the functional or physical properties of these interesting variants of the luciferase.

E. FOLDING OF THE SUBUNITS AND ASSEMBLY OF THE HETERODIMER The ability to reversibly unfold and refold the subunits of luciferase was developed in the early work of Friedland and Hastings,138'139 who demonstrated that luciferase is comprised of two nonidentical subunits that can be separated by chromatography in urea and recombined to form active enzyme. Their work was followed by that of Gunsalus-Miguel et al.,140 who described the purification and properties of luciferase from a new strain of bacteria,' 'MAV 1 ', which is now known as V. harveyi B392, and demonstrated that the luciferase subunits from V. harveyi (isolated by chromatography in urea) do not cross react immunochemically with, and cannot recombine with, the subunits of the luciferase from V. fischeri (formerly P. fischeri). 1. Refolding of Luciferase Subunits and Formation of Active Enzyme Since the initial studies of Friedland and Hastings138-139 and Gunsalus-Miguel et al.,140 several workers have employed the general approach delineated by Friedland and Hastings to accomplish in vitro subunit complementation. In his detailed investigation of mutant forms of luciferase, Cline relied heavily on the ability of the subunits to associate upon dilution from urea to assign the subunit location of lesions.78-136 Ziegler-Nicoli used subunit complementation as the initial step in determining the subunit modified in chemical modification experiments.41

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2. Formation of Hybrid Luciferases Meighen and Bartlet17 succeeded in obtaining an active hybrid luciferase consisting of the V. harveyi a subunit and the P. phosphoreum P subunit, which they characterized very thoroughly (see discussion in Section III.C.6). A more extensive subunit refolding experiment was performed by Ruby and Hastings,141 who reported the formation of hybrid luciferases from the subunits of luciferases from V. harveyi, V. fischeri, P. leiognathi, and P. phosphoreum. In these experiments, active hybrids were formed for all heterologous a-(3 combinations of the luciferases from V. fischeri, P. leiognathi, and P. phosphoreum except for the P. phosphoreum a:V. fischeri p combination. These authors were unable to obtain active hybrids involving subunits from V. harveyi with any of the other species. Meighen and Bartlet17 employed the NCMB 844 strain of P. phosphoreum, while Ruby and Hastings141 used the NZ-ll-D strain, and the difference in strains may be the source of the difference in results between the two labs, since both used the V. harveyi strain B392. Ruby and Hastings141 found that the enzymatic properties of the hybrid luciferases approximated those of the source of the a subunit, as would be expected if the active center were on the a subunit, a result consistent with those of Meighen and Bartlet.17 The p subunit of P. leiognathi luciferase conferred enhanced thermal stability on all hybrid luciferases containing that subunit.141 3. Subunit Dissociation and Reassociation The experiments of Friedland and Hastings,138-139 Gunsalus-Miguel et al.,140 and others have shown that the luciferase subunits can be unfolded in urea and refolded in buffer to form active heterodimeric luciferase. However, little has been done to delineate the mechanism of folding and association of the subunits. Using the enzyme from V. fischeri, Hastings and his colleagues demonstrated that the activity of the enzyme was linear with concentration over approximately six orders of magnitude in enzyme concentration.142 At very low concentrations, if subunit dissociation were occurring, one would have expected to see a marked decrease in enzyme activity. None was seen, suggesting that the luciferase subunits do not dissociate at concentrations down to the nmolar range. The only published experiment that addresses the possibility that the luciferase subunits can dissociate under nondenaturing conditions was reported by Anderson et al.143 Two of the mutant luciferases characterized by Cline and Hastings, AK-678-132 and FB-1,133 were incubated together in buffer solutions for prolonged times. The AK-6 luciferase, discussed in detail above, has a defective a subunit and FB-1 has a defective p subunit. With a time course of hours, wild-type luciferase was formed, demonstrating that the subunits of the mutant luciferases can dissociate under nondenaturating conditions and reassociate to form active wild-type enzyme.143 4. Association of Subunits Folded Independently In Vivo Bacterial luciferase subunits are normally translated from a single transcript. However, if the two genes are expressed from separate plasmids in the same cell, active luciferase is formed.144'145 Waddle et al.145 have extended these studies to analyze the folding and association of luciferase subunits in vivo. With the luxA and luxB genes cloned separately on pUC-derived plasmids, one carrying an ampicillin resistance marker and one carrying a kanamycin resistance marker, E. coli cells could be employed to produce only a subunit or only p subunit, or if transformed with both plasmids, E. coli could produce both subunits but translated from separate mRNA molecules. The individual subunits accumulated to high levels in the cells carrying the separate plasmids, and cells carrying both plasmids expressed a high level of bioluminescence when supplied with aldehyde substrate. However, when the lysates of cells carrying a subunit only were mixed with lysates of cells carrying p subunit only, minimal active heterodimeric luciferase was formed, even after prolonged incubation. If the subunits in the lysates were denatured with urea, mixed and allowed to refold by

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FIGURE 17. Proposed mechanism for subunit folding and luciferase assembly. If the subunits are obliged to fold independently in vivo (by expression of only a single gene), they appear to assume stable structures which when mixed together do not interact to form active luciferase unless first unfolded with urea. These observations,145 and more recent results of Ziegler and Baldwin (unpublished) suggest the existence of inactive folding intermediates both preceding and following the dimerization step.

dilution of the urea, a high level of active enzyme was formed, demonstrating that subunits folded individually in vivo were unable to associate to form active enzyme because one or both were in a stable but incorrect conformational state to dimerize. Based on these observations, Waddle et al.145 proposed that the dimerization step in the folding of bacterial luciferase occurs between conformational intermediates on the folding pathway; if dimerization does not occur at the required time, the subunits continue to fold into structures that are stable and unable to associate to form the active enzyme. The model to describe the folding and assembly of the luciferase subunits proposed by Waddle et al. is shown in Figure 17. F. TEMPERATURE SENSITIVE FOLDING MUTANTS Sugihara and Baldwin24 have reported an interesting class of luciferase mutant that produces normal levels of thermally stable luciferase if cells are grown at lower temperatures (about 20 to 25°C) but greatly reduced levels at higher temperatures (about 35°C). The mutants reported comprised a set of deletions having truncated (3 subunit C-termini (Figure 18).24 A variant which changed 10 of the 11 C-terminal residues and lengthened the (3 subunit by one residue, pJS23, was indistinguishable from the wild-type enzyme in folding and dimerization in vivo and in vitro. Deletion of the C-terminal 9 residues and alteration of the sequence of the 10th to 12th residues from the C-terminus of the (3 subunit resulted in a protein, pJS12, which folded normally in vivo and in vitro at 20°C but at 37°C neither folded in vivo to yield active enzyme, nor refolded from urea in vitro. Deletion of 11 residues (pJS2) resulted in a protein which accumulated to very low levels even at 20°C and was unable to refold from urea. All three proteins, pJS23, pJS12 and pJS2, had normal catalytic properties and normal thermal and protease lability. The enzyme from pJS2 appeared to be normal in all respects except that it was incapable of refolding from urea to form active enzyme. Based on these results, Sugihara and Baldwin24 proposed that the C-terminal region of the 3 subunit has little or no effect on the structure or stability of the active form of luciferase, but it does appear to play a crucial role in the efficiency with which the active form of the

FIGURE 18. Deletions from the C-terminus of the p subunit. The construction designated pTB7 is the wild type, pJS23 appears to fold like the wild type, pJS12 is temperature sensitive in folding, and pJS2 is a nonconditional folding mutant.24

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enzyme is achieved. They suggested that the C-terminal region of the (3 subunit might be involved, perhaps in a transient manner, in a specific interaction that is required for the correct folding and assembly of the luciferase enzyme. To further probe this hypothesis, Ke Wei146 generated a series of point mutations in the C-terminal region of the [3 subunit in which he has incorporated proline codons at sites (3313Asp —> Pro, (3317Ala —» Pro, (3320Val —> Pro, and (3323His —> Pro. The only mutation with any impact on the folding and assembly of the luciferase heterodimer was the 3313Asp —» Pro, which behaved the same as the pJS12 mutant described by Sugihara and Baldwin.24 That is, cells carrying the plasmid encoding the (3313 Asp -^ Pro mutation were dark when grown at 37°C, but produced normal levels of luciferase that was thermally stable when grown at 20°C. In addition, the refolding in vitro of the (3313Asp —> Pro luciferase showed the same large temperature dependence as did the luciferase encoded by pJS12.146

IV. BACTERIAL BIOLUMINESCENCE IN VIVO The subject of the physiology of bioluminescent bacteria has been reviewed in detail by other authors within the past 5 years.7'8 The topics of the secondary emitter proteins, the lumazine protein and the yellow fluorescent protein, have been recently reviewed by Lee et al. in a contribution to this series.10 Therefore, an in-depth review will not be undertaken here. Rather, a few of the more recent observations not covered by others will be discussed, and correlations with other topics covered in this review will be attempted. A. ACCESSORY ENZYMES For bioluminescence in vivo, the enzyme luciferase must be supplied with reduced flavin mononucleotide and tetradecanal (see Reference 11 for a recent review). Meighen and his colleagues have made enormous strides in elucidating the enzymology of the aldehyde metabolizing apparatus in bioluminescent bacteria.11 While there are species differences in the physical properties of the enzymes involved in aldehyde biosynthesis, the overall strategy is conserved in all strains of bioluminescent bacteria. There are three genes known to encode proteins required for aldehyde production, the luxC, luxD, and tuxE genes (Figure 6).5 Lesions in luxD yield a phenotype which responds to addition of both exogenous aldehyde and exogenous aliphatic acids, especially tetradecanoic acid.5-147 Lesions in either luxC or luxE result in a phenotype which will emit bioluminescence in the presence of exogenous aliphatic aldehyde but have lost the ability to respond to the aliphatic acid.148 Based initially on this observation but supported by biochemical data from Meighen and his students,11-149151 it has been concluded that the luxC and luxE gene products are necessary and sufficient for the reduction of tetradecanoic acid to tetradecanal. The luxC and luxE gene products, designated the acyl protein synthase and the acyl protein reductase, form a complex, the fatty acid reductase complex, which contains four copies of each protein.152 The LuxD protein apparently participates as a weakly associated member of the complex and probably plays the role of carrying fatty acid to the fatty acid reductase complex.152 The enzymatic source of the supply of reduced flavin mononucleotide in vivo is much less certain. There is no gene encoded by the lux regulon known to be involved in flavin reduction. Furthermore, there are enzymes present in the cytoplasm of E. coli which can supply luciferase with adequate FMNH2 for brilliant luminescence in vivo.4 In fact, the observation of bioluminescence in E. coli4 was a significant surprise in that it had been assumed that the levels of free reduced FMN in E. coli would be exceedingly low, which may indeed be the case. It is possible that in vivo, the enzyme-bound FMN that results from the bioluminescence reaction could be reduced by a disproportionation reaction without dissociation from the enzyme. The source of the reducing equivalents could be reduced flavin bound to another flavoprotein, such as an NAD(P)H:FMN oxidoreductase, or some

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other flavoprotein, or possibly a small molecule metabolite. There are no reports in the current literature of investigations of this critical issue. The fact remains that luciferase in E. coli is supplied with adequate reduced FMN to carry out the bioluminescence reaction without access to the flavin reductase of V. harveyi or V. fischeri.4'5

B. YELLOW FLUORESCENT PROTEIN

Ruby and Nealson153 reported isolation from the ocean of a strain of bioluminescent bacteria that emit yellow light when grown at temperatures of about 18°C, but emit bluegreen light when grown at the higher temperature of 25°C more commonly employed to grow bioluminescent bacteria. Nealson and his students demonstrated that the bacteria, V. fischeri strain Y-1, contain a protein which they designated the yellow fluorescent protein (YFP) which when mixed with luciferase in vitro causes a shift in the emission spectrum from the normal blue-green spectrum to a yellow spectrum centered at about 540 nm.153'154 The yellow fluorescent protein has been purified in two laboratories129-155 and has been conclusively demonstrated to possess noncovalently bound FMN. Daubner et al.129 have shown that the protein is a homodimer of about 22,000 Da subunits with one FMN per subunit. The fluorescence emission spectrum of YFP superimposes on the bioluminescence emission of V. fischeri Y-l grown at 18°C and on the 540 nm emission of in vitro reactions that include YFP.129 The YFP has two effects. First, it causes an acceleration in the rate of the luciferase catalyzed reaction, resulting in an increased intensity of bioluminescence in reactions in vitro.96-97 acetolactate + CO2 Acetolactate + CO2 — > pyruvate Pyruvate + O2 ^» peracetate + CO2 Acetolactate + O2 —> peracetate + pyruvate a b c

(a1"')

pyrb

acec

CO2

02

26

10 —

— 1.2 — 1.2

— 6 — —

— — 0.2 0.2

0.010 0.26 0.23

1000-fold), is attributable to the structural similarity of the analogue to the ene-amine form of the hydroxyethyl-thiamin pyrophosphate

Volume HI

541

reaction intermediate. Binding of thiamin pyrophosphate and its analogues to ALSII is dependent on the presence of a divalent metal, which in the presence of Mg2+ or Mn2+ (but not Co 2+ , Ca 2 + , or Zn2 + ) is a slowly-reversible complex.4 ALSII-FAD, that has been activated by thiamin pyrophosphate and Mg 2 + , can only be inhibited by thiamin pyrophosphate analogues as rapidly as that cofactor dissociates from the enzyme (with a half-time of 5 min).

REFERENCES 1. Stormer, F. CM Isolation of crystalline pH 6 acetolactate-forming enzyme from Aerobacter aerogenes, J. Biol. Chem., 242, 1756, 1967. 2. Stormer, F. C., The pH 6 acetolactate-forming enzyme from Aerobacter aerogenes. II. Evidence that it is not a flavoprotein, J. Biol. Chem., 243, 3740, 1968. 3. Stormer, F. C. and Umbarger, H. E., The requirement for flavin adenine dinucleotide in the formation of acetolactate by Salmonella typhimurium extracts, Biochem. Biophys. Res. Commun., 17, 587, 1964. 4. Schloss, J. V., Van Dyk, D. E., Vasta, J. F., and Kutny, R. M., Purification and properties of Salmonella typhimurium acetolactate synthase isozyme II from E. coli HB101/pDU9, Biochemistry, 24, 4952, 1985. 5. Barak, Z., Calvo, J. M., and Schloss, J. V., Acetolactate synthase isozyme III (Escherichia coli), Methods Enzymol., 166, 455, 1988. 6. Schloss, J. V., Ciskanik, L., Pai, E. F., and Thorpe, C., Acetolactate synthase: a deviant flavoprotein, in Flavins and Flavoproteins, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 907. 7. Poulsen, C. and Stougaard, P., Purification and properties of Saccharomyces cerevisiae acetolactate synthase from recombinant Escherichia coli, Eur. J. Biochem., 185, 433, 1989. 8. Poulsen, C. and Stougaard, P., Solubilization and renaturation of yeast acetolactate synthase expressed in Escherichia coli, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J. V., Eds., VCH, Weinheim, 1990, 269. 9. Durner, J. and Boger, P., Acetolactate synthase from barley Hordeum vulgare L. Purification and partial characterization, Z. Naturforsch. Teil C, 43, 850, 1988. 10. Durner, J. and Boger, P., Oligomeric forms of plant acetolactate synthase depend on flavin adenine dinucleotide, Plant Physiol., 93, 1027, 1990. 1 1 . Gollop, N., Damri, B,, Barak, Z., and Chipman, D. M., Kinetics and mechanism of acetohydroxy acid synthase isozyme III from Escherichia coli, Biochemistry, 28, 6310, 1989. 12. Huseby, N.-E. and Stormer, F. C., The pH 6 acetolactate-forming enzyme from Aerobacter aerogenes. The effect of 2-oxobutyrate upon the enzymic activity, Eur. J. Biochem., 20, 215, 1971. 13. Barak, Z., Kogan, N., Gollop, N., and Chipman, D. M., Importance of AHAS isozymes in branched chain amino acid biosynthesis, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J, V., Eds., VCH, Weinheim, 1990, 91. 14. Wek, R. C., Mauser, C. A., and Hatfield, G. W., The nucleotide sequence of the *7vBN operon of Escherichia coli sequence homologies of the acetohydroxy acid synthase isozymes, Nucleic Acids Res. ,13, 3995, 1985. 15. Squires, C. H., DeFelice, M., Devereux, J., and Calvo, J. M., Molecular structure of i/vIH and its evolutionary relationship to ilvG in Escherichia coli K12, Nucleic Acids Res., 11, 5299, 1983. 16. Lawther, R. P., Calhoun, D. H., Adams, C. W., Hauser, C. A., Gray, J., and Hat field, G. W., Molecular basis of valine resistance in Escherichia coli K-12, Proc. Nad. Acad. Sci. U.S.A., 78, 922, 1981. 17. Schloss, J. V. and Aulabaugh, A., Acetolactate synthase and ketol-acid reductoisomerase: a search for reason and a reason for search, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J, V., Eds., VCH, Weinheim, 1990, 329. 18. Huseby, N.-E., Christensen, T. B., Olsen, B. R., and Stormer, F. C., The pH 6 acetolactate-forming enzyme from Aerobacter aerogenes subunit structure, Eur. J. Biochem., 20, 209, 1971. 19. Falco, S. C., Dumas, K. S., and Livak, K. J., Nucleotide sequence of the yeast i/v2 gene which encodes acetolactate synthase, Nucleic Acids Res., 13, 4011, 1985. 20. Mazur, B. J., Chui, C.-F., and Smith, J. K., Isolation and characterization of plant genes coding for acetolactate synthase, the target enzyme for two classes of herbicides, Plant Physiol., 85, 1110, 1987. 21. Singh, B. K., Newhouse, K. E., Stidham, M. A., and Shaner, D, L., Imidazolinones and acetohydroxyacid synthase from plants, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J, V., Eds., VCH, Weinheim, 1990, 357.

542

Chemistry and Biochemistry of Flavoenzymes

22. Grabau, C. and Cronan, J. E., Jr., Nucleotide sequence and deduced amino acid sequence of Escherichia coll pyruvate oxidase, a lipid activated flavoprotein, Nucleic Acids Res., 14, 5449, 1986. 23. Falco, S. C. and Dumas, K. S., Genetic analysis of mutants of Saccharomyces cerevisiae resistant to the herbicide sulfometuron methyl, Genetics, 109, 21, 1985. 24. Falco, S. C., Selectable markers for yeast transformation, U.S. Patent 4626505, 1986. 25. Mazur, B. J. and Falco, S. C., The development of herbicide resistant crops, Annu. Rev, Plant Physiol. Plant MoL Biol., 40, 441, 1989. 26. Hawkins, C. F., Borges, A., and Perham, R. N., A common structural motif in thiamin pyrophosphatebinding enzymes, FEES Lett., 255, 77, 1989. 27. Abell, L. M., O'Leary, M. H., and Schloss, J. V., Determination of carbon isotope effects and substrate preference for acetolactate synthase by isotope ratio mass spectroscopy, Biochemistry, 24, 3357, 1985. 28. Chipman, D. M., Gollop, N., Damri, B., and Barak, Z., Kinetics and mechanism of acetohydroxyacid synthases, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J. V., Eds., VCH, Weinheim, 1990, 243. 29. Zahler, S. A., Najimudin, N., Kessler, D. S., and Vandeyar, M. A., a-Acetolactate synthesis by Bacillus subtilis, in Biosynthesis of Branched Chain Amino Acids, Barak, Z., Chipman, D. M., and Schloss, J. V., Eds., VCH, Weinheim, 1990, 25. 30. Abell, L. M. and Schloss, J. V., Oxygenase side reactions of Acetolactate synthase and other carbanionforming enzymes, Biochemistry, 30, 7883, 1991. 31. Becker, W., Benthin, U., Eschenhof, E., and Pfeil, E., Zur Kenntnis der Cyanhydrinsynthese II. Reindarstellung und Eigenschaften der Oxynitrilase aus bittern Mandeln (Prunnus communis Stokes), Biochem. Z., 337, 156, 1963. 32. Jorns, M. S., Mechanism of catalysis by the flavoenzyme oxynitrilase, J. Biol. Chem., 254, 12145, 1979. 33. Morell, H., Clark, M. J., Knowles, P. F., and Sprinson, D. B., The enzymic synthesis of chorismic and prephenic acids from 3-enolpyruvylshikimic acid 5-phosphate, /. Biol. Chem., 242, 82, 1967. 34. Welch, G. R., Cole, K. W., and Gaertner, F. H., Chorismate synthase of Neurospora crassa, a flavoprotein, Arch. Biochem. Biophys., 165, 505, 1974. 35. Hasan, N. and Nester, E. W., Purification and properties of chorismate synthase from Bacillus subtilis, J. Biol. Chem., 253, 4993, 1978. 36. Gupta, N. K. and Vennesland, B., Glyoxylate carboligase of Escherichia coli: a flavoprotein, J. Biol. Chem., 239, 3787, 1964. 37. Gupta, N. K. and Vennesland, B., Glyoxylate carboligase of Escherichia coli: some properties of the enzyme, Arch. Biochem. Biophys., 113, 255, 1966. 38. Chung, S.-T., Tan, R. T. Y., and Suzuki, L, Glyoxylate carboligase of Pseudomonas oxalaticus. A possible structural role for flavine-adenine dinucleotide, Biochemistry, 10, 1205, 1971. 39. Cromartie, T. H. and Walsh, C. T., Escherichia coli glyoxalate carbo-ligase, properties and reconstitution with 5-deaza-FAD and 1,5-dihydro-deaza-FADH, /. Biol. Chem., 251, 329, 1976. 40. Koland, J. G., Miller, M. J., and Gennis, R. B., Reconstitution of the membrane bound ubiquinone dependent pyruvate oxidase respiratory chain of Escherichia coli with cytochrome D terminal oxidase, Biochemistry, 23, 445, 1984. 41. Chang, Y.-Y. and Cronan, J. E., Jr., Common ancestry of Escherichia coli pyruvate oxidase and the acetohydroxy acid synthases of the branched-chain amino acid biosynthetic pathway, J. BacterioL, 170, 3937, 1988. 42. Schloss, J. V., Ciskanik, L. M., and Van Dyk, D. E., Origin of the herbicide binding site of acetolactate synthase, Nature, 331, 360, 1988. 43. LaRossa, R. A. and Smulski, D. R., ilvB-encoded acetolactate synthase is resistant to the herbicide sulfometuron methyl, J. BacterioL, 160, 391, 1984.

Volume III

543

Chapter 18

PHTHALATE DIOXYGENASE REDUCTASE AND RELATED FLAVIN-IRON-SULFUR CONTAINING ELECTRON TRANSFERASES Christopher J. Batie, David P. Ballou, and Carl C. Correll

TABLE OF CONTENTS I.

Introduction

544

II.

Biological Role of Phthalate Dioxygenase Reductase

544

III.

Electron Transfer Chains for [2Fe-2S]-Fe(II) Oxygenases

545

IV.

Structural Properties of PDR

548

V.

Spectroscopic Properties of PDR

548

VI.

Electron Transfer Reactions

551

VII.

Kinetics

552

VIII. Other Observations on Related Systems

553

Acknowledgments

554

References

554

544

Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION Numerous organisms found in the soil express multicomponent oxygenase systems that use NADH and O2 to attack unactivated aromatic compounds.' These dioxygenase or monooxygenase reactions initiate the degradation of benzene rings. Scheme 1 shows the functional units found in most types of monooxygenase and dioxygenase systems (with the exception of the flavoprotein hydroxylases). Oxido-reductase systems mediate electron transfer between NADH and the oxygenase. The active sites of the oxygenase systems often contain iron, either in heme or in some other type of ligation. Since NADH must transmit both of its electrons simultaneously as a hydride, whereas iron is generally restricted to single-electron reactions, a suitable reductase is required that can mediate this transition. Thus electron transfer chains generally contain both flavin and iron-sulfur (Fe/S) redox components. Flavins are distinctive among biological coenzymes in that they can accept two electrons (as hydrides) from NADH and transmit these electrons one at a time (via the semiquinone redox state) to other electron transfer components. Fe/S Redox centers are well suited for transfer of electrons of moderately low potential (- 100 to -400 mV), a process common to many oxygenases as well as to the electron transport chains of the respiratory systems. This review will be restricted to the reductase systems of Rieske center non-heme iron-containing oxygenases, although the principles presented are also generally valid of the reductases for hemoproteins, other oxygenases, and electron transport chains. We will use phthalate dioxygenase reductase (PDR) as a focus for describing this family of reductases. PDR is a small protein (34 kDa)2 containing both a FMN and a plant-type ferredoxin (Fd) [2Fe-2S] center and serves to transmit electrons from NADH to its oxygenase. Structural, kinetic, and mechanistic data for PDR are more complete than for any of the other reductases of this class.2"6 Furthermore, since both redox centers are on the same small protein, association-dissociation reactions will not complicate electron transfer studies.6 The protein has been crystallized,3 partially sequenced (unpublished), and X-ray structural work is in progress.4 Cloning is also well underway to further enable detailed structural and mechanistic studies. Moreover, in contrast to many of the other systems of this class, PDR is reasonably stable and can be obtained in 100 mg quantities.

II. BIOLOGICAL ROLE OF PHTHALATE DIOXYGENASE REDUCTASE Phthalate dioxygenase reductase is isolated from Pseudomonas cepacia DB01 cells that are grown with phthalate as the sole carbon and energy source.2-5 PDR transfers electrons from NADH to phthalate dioxygenase (PDO) during the oxygenation reaction shown in Scheme 2, which results in formation of 4,5-dihydro-dihydroxyphthalate. The phthalate dioxygenase system thus carries out the first step in the metabolism of phthalate by this bacterial strain. The next metabolic reaction (not shown) is a rearomatization catalyzed by a diol dehydrogenase to regenerate the NADH and produce the corresponding catechol. Thus via these two steps a stable aromatic compound is converted to its much less stable catechol with the net consumption of one O2. This process is widely used by pseudomonads and other soil bacteria and provides an efficient mechanism of activating aromatic rings for further metabolism.1'5'7-8 The biological redox components used for these transformations are all quite similar and will be described below. Phthalate dioxygenase has been purified and found to be a tetramer of identical 48 kDasubunits, each of which contains two distinct types of Fe centers: a Rieske-type [2Fe-2S] center and a mononuclear Fe(II) center.2'5'9"n Similar pairs of centers have also been found in other oxygenase systems that catalyze formation of m-dihydrodiols including: benzoate dioxygenase,12'15 benzene dioxygenase,16"20 naphthalene dioxygenase,21"25 toluene dioxygen-

Volume III

SCHEME 1.

SCHEME 2.

545

Functional units of oxygenase systems.

The reaction catalyzed by the PDR/PDO system.

ase,8'2632 and pyrazon dioxygenase.33 This comprises a distinct class of oxygenases, the Rieske-type [2Fe-2S] center-Fe(II)-containing oxygenases (abbreviated, (Fe/S)Rieske-Fe(II)). Biphenyl dioxygenase, which has been cloned and partially sequenced,34'35 catalyzes a similar reaction and may be another member of this class, but the enzyme has not yet been purified. The (Fe/S)Rieske-Fe(II) class also includes oxygenases that do not yield dihydrodiol products. For example, 4-chlorophenylacetate 3,4-dioxygenase has a Rieske-type [2Fe-2S] center and a requirement for Fe(II). This enzyme converts its substrate to a catechol, 4,5-dihydroxyphenylacetate, eliminating Cl~ in the process; this reaction may involve formation of a nascent ds-dihydrodiol, which rearomatizes by elimination of HC1.36-37 Similarly, the conversion of 4-sulfobenzoic acid to protocatechuic acid by an NADH requiring dioxygenase may proceed by a similar reaction; both atoms of oxygen in the product come from O2.38 In addition to these dioxygenases, a monooxygenase, 4-methoxybenzoate demethylase, has been shown to be a member of this class.39 47 Vanillate demethylase, which has been cloned and sequenced, may also be similar.48-49 Vanillate is an intermediate in the degradation of lignin. l These data indicate the important roles that the (Fe/S)Rieske-Fe(II) class of oxygenases play in the aerobic biodegradation of unreactive aromatic compounds. We and our co-workers have recently been able to determine the structure of the Riesketype [2Fe-2S] center of phthalate dioxygenase.2'9-11 EPR, ENDOR and EXAFS spectroscopic analysis of phthalate dioxygenase has shown that this center has two N-Fe bonds. As shown in Figure 1, both N ligands are histidyl residues10 and are coordinated to the same Fe through the 8-N of each histidine (data unpublished). The geometry and interatomic distances of the center are very similar to those of plant-Fd-type [2Fe-2S] centers,9"1! which have two cysteinyl thiolates coordinated to each of the two irons, Fe2S2Cys4.

III. ELECTRON TRANSFER CHAINS FOR [2FE-2S]-FE(II) OXYGENASES All of the [2Fe-2S]-Fe(II) type of oxygenases require NADH and an electron transfer system. Although the oxygenases are quite similar, the electron transfer systems are widely divergent (Table 1). These electron transfer chains have one flavin and one or two Fe/S centers. However, there is great variety in (1) the location of the Fe/S center(s) — on the flavoprotein or on a separate protein; (2) the number of Fe/S centers — one or two; and (3) the type of iron sulfur center(s) — Rieske-type or plant-Fd-type [2Fe-2S].

546

Chemistry and Biochemistry of Flavoenzymes

FIGURE 1. A drawing of the proposed coordination of the Rieske [2Fe2S] metal cluster.

The simplest electron transfer chains (Class I) have one flavin and one plant-Fd-type [2Fe-2S] center on a single relatively small protein. In addition to the phthalate dioxygenase system, which has PDR2, four other enzymatic systems have been shown to have a oneprotein electron transfer chain: benzoate oxygenase,13 putidamonooxin (4-methoxy-benzoate demethylase),44'45 4-sulfobenzoate dioxygenase,38'53 and 4-chlorophenylacetate-3,4-dioxygenase (component B).37 Four of these systems utilize FMN, but benzoate dioxygenase reductase uses FAD as its flavin. More complex are the two protein-two redox center electron transfer chains (Class II). Examples of systems using these are benzene dioxygenase,16'20 toluene dioxygenase,28'29 and pyrazon dioxygenase.33 These systems all consist of an FADcontaining flavoprotein and a [2Fe-2S] protein. The pyrazon dioxygenase system contains an Fe/S protein with a center like that of spinach Fd — Fe2S2Cys4; in contrast, benzene and toluene dioxygenases utilize an Fe/S protein with a Rieske-type Fe/S center — Fe2S2Cys2His2 (see Figure 1). Most complex is the naphthalene dioxygenase system (Class III). It has an oxido-reductase with both a flavin and a plant-Fd-type [2Fe-2S] center, and also requires another electron carrier, ferredoxinNAP, which has a Rieske-type [2Fe-2S] center.21"25 The genes for the reductase components of the toluene 2,3-dioxygenase system,31*32 as well as for the benzene dioxygenase systems,51 have been sequenced and the protein sequence for the ferredoxin component of a different benzene dioxygenase system has also been published.50 The gene sequences of the toluene and benzene systems are nearly identical (>99%).32 The naphthalene dioxygenase system has also been cloned and sequenced.25'54 Kurkela et al.25 found three naphthalene dioxygenase genes (ndoA, ndoB, and ndoC), again with significant sequence similarity to those of the benzene dioxygenase51 (pl,/?2, and/?3). Comparing the benzene and naphthalene systems, ndoA corresponds to/?3, ndoB topi, and ndoC to p2 (pi and p2 are the subunits of the dioxygenase). Gene p3 has been shown to code for the Fd component of benzene dioxygenase,50 which suggests that ndoA codes for FdNAP. However, the naphthalene dioxygenase system isolated by Ensley, Haigler and Gibson22"24 has 4 polypeptides and 36 kDa more total protein. Kurkela et al.54 did not sequence the reductase, which accounts for the missing 36 kDa. The vanillate demethylase genes have been sequenced48 and also fit the pattern common to Class I. The amino acid sequence of PDR is nearly complete (unpublished data); it is quite similar to that of the vanillate

Volume III

547

TABLE 1 Electron Transfer Chains for Rieske-Type [2Fe-2S] Oxygenases1

Class

Number of Proteins

NADH Oxido-reductase Chromophores

References

Fe/S Protein Chromophore

Type

Size (kDa)

Stable Semiquinone

Type

Size (kDa)

IA

Phthalate P. cepacia DB01

1

FMN + [Fe2S2Cys4]

34 (S)

Yes

None

2,3,6

4-Methoxybenzoate P.putida (DSM-1868)

1

FMN + [Fe2S2Cys4]

42

Yes

None

43-46

p-Sulfobenzoate C. testosterom 1-2

1

FMN + [Fe2S2Cys4]

35

None

38

4-Chlorophenylacetate P.sp. strain CBS 3

1

FMN + [Fe2S2Cys4]

35

Yes

None

36,37

1

FAD + [Fe2S2Cys4]

38 (S)

N.D.2

None

13,15

IB

Benzoate P.arvitlaC-l (ATCC 23974) A. calcoaceticus BD413

55

HA

Pyrazon

2

FAD

67

N.D.2

[Fe2S2Cys4]

12

33

Benzene P. putida ML2 (NCIB 12190)

2

FAD

81 (S)

N.D.2

[Fe2S2Cys2 His2]

12 (S)

16-20,50,51

Toluene P. putida Fl

2

FAD

42.9 (S)

N.D.2

[Fe2S2Cys2 His2]

11.9(S)

8,26-32

2

FAD + [Fe2S2Cys4]

36.3 (S)

[Fe2S2Cys2 His2]

13.6 (S)

21-25, 52

IIB

III

Naphthalene P. sp. (NCIB 9816)

1. In addition to the electron transfer chains, each of these systems has an oxygenase consisting of one or two types of subunits with one of the subunits containing both a Rieske [2Fe-2S] center and a mononuclear Fe2+ center. The latter is believed to be the site of oxygenation. 2. N.D.; not determined.

548

Chemistry and Biochemistry of Flavoenzymes

demethylase oxido-reductase (vanE). The benzoate system from Acinetobacter calcoaceticus has been sequenced55 and shows considerable homology. Little is known about the sequence of members of the other structural classes of oxidoreductases listed in Table 1. It will be interesting to see whether any of the other classes will show as much homology as Class II and Class III. The variety of electron transport chains that serve very similar oxygenases raises an interesting question. Have different combinations of components been selected for their particular properties, or is the incorporation of a particular set of electron transfer proteins into a new catabolic pathway an evolutionary accident? PDO shows considerable specificity towards its electron donor. Although a reduced PDO on its own can effect slow oxygenation, a significant rate of oxygenation by PDO occurs only in the presence of PDR; other oxidoreductase systems do not stimulate activity (including benzoate dioxygenase reductases from two different pseudomonads, liver microsomal P-450 reductase and spinach ferredoxin reductase with ferredoxin). Thus some specific interaction of PDR with PDO is crucial to catalysis. Other systems reported to show specificity for their particular reductases are naphthalene dioxygenase,24 toluene dioxygenase,28'29 4-methoxybenzoate monooxygenase,46 and benzene dioxygenase.46

IV. STRUCTURAL PROPERTIES OF PDR Crystallographic studies of PDR,3>4'56 now at a resolution of 2.0 A with an R = 0.19, show that the chain folds into three domains, each associated with one of the cofactors. In PDR the FMN domain is at the N-terminus of the chain, and the [2Fe-2S] domain is at the C-terminus. The domains are arranged to bring the FMN and [2Fe-2S] prosthetic groups together near the center of the molecule (Figure 2A), with the flavin adjoining the binding site for pyridine nucleotide. The binding site for pyridine nucleotides has been identified from experiments in which NAD, NADH (unpublished), AAD, and TNAD58 have been added to crystals of PDR.4 As one might anticipate from the functional similarity between PDR and the ferredoxin reductase system (FNR/Fd), the structures of the FMN and NAD domains of these molecules are closely related,59 with equivalent topologies. The fold of the FMN domain, a six-stranded antiparallel (3-barrel with a helix at one end, is a new motif for flavin binding domains, found so far only in FNR59 and PDR. The NAD domain resembles the classic a/0 nucleotide binding fold;60 it consists of a five-stranded parallel sheet surrounded by four helices. The [2Fe-2S] domain includes a four-stranded mixed p-sheet and two helices in a fold that resembles the [2Fe-2S] ferredoxin from Anabaena.61 The orientation of the FMN and [2Fe-2S] centers is shown in Figure 2B. The electron density at 2.0 A and the flavin-protein contacts confirm that the re-side of the flavin ring faces the NAD domain. This orientation is consistent with the stereochemistry of hydride transfer observed in related proteins.62 The dimethylbenzene end of the flavin is situated in the cleft between the NAD and FMN domains and points toward the [2Fe-2S] center. The folding of the polypeptide around the [2Fe-2S] center is similar to the other known plant type [2Fe-2S] ferredoxins.61*63

V. SPECTROSCOPIC PROPERTIES OF PDR As purified, PDR is an orange protein having the absorbance spectra shown in Figure 3 (solid line). Peaks are found at 273, 280 (not shown), 350, and 462 nm; in addition, the spectrum of oxidized PDR has shoulders at 410, 430, 488, 560 nm, and a broad longwavelength band from 640 to 720 nm. This spectrum is consistent with the sum of absorbance from a flavin and a plant-Fd-type [2Fe-2S] center; chemical analysis indicates 1 FMN, 2

Volume III

FIGURE 2. (A) A ribbon drawing produced from the program PAP57 of PDR with the FMN and [2Fe-2S] center included, and with the N and C-termini labeled. The regions labeled 1 and 2 are not well defined and may be disordered. Region 1 is the linker between the NAD and [2Fe-2S] domains and region 2 is the second helix in the [2Fe-2S] domain. (B) A closeup view, rotated slightly from 1(A), of the FMN and the [2Fe-2S] center. The Ca chain trace is drawn with bold lines and the FMN, [2Fe-2S], and cysteine side chains are represented with thin lines. The direction of the Ca chain trace of the large loop is from left to right, whereas the lower segment direction is from right to left. The distance between the C8-methyl of the FMN (C8M) and the nearest cysteine sulfur ligand (SG) is 5.0 A, the distance between the C8-methyl and Pel is 7.3 A, and the distance between Fel and Fe2 is 2.7 A.

549

550

Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. Absorption spectra of PDR. Oxidized PDR (14.5 fiM) was mixed with an equal volume of buffer to give the spectrum of the oxidized protein ( ). Two electron reduced PDR (—) was recorded 3 to 10 seconds after mixing excess NADH anaerobically with oxidized PDR. Fully reduced PDR (—-—) was recorded 20 min later. Spectra were recorded in a stopped flow spectrophotometer with a 2 cm optical path. Conditions: 4°C, 0.1 M HEPES, pH 8.0. (From Batie, C. J., and Ballou, D. P., Electron transfer kinetics of phthalate oxygenase reductase, an iron-sulfur containing flavoprotein, in Flavins and Flavoproteins 1987, Edmonson, D. E. and McCormick, D. B., Eds. Copyright © 1987 Walter de Gruyter & Co., Berlin with permission.)

Fe, and 2 S 2 ~ per polypeptide.2 The relative amplitude of the shoulders at 410 and 430 nm varies with pH; absorbance increases at 410 nm at higher pH. This effect may be due to an ionization linked to the [2Fe-2S] center, as the ferredoxins generally have a peak in this region.64 The other members of Class I have similar absorbance spectra.13-37'39'44-45 Thermodynamic stabilization of the neutral flavin radical appears to be a general characteristic of enzymes of Class I; addition of one equivalent of NADH results in reduction of the Fe/S center and the appearance of a neutral radical.2-6'37'39'44-45 Phthalate dioxygenase reductase can be reduced by either NADH or dithionite. In titrations at pH 6.8 a set of absorbance changes is observed throughout addition of the first two electron equivalents; these include an increase in absorbance between 520 and 675 nm, and a decrease in absorbance between 340 and 520 nm (Figure 3). The first two electrons are distributed nearly equally between the flavin and the [2Fe-2S] center, almost quantitatively generating the neutral semiquinone and the reduced Fe/S center ([2Fe-2S]1 + ). EPR spectra of two-electron reduced enzyme corroborate this observation, showing both flavin semiquinone and reduced Fe/S signals (Figure 4). A third electron equivalent can be added to the enzyme (from dithionite or in a slow process from NADH), thus reducing the semiquinone to fully reduced flavin, but not affecting the EPR signal of the reduced Fe/S center. Similar observations have been reported for the reductase components of 4-methoxybenzoate demethylase39'44-45 and 4-Cl-phenylacetate dioxygenase.37 The data from titrations of PDR with NADH allow us to place lower limits on the redox potentials of the flavin and the [2Fe-2S] center. At pH 6.8, the Em values of the FMN/ FMNH • (El) and the [2Fe-2S]2+/[2Fe-2S]1+ couples are nearly equal, and at least 60 mV more positive than the Em of the FMNH/FMNH2 (E2) couple (i.e., nearly complete formation

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FIGURE 4. EPR spectrum of PDR. Spectrum was recorded after anaerobically reducing PDR (96 juiAf) with 1 equivalent of NADH. The spectrum was recorded at 20°K, with 10 G modulation at 0.1 mW power at 9.224 GHz.

of the semiquinone is observed). The latter redox couple, in turn, is at least 50 mV more positive than the NAD/NADH couple (- 320 mV). Hence, the Em for the first two electrons into PDR must be greater than -210 mV, which is quite high for [2Fe-2S] ferredoxins. The redox titrations show a pH dependence indicating that the two-electron reduced form of the FMN is largely in the anionic state. This conclusion rests on the following observations. When the protein is reduced by two electrons at pH 6.8, most of the flavin appears as the neutral semiquinone; thus Ej > E2. With increasing pH less semiquinone is present; thus E^ which decreases by 60 mV/pH, approaches E2. For that to happen, the flavin must bind less than one H + when the reduced flavin is formed from FMNH* (i.e., above pH 6.8 the E2 decreases < 60 mV/pH); thus the reduced flavin is largely anionic. These data may be compared with the two-electron redox potentials of the reductase of the 4-methoxy-benzoate demethylase (-200 to -240 mV43), and of the Fds of benzene dioxygenase (—155 mV19) and toluene dioxygenase ( — 109 mV29). We have used EPR spectra to investigate possible interactions between the two centers of PDR (Figure 4). The X-ray data indicates that the point of closest approach from one of the sulfur ligands of the Fe/S center to the 8-methyl group of the flavin is about 5 A. However, the shape of the paramagnetic signals of the FMNH- and [2Fe-2S]1+ centers appears to be independent of the redox state of the other center. This would indicate that the two centers are separated by at least 10 A.65 This dilemma needs to be addressed. Very similar EPR observations have been reported on the reductase moiety of the 4-chlorophenylacetate dioxygenase system.37 In this case the X-ray structure is not available.

VI. ELECTRON TRANSFER REACTIONS Phthalate dioxygenase reductase catalyzes electron transfer from NADH to PDO. PDR is specific for NADH; NADPH yields no detectable activity. Isotopic labeling studies show that PDR removes the pro-R hydrogen from NADH and a deuterium isotope effect on the hydride transfer from NADH to PDR of 3.0 ± .3 has been observed (data unpublished). PDR will reduce a variety of electron acceptors other than PDO, including: cytochrome c, DCIP, ferricyanide, FMN, and nitro-blue tetrazolium. The kcat with several of these substrates was found to be similar to that observed with PDO (50 to 100 s"1). This suggests that electron transfer from PDR is not rate-limiting in these cases, but instead, the overall rate

552

Chemistry and Biochemistry of Flavoenzymes

SCHEME 3.

Steps in the reduction of PDR.

of electron transfer is limited by some step or steps in the reduction of PDR by NADH. This also fits with the observation of the isotope effect. Curiously, PDR does not reduce oxygen very well (0.05 s"1); this raises the question of how an apparently indiscriminate oxido-reductase selectively avoids reducing O2. The other members of this class are relatively similar to PDR in their catalytic characteristics. As a rule, they are specific for NADH and will reduce cytochrome c, ferricyanide, and other dyes; the optimal activity is generally observed near pH g.o.13'15-16'23'28-37'43

VII. KINETICS We have examined the kinetics of electron transfer by phthalate dioxygenase reductase. This is the only member of this class of oxido-reductases to have been extensively studied by kinetic analysis. Rapid reaction studies, combined with steady state kinetic analysis, indicate that electron transfer by PDR from NADH to an electron acceptor is a multistep process, with no one step being rate-limiting. Observation of the reduction of PDR by NADH in a stopped flow spectrophotometer has allowed us to determine the mechanism outlined in Scheme 3. We first observe an intermediate with spectral characteristics of an FMN-NADH chargetransfer complex (CT1). Saturation-type kinetics in the formation of this species indicates a preceding PDR-NADH complex (MC). Hydride transfer then occurs; we observe not the expected FMNH2-[2Fe-2S]2+ species, but instead, an FMNH--[2Fe-2S]1 + species (CT2). Thus electron transfer from reduced flavin to [2Fe-2S] is much faster than reduction of the flavin. The initially observed FMNH--[2Fe-2S]1+ species has a long-wavelength band with absorbance extending beyond 800 nm. This species results from a reversibly formed chargetransfer interaction between the flavin semiquinone and NAD; as such it is a novel species, not previously observed with other flavoproteins. NAD then dissociates from CT2, yielding either free PDR in the FMNH--[2Fe-2S]1+ state or (with excess NADH) an NADH-bound species. Simulations of the data from the reactions of PDR with either NADH or 4R-NADD can fit the model of Scheme 3. The data were collected at 4°C, pH 8.0. These simulations indicate that the Kd of the initially formed PDR-NADH complex (Reaction 1) is about 50 |xM. Formation of CT1 (Reaction 2) and hydride transfer from NADH (Reaction 3) occur

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at rates of about 80 to 90 s" 1 and 45 s" 1 , respectively. Internal electron-transfer from FMNH 2 to [2Fe-2S]2+ (Reaction 4) must occur at a rate >200 s" 1 . Dissociation of NAD from CT2 (Reaction 5) occurs at a rate of 40 to 50 s"1; titrations indicate a Kd of 2 to 5 mAf for NAD. We conclude from the simulations that a combination of steps B, C, and E limit the overall rate of electron transfer. The rates of these steps are sufficiently rapid to account for turnover at this temperature. This conclusion is also compatible with the observed steady-state kinetic isotope effect of 3.

VIII. OTHER OBSERVATIONS ON RELATED SYSTEMS The oxido-reductases that have been purified have several properties in common. The oxido-reductase of the 4-methoxybenzoate monooxygenase system is very similar in structure to PDR, having FMN and a plant-Fd-type [2Fe-2S] center on a peptide of 42 kDa. In addition, in the partially reduced state both a neutral-semiquinone and a reduced [2Fe-2S] center appear.43 Unlike phthalate dioxygenase reductase, however, NADH-putidamonooxin oxido-reductase is reported to be unstable in aerobic solution. Component B of 4-chlorophenylacetate 3,4-dioxygenase also has FMN and a Fd-type [2Fe-2S] center; when reduced by excess NADH both centers are observed in the one-electron reduced state. The NADHbenzoate dioxygenase oxido-reductase also resembles the preceding enzymes, but has FAD as the flavin moiety. Yamaguchi and Fujisawa15 have reported removal and reconstitution of the Fe/S cluster of benzoate dioxygenase reductase. This enzyme is reported to be unstable in the absence of thiol reducing agents, but with dithiothreitol has stability comparable to phthalate dioxygenase reductase.13 Class II enzymes have a flavoprotein and a separate ferredoxin that has a Rieske-type [2Fe-2S] center. The benzene dioxygenase electron transfer chain has been best characterized physically. The flavoprotein has FAD bound to an 81 kDa polypeptide and catalyzes electron transfer from NADH to the 12 kDa ferredoxin.19 The latter component is required for reduction of either the oxygenase or cytochrome c. EPR, Mossbauer and UV-visible absorbance spectra are consistent with the iron-sulfur center in the 12 kDa ferredoxin being a Rieske-type structure.16'19 The midpoint redox potential at pH 7.0 is - 155 mV. 19 The toluene dioxygenase electron transfer chain is the other well-characterized Class II system. The flavoprotein of the system is a 46 kDa protein that contains FAD. Most FAD dissociates from the ferredoxinTOL reductase during purification, but the enzyme can be reconstituted; the KM for FAD added to assays was 2.5 nM.28 Upon addition of excess NADH, long-wavelength absorbance was observed; this broad absorbance band is probably due to charge-transfer interactions between reduced flavin and NAD.28 FerredoxinTOL has also been purified. Its characteristics are similar to the ferredoxin of the benzene dioxygenase system: Mr = 11,90032 (SDS-PAGE experiments imply Mr = 15,300),29 there are 2 equivalents each of Fe and S 2 ~; optical spectra are indicative of a Rieske-type [2Fe-2S] center, and the redox potential at pH 7 is -109 mV.29 Naphthalene dioxygenase is unusual in having a two protein electron transfer chain that contains one flavin and two Fe/S centers. 2125 ReductaseNAP, as isolated, has a plant-Fd-type [2Fe-2S] center; addition of flavin is required for restoration of activity. Curiously, either FMN or FAD can reconstitute activity.23 ReductaseNAP transfers electrons from reduced pyridine nucleotides to ferredoxinNAP, which has a Rieske-type [2Fe-2S] center.23-24 There is much variability in the overall structure of the electron transport chains of the Fe/S-Fe(II)-type of oxygenases; however, gene sequencing has indicated similarities in the structure of peptides associated with individual redox centers. Two genes required for vanillate degradation in a pseudomonad have been sequenced. One, vanB, is similar to many members of the ferredoxin class of proteins;48 the gene product of vanB is 33 kDa in size, suggesting that it may be similar to PDR and related enzymes. Indeed, a comparison of the

554

Chemistry and Biochemistry of Flavoenzymes

translated gene sequence of vanE with the protein sequence of PDR shows considerable homology. The genes for the toluene,32 benzene,51 and naphthalene25 dioxygenase systems have also been cloned and sequenced and found to be quite similar.

ACKNOWLEDGMENTS Biochemical studies of PDR and PDO have been supported by NIH Grant GM 20877, and structural studies of PDR have been supported by NIH GM 16429. We would like to especially acknowledge Dr. Martha L. Ludwig for her support in structural studies of PDR and discussions of the manuscript. We would also like to acknowledge Dr. W. R. Dunham for the EPR spectrum and Dr. Gerben Zylstra and George Gassner for discussions of this manuscript.

REFERENCES 1. Dagley, S., Biochemistry of aromatic hydrocarbon degradation in Pseudomonads, in The Bacteria, Vol. X, Gunsalus, I. C., Sokatch, J. R., and Ornston, L. N., Eds., Academic Press, Orlando, 1986, 527. 2. Batie, C. J., LaHaie, E., and Ballou, D. P., Purification and characterization of phthalate oxygenase and phthalate oxygenase reductase from Pseudomonas cepacia, J. Biol. Chem., 262, 1510, 1987. 3. Correll, C. C., Batie, C. J., Ballou, D. P., and Ludwig, M. L., Crystallographic characterization of phthalate oxygenase reductase, an iron-sulfur flavoprotein from Pseudomonas cepacia, J. Biol. Chem., 260, 14633, 1985. 4. Correll, C. C. and Ludwig, M. L., Structure determination of an iron-sulfur flavoprotein, in Flavins and Flavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 743. 5. Ballou, D. P. and Batie, C., Phthalate oxygenase, a Rieske iron-sulfur protein, in Oxidases and Related RedoxSystems, King, T. E., Mason, H. S., and Morrison, M., Eds., Alan R. Liss, New York, 1988, 211. 6. Batie, C. J. and Ballou, D. P., Electron transfer kinetics of phthalate oxygenase reductase, an iron-sulfur containing flavoprotein, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 377. 7. Ziffer, H., Kabuto, K., Gibson, D. T., Kobal, V. M., and Jerina, D. M., The absolute stereochemistry of several m-dihydrodiols microbially produced from substituted benzenes, Tetrahedron, 33, 2491, 1977. 8. Gibson, D. T., Yen, W. K., Liu, T. N., and Subramanian, V., Toluene dioxygenase: a multienzyme system from Pseudomonas putida, in Oxygenases and Oxygen Metabolism, Nozaki, M., Yamamoto, S., Ishimura, Y., Coon, M. J., Ernster, L., and Estabrook, R. W., Eds., Academic Press, New York, 1982, 211. 9. Cline, J. F., Hoffman, B. M., Mims, W. B., Lahaie, E., Ballou, D. P., and Fee, J. A., Evidence for N coordination of Fe in the [2Fe-2S] clusters of Thermus Rieske protein and phthalate dioxygenase, /. Biol. Chem., 260, 3251, 1985. 10. Gurbiel, R. J., Batie, C. J., Sivaraja, M., True, A. E., Fee, J. A., Hoffmann, B. M., and Ballou, D. P., Electron-nuclear double resonance spectroscopy of 15N-enriched phthalate dioxygenase from Pseudomonas cepacia proves that two histidines are coordinated to the [2Fe-2S] Rieske-type clusters, Biochemistry, 28, 4861, 1989. 11. Tsang, H. T., Batie, C. J., Ballou, D. P., and Penner-Hahn, J. E., X-ray absorption spectroscopy of the [2Fe-2S] Rieske cluster in Pseudomonas cepacia phthalate dioxygenase. Determination of core dimensions and iron ligations, Biochemistry, 28, 7233, 1989. 12. Yamaguchi, M., Yamaguchi, T., and Fujisawa, H., Studies on mechanism of double hydroxylation. I. Evidence for participation of NADH-cytochrome c reductase in the reaction of benzoate 1,2-dioxygenase (benzoate hydroxylase), Biochem. Biophys. Res. Commun., 67, 264, 1975. 13. Yamaguchi, M. and Fujisawa, H., Characterization of NADH-cytochrome c reductase, a component of benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-l, /. Biol. Chem., 253, 8848, 1978. 14. Yamaguchi, M. and Fujisawa, H., Purification and characterization of an oxygenase component in benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-l, J. Biol. Chem., 255, 5058, 1980. 15. Yamaguchi, M. and Fujisawa, H., Reconstitution of iron-sulfur cluster of NADH-cytochrome c reductase, a component of benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-l, J. Biol. Chem., 256, 6783, 1981.

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16. Axcell, B. C. and Geary, P. J., Purification of some properties of a soluble benzene-oxidizing system from Pseudomonas putida, Biochem. J., 146, 173, 1975. 17. Crutcher, S. E. and Geary, P. J., Properties of the iron-sulfur proteins of the benzene dioxygenase system from Pseudomonas putida, Biochem, J,, 177, 393, 1979. 18. Geary, P. J. and Dickson, D. P. E., Mossbauer spectroscopy studies of the terminal dioxygenase from Pseudomonas putida by electron-spin-resonance spectroscopy, Biochem, J., 195, 199, 1981. 19. Geary, P. J., Saboo walla, F., Patil, D., and Cam mack, R., An investigation of the iron-sulfur proteins of benzene dioxygenase from Pseudomonas putida by electron-spin-resonance spectroscopy, Biochem. J., 217, 667, 1984. 20. Zamanian, M. and Mason, J. R., Benzene dioxygenases in Pseudomonas putida; subunit composition and immuno-cross-reactivity with other aromatic dioxygenases, Biochem. J,, 244, 611, 1987. 21 Jeffrey, A. M., Yeh, H. J. C., Jerina, D. M., Patel, T. F., Davey, J. R., and Gibson, D. T., Initial reaction in the oxidation of naphthalene by Pseudomonas putida, Biochemistry, 14, 575, 1975. 22. Ensley, B. D. and Gibson, D. T., Naphthalene dioxygenase: purification and properties of a terminal oxygenase component, J. BacterioL, 155, 505, 1983. 23. Haigler, B. E. and Gibson, D. T., Purification and properties of NADH-ferredoxinNAP reductase, a component of naphthalene dioxygenase from Pseudomonas sp. strain NCIB 9816, J. BacterioL, 172, 457, 1990. 24. Haigler, B. E. and Gibson, D. T., Notes: purification and properties of ferredoxinNAP, a component of naphthalene dioxygenase from Pseudomonas sp. strain NCIB 9816, J. BacterioL, 172, 465, 1990. 25. Kurkela, S., Lehvaslaiho, H., Palva, E. T., and Reeri, T. H., Cloning, nucleotide sequence and characterization of genes encoding naphthalene dioxygenase of Pseudomonas putida strain NCIB 9816, Gene, 73, 355, 1988. 26. Yeh, W. K., Gibson, D. T., and Liu, T. N., Toluene dioxygenase: a multicomponent enzyme system, Biochem. Biophys. Res. Commun., 78, 401, 1977. 27. Subramanian, V., Liu, T. N., Yeh, W. K., and Gibson, D. T., Toluene dioxygenase: purification of an iron-sulfur protein by affinity chromatography, Biochem. Biophys. Res. Commun., 91, 1131, 1979. 28. Subramanian, V., Liu, T. N., Ye, W. K., Narro, M., and Gibson, D. T., Purification and properties of NADH-ferredoxinTOL reductase, a component of toluene dioxygenase from Pseudomonas putida, J. Biol. Chem., 256, 2723, 1981. 29 Subramanian, V., Liu, T. N., Yeh, W. K., Serdar, C. M., Wackett, L. P., and Gibson, D. T., Purification and properties of ferredoxinTOL, a component of toluene dioxygenase from Pseudomonas putida Fl, J. Biol. Chem., 260, 2355, 1985. 30. Finette, B. A., Subramanian, V., and Gibson, D. T., Isolation and characterization of Pseudomonas putida PpFl mutants defective in the toluene dioxygenase enzyme system, J. BacterioL, 160, 1003, 1984. 31. Zylstra, G. J., McCombie, W. R., Gibson, D. T., and Finette, B. A., Toluene degradation by Pseudomonas putida Fl: genetic organization of the /ot/operon, Appl. Environ. MicrobioL, 54, 1498, 1988. 32. Zylstra, G. J. and Gibson, D. T., Toluene degradation by Pseudomonas putida Fl: nucleotide sequence of the te*/ClC2BADE genes and their expression in Escherichia coli, J. Biol. Chem., 264, 14940, 1989. 33. Sauber, K., Froehner, C., Rosenberg, G., Eberspacher, J., and Lingens, F., Purification and properties of pyrazon dioxygenase from pyrazon-degrading bacteria, Eur. J. Biochem., 74, 89, 1977. 34. Furukawa, K. and Miyazaki, T., Cloning of a gene cluster encoding biphenyl and chlorophenyl degradation in Pseudomonas pseudoalcaligenes, J. BacterioL, 166, 392, 1986. 35. Hayase, N., Taira, K.» and Furukawa, K., Pseudomonas putida KF715 bph ABCD operon encoding biphenyl and poly chlorinated biphenyl degradation: cloning, analysis, and expression in soil bacteria, J. BacterioL, 172, 1160, 1990. 36. Markus, A., Krekel, D., and Lingens, F., Purification and some properties of component A of the 4chlorophenyacetate 3,4-dioxygenase from Pseudomonas species strain CBS, J. Biol. Chem., 261, 12883, 1986. 37. Schweizer, D., Markus, A., Seez, M., Ruf, H. H., and Lingens, F., Purification and some properties of component B of the 4 chlorophenylacetate 3,4 dioxygenase from Pseudomonas species strain CBS 3, /. Biol. Chem., 262, 9340, 1987. 38. Locher, H. H., Leisinger, T., and Cook, A. M., Degradation of p-toluenesulphonic acid via sidechain oxidation, desulphonation and meta ring cleavage in Pseudomonas (comamonas) testosteroni T-2, /. Gen. MicrobioL, 135, 1969, 1989. 39. Bernhardt, F. H., Ruf, H. H., Staudinger, H., and Ullrich, V., Purification of a 4-methoxybenzoate 0-demethylase from Pseudomonas putida, Hoppe-Seyler's Z. Physiol. Chem., 352, 1091, 1971. 40. Bernhardt, F. H., Heymann, E., and Traylor, S., Chemical and spectral properties of putidamonooxin, the iron-containing and acid-labile-sulfur-containing monooxygenase of a 4-methoxybenzoate 0-demethylase from Pseudomonas putida, Eur. J. Biochem., 92, 209, 1978. 41. Bernhardt, F. H. and Kuthan, H., Kinetics of reduction of putidamonooxin by NADH-puditamonooxin oxidoreductase, sodium dithionite and superoxide radicals, Eur. J. Biochem., 130, 99, 1983.

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42. Bernhardt, F. H., Nastainczyk, W., and Seydewitz, V., Kinetics studies on a 4-methoxybenzoate Odemethylase from Pseudomonas putida, Eur. J, Biochem,, 72, 107, 1977. 43. Bernhardt, F. H., Bill, E., Trautwein, A. X., and Twilfer, H., 4-Methoxybenzoate monooxygenase from Pseudomonas putida: isolation, biochemical properties, substrate specificity, and reaction mechanisms of the enzyme components, Methods Enzymol., 161, 281, 1988. 44. Bernhardt, F. H., Pachowsky, H., and Staudinger, H., A 4-methoxybenzoate Odemethylase from Pseudomonas putida: a new type of monooxygenase system, Eur. J. Biochem., 57, 241, 1975. 45. Twilfer, H., Bernhardt, F. H., and Gersonde, H., An electron-spin-resonance study on the redox-active centers of the 4-methoxybenzoate monooxygenase from Pseudomonas putida, Eur. J. Biochem., 119, 595, 1981. 46. Eich, F., Geary, P. J., and Bernhardt, F. H., Protein-protein interactions and antigenic relationships between the components of 4-methoxybenzoate monooxygenase and the benzene 1,2-dioxygenase from Pseudomonas putida, Eur. J. Biochem,, 153, 407, 1985. 47. Wende, P., Bernhardt, F. H., and Pfleger, K., Substrate-modulated reactions of putidamonooxin: the nature of the active oxygen species formed and its reaction mechanism, Eur. J. Biochem., 181, 189, 1989. 48. Brunei, F. and Davison, J., Cloning and sequencing of Pseudomonas genes encoding vanillate demethylase, /. Bacterial., 170, 4924, 1988. 49. Buswell, J. A. and Ribbons, D. W., Vanillate 0-demethylase from Pseudomonas species, Methods Enzymol., 161, 294, 1988. 50. Morrice, N., Geary, P., Cammack, R., Harris, A., Beg, F., and Aitken, A., Primary structure of protein B from Pseudomonas putida, member of a new class of 2Fe-2S ferredoxins, FEES Lett,, 231, 336, 1988. 51. Irie, S., Doi, SM Yorifuji, T., Takagi, M., and Yano, K., Nucleotide sequencing and characterization of the genes encoding benzene oxidation enzymes of Pseudomonas putida, J. BacterioL, 169, 5174, 1987. 52. Schell, M. A., Homology between nucleotide sequences of promoter regions of nah and sal operons of NAH7 plasmid of Pseudomonas putida, Proc. Natl. Acad. Sci. U.S.A., 83, 369, 1986. 53. Locher, H. H., Leisinger, T., and Cook, A. M., 4-Toluene sulfonate methyl-monooxygenase from Comamonas testosteroni T-2: purification and some properties of the oxygenase component, /. BacterioL, 173, 3741, 1991. 54. Gibson, D. T., Zylstra, G. J., and Cruden, D., personal communication. 55. Neidle, H. and Ornston, L. N., personal communication. 56. The reported R is for all the data between 40 to 2.0 A. The current structure incorporates sequence information from peptides that account for 215 of the 318 residues in the model; an additional 63 residues have been assigned based on features in the electron density maps, and the remaining residues were modeled as alanine or glycine. The model also contains 143 water molecules. The peptide fragments were positioned by alignment with the vanB gene sequence from vanillate demethylase,48 which is 38% identical in the known regions. 57. Callahan, T. J., Gleason, W. B., and Lybrand, T. P., PAP: A protein analysis package, Am. CrystallographicAbstr., 18, 73, 1990. 58. Abbreviations: AAD, 3-aminopyridine adenine dinucleotide and TNAD, thionicotinamide. 59. Karplus, P. A., Daniels, M. J., and Herriott, J. R., Atomic structure of ferredoxin-NADP+ reductase: prototype for a structurally novel flavoenzyme family, Science, 251, 60, 1991. 60. Rossman, M. G., Liljas, A., Branden, C.-I., and Banaszak, L. J., Evolutionary and structural relationships among dehydrogenases, The Enzymes, 11A, 62, 1975. 61. Rypniewski, W. R., Breiter, D. R., Benning, M. M., Wesenberg, G., Oh, B.-H., Markley, J. L., Rayment, I., and Holden, H. M., Crystallization and structure determination of 2.5 A resolution of the oxidized [2Fe-2S] ferrodoxin isolated from Anabaena 7120, Biochemistry, 30, 4126, 1991. 62. Manstein, D. J., Massey, V., and Pai, E. F., Absolute stereochemistry of flavins in enzyme-catalyzed reactions, in Flavins and Flavoproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 3. 63. Fukuyama, K., Hase, T., Matsumoto, S., Tsukihara, T., Katsube, Y., Tanaka, N., Kakudo, M., Wada, K., Matsubara, H., Structure of S. platensis [2Fe-2S] ferredoxin and evolution of chloroplast-type ferredoxins, Nature, 286, 522, 1980. 64. Buchanan, B. B. and Arnon, D. I., Ferredoxins from photosynthetic bacteria, algae, and higher plants, Methods EnzymoL, 23A, 413, 1971. 65. Beinert, H. and Orme-Johnson, W. H., Electron spin resonance as a probe for active centers of paramagnetic enzyme species, in Magnetic Resonance in Biological Systems, Vol. 9, Ehrenberg, A., Malmstrom, B. G., and Vanngard, T., Eds., Pergamon Press, New York, 1967, 221.

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Chapter 19

NUCLEAR MAGNETIC RESONANCE STUDIES ON FLAVOPROTEINS Franz Miiller

TABLE OF CONTENTS I.

Introduction

558

II.

31

P NMR A. FMN-Dependent Enzymes B. FAD-Dependent Enzymes

560 560 565

III.

15

569

IV.

13

574

V.

19

581

VI.

17

583

VII.

'H NMR A. Conventional J H NMR B. Two-Dimensional (2D) NMR

584 584 586

VIII.

Conclusion

588

Acknowledgments

589

References

590

Appendix with References

594

N NMR

C NMR F NMR

O NMR

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Chemistry and Biochemistry of Flavoenzymes

I. INTRODUCTION Unlike other physical techniques nuclear magnetic resonance (NMR) has not been applied routinely to flavoproteins until recently. This is partly due to the fact that flavoproteins possess generally molecular weights exceeding 20,000. In 'H NMR spectra this physical property of flavoproteins leads to highly complex and often little resolved spectra because of the wealth of protons present in the proteins leading to overlaps of resonance lines. Also with increasing molecular weight the resonance lines broaden, generally counteracting resolution. During the last decade the hardware and software of NMR instruments have been developed to a high degree of sophistication. It is now possible to resolve the solution structure of proteins up to a molecular weight of about 20,000 by two-dimensional ]H NMR techniques. In addition other nuclei like 31P, 13C, 15N are also amenable to NMR techniques. All these combined make the NMR techniques into a powerful tool allowing to obtain information on molecular, submolecular, and atomic levels. Since the NMR techniques will undoubtedly play a more prominent role in the future in flavoprotein research, it seems appropriate to summarize for the first time briefly the presently available NMR data on flavoproteins. The interpretation of especially the 13C and 15N NMR spectra of flavoproteins relies heavily on the interpretation of spectra of free flavins in different solvents. Therefore, the 13 C and 15N NMR spectra of free flavins are discussed briefly in order to facilitate the discussion on the 13C and 15N NMR data of flavoproteins. The following discussion is an extension of the NMR data presented in Volume I of this series by the present author.1 Flavoproteins contain in most cases either FMN or FAD as a prosthetic group which is generally tight but noncovalently bound to the apoflavoprotein. The structures of FMN and FAD in the oxidized state are shown in Figure 1 together with the two-electron reduced state in the neutral and ionized forms. These redox states and the neutral form of the flavosemiqumone bound to apoflavoproteins will play a role in the discussions presented below. To study the protein-bound flavin by 13C and 15N NMR techniques, selective, isotopic enrichment of flavin is required. The synthesis of these molecules has been described.1-2 In the oxidized and two-electron reduced states there exists a fairly good linear relationship between the experimental 13C chemical shift and the calculated TT electron density of the corresponding carbon atom.2'3 This fact allows to observe TT electron density changes in the flavin molecule induced by different environments (solvent polarity) or upon binding to apoflavoproteins. For instance a TT electron density increase on a particular carbon atom will lead to an upfield shift, and a decrease in TT electron density will show an opposite effect on the 13C chemical shift. Tetraacetylriboflavin (TARF) in an apolar organic solvent (e.g., CHC13) serves as a reference where hydrogen bonding interaction with the solvent is diminished and self-association of flavin is minimized. Hydrogen bond formation with the carbonyl groups at position 2 and 4 of flavin occurs even in protic organic solvents and leads to downfield shifts of the corresponding 13C chemical shifts. The extent of the shifts depends on the dielectric constant of the solvent and leads in water also to a high degree of polarization of the flavin ring and to various possible mesomeric structures.4 The nitrogen atoms in flavin can be characterized as pyridine- or (3-type nitrogen atoms [N(l) and N(5) in the oxidized molecule] and pyrrole- or a-type nitrogen atoms [N(10) and N(3) in oxidized and all four nitrogen atoms in two-electron reduced flavin]. The 15N chemical shifts of pyridine-like nitrogen atoms are very sensitive to hydrogen bonding interactions leading to upfield shifts. The I5N chemical shifts of pyrrole-like nitrogen atoms are little sensitive to hydrogen bonding exhibiting a small downfield shift upon hydrogen bond formation. The unexpected downfield shift of the resonance due to N(10) when the polarity of the

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559

FIGURE 1. The chemical structures of FMN and FAD in the oxidized and two-electron reduced states. The internationally accepted numbering of the atoms is also shown.

solvent is increased, is interesting. Since this atom cannot form a hydrogen bond, the downfield shift must be caused by some other effect. The clue to the solution is found in the downfield shifts of the resonances due to C(9) and C(7), which parallel the downfield shift of the N(10) atom. These data strongly indicate that the N(10) atom carries a partial positive charge, which is distributed to the C(7) and C(9) atoms of the xylene subnucleus of the flavin, leading to the downfield shifts. The partial positive charge on N(10) is created by the delocalization of the lone electron pair of the N(10) atom to C(4a) (upfield shift) and further to 0(4ot).4 From the above described data, it follows that the degree of hybridization of the N(10) atom depends strongly on the polarity of the microenvironment of the flavin molecule, i.e., in CHC13 the N(10) atom possesses a higher degree of sp3 hybridization than in a polar solvent. Obviously this could provide a mechanism by which the chemical reactivity of protein-bound flavin could be regulated by specific hydrogen bond formation with the apoflavoprotein and by providing the appropriate microenvironment.5 Oxidized flavin contains one- and two-electron reduced flavin two or three, depending on the ionization, NH groups. The 15N-!H coupling constants yield valuable information about the degree of hybridization of the nitrogen atoms in question and the accessibility of these groups to bulk solvent in the protein-bound flavin. The observation of a doublet in protein-bound flavin demonstrates that no or only a very slow hydrogen exchange occurs,

560

Chemistry and Biochemistry of Flavoenzymes

a singlet would indicate a fast proton exchange and hence accessibility to bulk solvent. A I5 N- J H coupling constant of about 75 Hz indicates a high degree of sp3 hybridization and a value of about 90 Hz a high degree of sp2 hybridization of the corresponding nitrogen atom. In flavoproteins the situation may not be as clear cut as suggested above, because involvement of a NH group in hydrogen bonding would decrease the 15N-'H coupling constant while the hybridization of the nitrogen atom could remain unaffected. This is due to the fact that the coupling constant is a scalar term, i.e., hydrogen bonding increases the N-H bond length thereby decreasing the coupling constant. Recently it has been discovered that N(5)H in two-electron reduced free flavin exhibits a peculiar behavior towards a proton exchange.6 In the pH range 5 to 6 and above 11 a sharp line is observed for N(5) in the 15N NMR spectra indicating a fast acid and base catalyzed exchange. In the pH range 6 to about 8, an intermediate exchange rate, and in the pH range 8 to 10, a doublet is observed indicating no or only a very slow exchange reaction. Therefore, the observation of a doublet of N(5)H in reduced protein-bound flavin does not imply that this group is not accessible to bulk solvent because the very slow proton exchange reaction seems to be an inherent property of ionized reduced free flavin.6 The ionization of N(3)H in oxidized and N(1)H in reduced flavin has large effects on the neighboring carbon atoms. As shown in Figure 2 the pKa values determined by 15N NMR are also reflected by the neighboring carbon atoms due to the electric field effect of the negatively charged N atoms on the neighboring carbon atoms.4-7~12 All the resonances are shifted downfield. This is also true for atoms in para position to the ionization site, but the downfield shifts are less extensive. The fact that C(2) and C(10a) in reduced flavin are strongly affected upon ionization of N(l) lends the 13C chemical shifts of these atoms as reliable reporters of the ionization state of protein-bound flavin in 13C NMR spectra. Finally it should be mentioned that to obtain 13C and 15N NMR spectra of flavoproteins with a reasonable signal-to-noise ratio, a large concentration is needed. Although the isolation and purification of large amounts of proteins is time consuming, this problem can be handled. A by-far-larger hurdle is the development of methods allowing the preparation of large amounts of apoflavoproteins which are highly reconstituted. Such methods have been reviewed recently.13

II.

31

P NMR

The phosphorus atoms of the protein-bound flavins can be easily observed in native proteins. Caution must however, be exercised to avoid the possible presence of paramagnetic metal ions in the samples. Paramagnetic metal ions can lead to extensive line broadening of phosphorus resonances. However paramagnetic metal ions can be removed from solutions by Chelex chromatography. On the other hand, manganese ions causing line broadening if the phosphate group is accessible, are often purposely added to samples in order to investigate the possible access of bulk solvent to protein-bound phosphates. The 31P chemical shifts can be influenced 1. 2. 3.

By conformational changes altering the O-P-O or P-O-P bond angle distribution: Gorenstein14'15 has shown that 13P chemical shifts of phosphate esters may depend on P-O-P and O-P-O bond angles; The proximity of the phosphate chain to an aromatic ring could expose the phosphates to ring-current shielding and deshielding; Hydrogen bonding and strong electrostatic interactions could also influence 31P chemical shifts.

A. FMN-DEPENDENT ENZYMES Some illustrative examples of 31P NMR spectra of FMN-dependent proteins are shown

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FIGURE 2. The pH dependence of 13C and I5 N chemical shifts of oxidized and two-electron reduced free flavin. These data were taken from various publications. 4 - 712

in Figure 3. The spectrum of Megasphaera elsdenii flavodoxin16'17 represents the usual spectra of flavodoxins containing only one resonance line due to the phosphate group of FMN. In contrast the spectrum of Azotobacter vinelandii flavodoxin18'19 shows additional resonance lines at high field. Both these phosphate residues are covalently bound to the apoflavodoxin. However, one phosphate residue is subject to acid hydrolysis and is lost when the protein is treated with trichloroacetic acid. The acid stable phosphate residue has

562

Chemistry and Biochemistry of Flavoenzymes

FIGURE 3. 3I P NMR spectra of Megasphaera elsdenii (left) and Azotobacter vinelandii flavodoxins in the oxidized (A), semiquinone (B) and reduced (C) state.

been shown to be disubstituted by 1H-31P two-dimensional NMR.20 The phosphate residue is bound to a seryl and a threonyl residue. The 31P NMR resonance line due to FMN in flavodoxins — this is in fact also valid for all other FMN-dependent proteins — is pH-independent or only little dependent on pHvalues indicating that the phosphate group is inaccessible to bulk solvent. This interpretation is fully supported by the fact that addition of Mn 2+ does not influence the resonance line of the phosphate group. Furthermore, the chemical shift due to the phosphate moiety of the protein-bound flavin in flavodoxins resembles that of FMN at pH 9 strongly indicating that the phosphate moiety of protein-bound FMN is in the diionic state. Reduction of flavodoxins to the semiquinone state yields 31P NMR spectra in which the FMN phosphate resonance line is broadened (Figure 3). The extent of broadening varies depending on the flavodoxin, suggesting that the distance between the phosphate and the isoalloxazine moiety in protein-bound FMN varies. The distances calculated from the 31P NMR spectra are in good agreement with X-ray data.16-21 Upon stepwise reduction of oxidized flavodoxin to the semiquinone state, a concomitant increase of the line width is observed indicating a slow electron exchange between the two states.16 Further reduction to the twoelectron reduced state yields intermediate spectra showing a sharp line superimposed on the broad line. In reduced flavodoxin a sharp line is observed again whose chemical shift differs little from those of oxidized flavodoxins. Kinetic measurements by 31P NMR indicate that the electron exchange between the semiquinone and the two-electron reduced state is fast, and slow between the oxidized and the semiquinone state (see also below). The presently available data on FMN-containing proteins are collected in Table 1. The 31 P chemical shifts of flavodoxins from Clostridium MP17 (more recently characterized as Clostridium beijerinckii), M. elsdenii^11 Desulfovibrio vulgaris17-22 D. gigas,22 A. vinelandii,17'19 and Anabaena 712023 are all very similar. The values determined at earlier times are at somewhat higher field and are partly due to the lower resolution of the instruments used at that time and partly due to the reference standard used.

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563

TABLE 1 P NMR Chemical Shifts in Flavoproteins Chemical shift (ppm)a of

Compound Free FMN Oxidized

Reduced Flavodoxin from Clostridium MP Oxidized Reduced Megasphaera elsdenii Oxidized

Semiquinone Hydroquinone Reconstituted with riboflavin 3', 5'bisphosphate Oxidized Reduced Desulfovibrio vulgaris Oxidized Semiquinone Hydroquinone

pH

FMN

4.0 7.0 9.0 9.0 9.0

0.90 4.01

8.0 8.0

FAD

Others

Ref.

—

— —

16 16 16 17 17

5.7 5.8

— —

— —

17 17

6.0—9.2 8.0 5.5 8.2 8.2 8.0

4.8 5.3 4.5

— — — — —

—

—

— — — — —

16 17 16 16 16 17

8.0 8.0

5.2 5.3

— —

2.0 1.6

24 24

7.8 8.0 7.8 7.8 8.0

4.99

—

5.4 4.99b 5.5

— — — —

— — —

5.5 5.6 4.99 4.99b

Reconstituted with riboflavin 3', 5'bisphosphate 8.0 Oxidized 8.0 Reduced Desulfovibrio gigas 7.8 Oxidized 7.8 Semiquinone 7.8 Hydroquinone Azotobacter vinelandii Oxidized 7.5 Form I 7.5 Form II 7.5 Form III 5.5—9.5 Oxidized 8.0 8.0 Semiquinone 8.0 Hydroquinone 8.0 Anabaena 7120 8.0 Oxidized 8.0 Reduced Bacterial luciferase 7.0 Oxidized 7.0 Reduced Old Yellow Enzyme 8.6, 8.0 Oxidized, OYE(O) 8.0 Oxidized, OYE(l)

4.7 5.1 5.1

4.9b

4.9 5.4

4.99

4.9 5.33 6.03 5.48

5.6 6.3 ~5.6C 5.4 6.4

— — — —

— —

22 17 22 22 17

— —

0.8 0.3

24 24

— — —

— — __

22 22 22

— — —

25 25 25

— — — — — — — —

0.8, o:>

18, 19

—

18, 19 18, 19

0.9

0.4 1.0

17

17

—

23 23

— —

—

—

26 26

— —

—

7, 28

5.4 5.9

—

5.2 5.6 8.6, 8.9

-12

— — —

—

—

—

7

564

Chemistry and Biochemistry of Flavoenzymes 31

TABLE 1 (continued) P NMR Chemical Shifts in Flavoproteins Chemical shift (ppm)11 of

Compound Free FAD Oxidized

pH 1 .7—10.5

Glucose oxidase Oxidized 5 .0—8.0 Reduced 8.0 Adrenodoxin reductase Oxidized 6.93 6-Hydroxy-L-nicotine oxidase Oxidized 7.5 Xanthine oxidase Oxidized 8.0 /?-Hydroxybenzoate hydroxylase Oxidized 7.0 Glutathione reductase Oxidized 7.0 Mercuric ion reductase Oxidized 7.5 — + Mn 2 + Lipoamide dehydrogenase Oxidized 7.0 D-Amino acid oxidase Oxidized 7.0 Oxynitrilase Oxidized 7.0 Ferredoxin-NADP+ reductase Oxidized 8.0 NADPH-Cytochrome P-450 reductase Protease solubilized 7.65 7.7 Detergent solubilized 7.7 FMN depleted6 7.7 a b c d e

FMN

FAD

Others

Ref.

— — —

-9.87, -10.55 -10.4, -11.1 -10.8, -11.3

— — —

29 30 31

— —

-10.8, --13. 3 -10.8, --13.3

-2.0 -2.0

31 31

—

-10.0, -14.3

—

32

-12.2d

1.3, 0.7, -0.3, -0.6 33

-8.8, -13.5

-1, ~3

—

-9.0, -9.6

34 35

—

-9.7, -10.5

—

36

— —

-12.1, -12.9 -12.1, -12.9

-10.5 —

36 36

-8.4, -12.4 —

-11.2, -13.5

36 —

36

-11.2, -12.3

—

-7.3, -10.0

4.4, 3.98 -7.33, -11.25 4.1 -7.4, -11.3 4.1 -7.4, -11.3 --9.6, --10.4

36

—

40

1.7 1.8 1.8,2.9,0.4, -0.3

29 40 40 40

Chemical shifts are reported relative to external H3PO4. The resonance line is considerably broadened in the presence of the flavosemiquinone. Resonance line broadened beyond detection. Unresolved. Various resonances.

Although apoflavodoxins bind quite specifically FMN, riboflavin 3',5'-bisphosphate is also strongly bound by apoflavodoxins from M. elsdenii and D. vutgaris.24 The 3'-phosphate group in these flavodoxins resonates at about 2 ppm in M. elsdenii and about at 0.5 ppm in D. vulgaris flavodoxins (Table 1). Both 3'-phosphate groups are accessible to Mn 2 + , the group in M. elsdenii flavodoxin to a lesser extent than the one in D. vulgaris flavodoxin.24 These data and the chemical shifts indicate that the 3'-phosphate group in the former protein is monoionized where it is probably protonated in the latter. Three different forms of flavodoxins have been discovered in A. vinelandii, distinguished by different chemical shifts of the phosphate group of the protein-bound FMN.25 The flavodoxins were obtained under different growth conditions which are believed to be the reason for the synthesis of three different forms of flavodoxins. As can be seen from Table 1 the 31P chemical shifts of protein-bound FMN in flavodoxins

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generally undergo a small downfield shift upon two-electron reduction. A larger downfield shift is observed upon reduction in Anabaena 7120 flavodoxin23 and in bacterial luciferase.26 Bacterial luciferase binds FMN relatively weakly in the oxidized state as compared with other flavoproteins, while reduced FMN is much more strongly bound. This is apparently reflected in the chemical shift. Although bacterial luciferase and A. vinelandii apoflavodoxin bind the prosthetic group more strongly in the reduced than in the oxidized state, the contrary is true for M. elsdenii and D. vulgaris flavodoxins, but the chemical shift trends do not reflect this fact (Table I). 27 It is therefore suggested that the downfield and upfield shifts are caused by some minor environmental and/or conformational changes in the immediate vicinity of the phosphate group. A small configurational change of the phosphate group is another possibility. So far an exception within the FMN-dependent proteins with respect to the 31P chemical shift represents Old Yellow Enzyme. Its chemical shift in the oxidized free state appears at about 8.6 ppm (Table I).7-28 The resonance is further downfield shifted by about 3 ppm on complexation with an organic ligand.7 It has been suggested7 that a positive charge is localized close to the phosphate group causing the resonance to shift downfield as compared to the 31 P chemical shifts of flavodoxins. Since the enzyme interacts with the ligand in the anionic state, a positive charge is neutralized in the enzyme upon complex formation. The removal of a positive charge in the enzyme would then predict the phosphorus resonance to shift upfield. This is not the case, so other effects must play an important role, e.g., ring current effects by an aromatic amino acid residue or a large conformational change in the phosphate binding site. B. FAD-DEPENDENT ENZYMES The 31P NMR spectrum of free FAD exhibits an AB splitting pattern (Figure 4). The chemical shifts published by various groups are given in Table I.29'31 The 31P-31P coupling constant has been determined to be 20.9 Hz.30 The lowfield part of the FAD signal was assigned to the FMN and the upfield signal to the AMP moiety (Table I).30 As could be expected the resolution observed with free FAD is lost on binding to apoflavoproteins but the two phosphorus residues can in most cases still be observed in the 31 P NMR spectra (Table 1). The assignment of the two resonances to the FMN and AMP moiety of FAD cannot be done with certainty since a positional change of the resonance lines cannot be excluded. The 31P NMR spectrum of glucose oxidase contains in addition to the FAD resonances a resonance at low field (Table I). 31 The phosphate residue was still present in the apoprotein, in the protein treated with periodate or pronase. From this it was concluded that the phosphate residue was covalently bound to the protein.31 The addition of Mn 2+ to a solution of native protein did not affect the FAD resonance lines but led to a considerable broadening of the line of the covalently bound phosphate indicating its accessibility to bulk solvent. The 31P chemical shifts of FAD bound to adrenodoxin reductase differ somewhat from those of glucose oxidase, the line widths in the spectrum of glucose oxidase are broader than those in adrenodoxin reductase.32 The complex between the enzyme and NADP + has also been studied. The resonance lines of the 2'-phosphate and the pyrophosphate group of bound NADP + are shifted downfield and upfield, respectively, as compared to those of free NADP + . 6-Hydroxy-L-nicotine oxidase from Arthrobacter oxidans is so far the only flavoprotein whose 31P NMR spectrum shows an unresolved resonance for the pyrophosphate group (Table I).33 This spectrum also exhibits several resonances at low field. The chemical structure of these phosphate residues is not known but probably represent diester phosphates. It was observed that the intensity of the various peaks differed from preparation to preparation, the relative intensity of the peaks remained almost constant. Based on the observation that

566

Chemistry and Biochemistry of Flavoenzymes

FIGURE 4. 31P NMR spectra of free FAD and various flavoproteins in the oxidized state. (A) Free FAD; (B) mercuric ion reductase; (C) mercuric ion reductase in the presence of Mn2 + ; (D)p-hydroxybenzoate hydroxylase; (E) lipoamide dehydrogenase; (F) D-amino acid oxidase; (G) oxynitrilase.

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extensive dialysis of the protein did not lead to an apparent loss of phosphate residues it was suggested that they are covalently bound to the protein.33 The phosphate residues are located on or close to the surface of the protein as indicated by Mn 2 + -induced line broadening.33 The 31P NMR spectrum of bovine milk xanthine oxidase shows four resonances due to the pyrophosphate group of FAD, the phosphate group of the pterin cofactor, and a phosphate residue covalently bound, accessible to Mn 2+ and monosubstituted.34 Despite the large size of the protein (360 kDa) a reasonable spectrum was obtained from about 60,000 acquisitions. Another surprising observation was that the intensity of the phosphoseryl residue (~3 ppm) was dependent on the functionality of the enzyme, i.e., it increased as the functionality of the enzyme was decreased by cyanide treatment. In Figure 4 several yet unpublished 31P NMR spectra of FAD-containing proteins are shown. All spectra of FAD-dependent proteins show, with the exception of 6-hydroxy-Lnicotine oxidase, a spacing of at least 2 ppm between the two resonances due to the pyrophosphate group. Lipoamide dehydrogenase from Azotobacter vinelandii, D-amino acid oxidase (Figure 4) and spinach ferredoxin-NADP+ reductase (Table 1) exhibit a similar pattern. On the other hand, the spacing between the pyrophosphate peaks is only about 1 ppm in the spectra of p-hydroxybenzoate hydroxylase from Pseudomonas fluorescens,35 glutathione reductase, mercuric ion reductase, and oxynitrilase (Table 1, Figure 4),36 Glutathione reductase, lipoamide dehydrogenase, and mercuric ion reductase belong to the same family of FAD-containing pyridine nucleotide: disulfide oxidoreductases,37 the 31P NMR spectral properties of lipoamide dehydrogenase do not reflect this fact. On the other hand, it has been shown that glutathione reductase and thep-hydroxybenzoate hydroxylase-substrate complex exhibit a great similarity with respect to the three-dimensional structure of the FAD domains.38 This is reflected to some extent by the 31P NMR spectra (Table 1). Yet the pyrophosphate group in /?-hydroxybenzoate hydroxylase is not accessible to Mn 2+ while the group in glutathione reductase is. Another difference between the two enzymes is that the pyrophosphate group of the former enzyme interacts with two arginine residues, the latter is not in direct contact with such residues. We have undertaken an attempt to explain the different pyrophosphate chemical shifts in FAD-dependent enzymes by searching for a correlation between the chemical shifts and the P-O-P, O-P-O angles using the available Xray data. No reliable correlation was found which is probably due to the still rather inaccurate torsional angles deduced from X-ray data, especially those from p-hydroxybenzoate hydroxylase. As seen in Table 1 mercuric ion reductase contains one additional *'phosphate" group resonating at about -10 ppm. This "phosphate" group is not lost upon apoprotein preparation and reconstitution of the enzyme although the procedure includes several steps (dialysis, column chromatography).39 The addition of Mn2+ to the enzyme broadens the resonance of the "phosphate" group beyond detection (Figure 4). This procedure also ascertained the assignment of the resonances due to protein-bound FAD and suggests that the "phosphate" group is covalently bound to the protein. NADPH-cytochrome P-450 reductase contains FMN and FAD as prosthetic groups,31? NMR spectra of the enzyme are shown in Figure 5 and 31P chemical shifts collected in Table 1. Two preparations have been studied by 31P NMR: the protease-solubilized and the detergent-solubilized proteins. The former preparation exhibits only an artificial activity using cytochrome c as an electron acceptor. One-electron reduction of both preparations yields the "air-stable" semiquinone. Surprisingly, in contrary to expectation, little line broadening is observed on the resonance due to the phosphate moiety of FMN.29 This observation was confirmed independently by another research group.40 The expected line broadening was observed in the semiquinone form of the native, detergent-solubilized enzyme.40 The different behavior of the two enzyme preparations was explained by different conformations.40 This

568

Chemistry and Biochemistry of Flavoenzymes

FIGURE 5. 31P NMR spectra of NADPH-cytochrome P-450 reductase. Protease-solubilized enzyme in the oxidized (A) and semiquinone (B) state. Detergent-solubilized enzyme in the oxidized (C) and semiquinone (D) state. (E) FMN-depleted protein (detergent-solubilized).

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interpretation was supported by the observation that A. vinelandii flavodoxin which did not contain the acid-labile phosphate group behaved like the protease-solubilized enzyme. The same flavodoxin in a complex with ferredoxin-NADP+ reductase exhibited again line broadening in the semiquinone state.40 The 31P NMR spectra of NADPH-cytochrome P-450 reductase shows additional lines beside the resonances of the flavin prosthetic groups. In both the detergent- and proteasesolubilized preparations a line resonating at 1.8 ppm is observed (Figure 5).29'40 This line was assigned to AMP which is noncovalently but very strongly bound to the enzymes, introduced by using an affinity chromatography requiring AMP in the elution solution. The protease-solubilized preparation contains additional phosphate residues resonating at high field (Figure 5).40 The number of lines and their intensities varied from preparation to preparation and seem also to be dependent on the source of the enzyme (liver from pig, rabbit, rat). It has been suggested that these lines are due to phospholipids, i.e., phosphatidylcholine. These residues are noncovalently bound to the enzyme since extensive dialysis of the enzyme leads to an intensity decrease of the corresponding lines. However a fully phospholipid-depleted preparation could not be obtained, not in concentrations needed for 31 P NMR.40 It cannot be excluded that phosphatidylethanolamine is the constituent in the enzyme as has been proposed previously by others based on data obtained by a combination of techniques.41 At any rate the phospholipids play probably an important role in the regulation of the activity of the enzyme, at least in vitro.40 Very recently it was shown that optimal cytochrome P-450 activity could be reconstituted in the presence of phosphatidylcholine, phosphatidylserine, diolenoylphosphatidylcholine (18:1), didecanoylphosphatidylcholine (10:0) or dilauroylphosphatidylcholine (12:0).42 The pyrophosphate resonances in FMN-depleted NADPH-cytochrome P-450 reductase undergo a considerable change as compared to those of the original enzyme, indicating a conformational change in the FAD binding domain (Figure 5).40 Reconstitution with FAD is accompanied with a time-dependent generation of the original 31P NMR spectrum.40 31 P NMR has only slightly been exploited in the field of flavoproteins. A solid base is now available on which further research can be built up. Especially in times where enzymes can be expressed in high yields by cloning, the 31P NMR spectroscopy has a high potential to study flavoproteins in their natural environments, and to obtain possibly further information about protein-protein interactions and the catalytic mechanisms under in vivo conditions.

III.

15

N NMR

As already outlined in the introduction, the 15N NMR spectra yield primarily information about the degree of hybridization, deduced from the 15N chemical shift in combination with 15 N-'H coupling constant, the involvement of a particular nitrogen atom in hydrogen bonding, the accessibility of NH groups to bulk solvent (proton exchange) with the exception of N(5)H (see above and also below) and the ionization state of an ionizable NH group. In addition the 15N chemical shifts of free and protein-bound flavins are dependent on the redox state of the molecule. A selection of 15N NMR spectra in the oxidized and reduced states are shown in Figure 6. From these spectra it is obvious that the resonances of all four nitrogen atoms in proteinbound flavin can be observed without any interference from 15N natural abundance nuclei. Depending on the size of the protein natural abundance, resonances appear at about 125 ppm in the spectra; these resonances are due to amide groups in the proteins. The 15N chemical shifts of free and protein-bound flavins are presented in Table 2. Compared to TARF in CHC13 and FMN in aqueous solution (Table 2) the five flavodoxins so far studied show that N(l) of the oxidized prosthetic group is hydrogen-bonded with

570

Chemistry and Biochemistry of Flavoenzymes

FIGURE 6. 15N NMR spectra of various flavoproteins in the oxidized and reduced state. (A) Desulfovibrio vulgaris flavodoxin; (B) (C) riboflavin-binding protein from egg white; (D) Old Yellow Enzyme; (E) (F) bacterial luciferase; (G) (H) glucose oxidase.

571

Volume III TABLE 2 N NMR Chemical Shifts of Free and Protein-Bound Flavins

!5

Chemical shift (ppm)a Compound

PH

kl)a

N(l)

N(3)

N(5)

N(10)

Ref.

200.1 190.8

159.6 160.5

346.0 334.7

151.9 163.5

26 26

Oxidized TARF FMN Flavodoxin from Megasphaera elsdenii Clostridium MP Azotobacter vinelandii Desulfovibrio vulgaris Anabaena 7120 Bacterial luciferase Old Yellow Enzyme Glucose oxidase 6-Hydroxy-L-nicotine Oxidase + Inhibitor /?-Hydroxybenzoate Hydroxylase + Substrate + Product Riboflavin-binding protein from Egg yolk Egg white

CHC1, 7.0 7.0 8.0 8.0 8.0 7.5 7.0 8.5 8.5C 5.6

15.0 15.8 21.5 15.4 19.2 ~80b ~49c — 80C

185.0 184.5 186.4 188.0 188.0 187.1 194.3 194.8 195.0

160.9 161.1 160.2 159.9 162.5 162.3 164.1 164.2 161.8

349.3 351.5 341.4 341.1 335.0 325.8 319.4 320.8 336.2

165.6 164.8 161.5 165.6 163.5 160.8 — 161.5 164.3

8, 17 17 17 43 44 26 47 47 45

7.0 7.0

48C —

192.2 193.8

160.6 160.1

335.3 330.0

162.2 161.7

33 33

7.0 7.0 7.0

44C — —

191.6 189.2 188.4

159.8 159.8 159.5

328.5 348.0 361.5

165.6 165.6 165.9

35 35 35

9.0 6.2

36 32

191.5 191.6

159.2 159.4

338.2 338.2

165.4 165.3

46 46

119.9 128.0 181.3

149.0 149.7 150.0

59.4 58.0 58.4

76.8 87.2 96.5

26 26 26

Reduced TARFH2 FMNH2 FMNHFlavodoxin from Megasphaera elsdenii Clostridium MP Azotobacter vinelandii Desulfovibrio vulgaris Anabaena 7120 Bacterial luciferase Old Yellow Enzyme Glucose oxidase 6-Hydroxy-L-nicotine oxidase Dithionite reduced Substrate reduced p-Hydroxybenzoate Hydroxylase + Substrate + Product Riboflavin-binding protein from Egg yolk Egg white a b c d

CHC13 5.0 8.5

7.0 8.0 8.0 8.0 7.0 7.0 8.5 8.5d 5.6 7.4

15.0 15.8 21.5 15.4 19.2 80b ~49C — 80C —

183.4 182.8 182.0 186.6 182.5 176.8 187.4 186.5 186.2 186.9

149.7 150.1 150.0 148.3 151.5 150.0 153.2 152.5 153.8 153.8

61.3 61.9 61.7 62.1 55.0 59.9 48.6 57.1 57.3 53.0

98.3 97.7 96.7 98.4 95.6 94.6 — 97.6 96.3 96.2

8, 17 17 17 43 44 26 47 47 45 45

7.0 7.0

48C —

192.7 192.5

149.2 147.7

52.7 60.5

100.6 102.6

33 33

7.0 7.0 7.0

44C — —

183.9 179.8 181.3

148.2 149.0 150.0

60.9 61.5 58.4

98.4 94.3 96.5

35 35 35

6.3 6.3

36 32

128.5 129.8

148.2 148.9

59.8 59.9

89.9 90.3

46 46

The chemical shifts are relative to liquid NH 3 . Heterodimer. Subunit molecular weight; the protein is a dimer. The prosthetic group used was 7-methyl-10-ribityl-isoalloxazine 5'-phosphate.

572

Chemistry and Biochemistry of Flavoenzymes TABLE 3 ^N-1!! Coupling Constants of Free and Protein-Bound Flavins in the Oxidized and Reduced State Coupling constant (Hz) Compound

I

1

J[15N(5)-1H(5)]

J[ISN(3)-1H(3)]

Ref.

Oxidized state TARF in CHC13 FMN in H2O Flavodoxin from Clostridium MP Megasphaera elsdenii Desulfovibrio vulgaris Azotobacter vinelandii Anabaena 7120 Bacterial luciferase Old Yellow Enzyme Glucose oxidase 6-Hydroxy-L-nicotine oxidase /7-Hydroxybenzoate hydroxylase Riboflavin-binding protein

a

—

8 8

90.3 88.2 90.6 90.6 89.9

— — — — — — — — — — —

17 8 43 17 44 26 47 45 33 35 46

92.7

-90 90

~90b ~90b Not resolved a

Reduced state TARFH2 FMNH2 FMNHFlavodoxin from Clostridium MP Megasphaera elsdenii Desulfovibrio vulgaris Azotobacter vinelandii Anabaena 7120 Bacterial luciferase Old Yellow Enzyme Glucose oxidase

93.1 — * —a

a

90

8 8 6

91.6 93.1 89.8 89.7

94.0 92.1 86.2 90.3

17 8 43 17

Not determined Not resolved

~85b

88

70 Not resolved

6-Hydroxy-L-nicotine oxidase /7-Hydroxybenzoate hydroxylase

90

Riboflavin-binding protein

Not observed

a b

87.5

78 Not resolved Not observed

26 47 45 33 35 46

Not observed, fast proton exchange. Approximate value.

amino acid residues of the protein. The hydrogen bonds are stronger in flavodoxins from M. elsdenii and C. MP than in others in which the strength of the hydrogen bonds still exceeds the one in FMN. Also the N(3)H group of FMN in all flavodoxins is hydrogen bonded. The weakest hydrogen bonds are observed in A. vinelandii17 and D. vulgaris43 flavodoxins, the strongest in Anabaena 7120 flavodoxin.44 This fact is also nicely reflected by the 1J[15N(3)-1H(3)] (Table 3). Only the coupling constant of M. elsdenii flavodoxin is not in line with expectation, this must most probably be ascribed to the lower accuracy with which the coupling constant could be determined at that time.8 The data also support the view that N(3) in all flavodoxins is highly sp2 hybridized. This conclusion seems in contradiction with the values of the

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573

coupling constants, but it must be considered that the coupling constants is a scalar term influenced by the N-H bond length (see above). At any rate the observation of a coupling constant in protein-bound flavin proves its inaccessibility to bulk solvent. Free FMN in an aqueous solution does not show the N(3)H coupling constant due to a fast proton exchange reaction. The 15N chemical shifts due to N(5) of FMN in flavodoxins indicate that this atom is not hydrogen bonded in M. elsdenii and C. MP flavodoxins, weakly hydrogen bonded in A. vinelandii and D. vulgaris flavodoxins and strongly in Anabaena 7120 flavodoxin. Since the N(5) atom of flavin is involved in the redox reactions, it could have been expected that a correlation between the 15N chemical shifts due to N(5) and the redox potentials of flavodoxins would exist. However, no correlation is apparent yet.17 The N(10) atoms in flavodoxins are all sp2 hybridized except that in A. vinelandii flavodoxin showing a considerable degree of sp3 hybridization (Table 2). Like in flavodoxins, all other oxidized flavoproteins listed in Table 2 possess a hydrogen bond at the N(l) atom. The strongest hydrogen bond is observed in bacterial luciferase,26 the weakest in glucose oxidase.45 It has been suggested that a helical microdipole is located close to N(l) in bacterial luciferase26 and /7-hydroxybenzoate hydroxylase,35 for the latter this suggestion is supported by X-ray data and by the fact that an additional upfield is observed in the enzyme-substrate and enzyme-product complexes. It is known that the substrate induces a conformational change in the enzyme thereby facilitating the reduction of the enzyme by NADPH. 1J[15N(3)-1H(3)] are observed in all flavoproteins indicating hydrogen bonding with N(3)H and inaccessibility of this group to bulk solvent. The value of the coupling constants (Table 3) indicate sp2 hybridization of the N(3) atom. In phydroxybenzoate hydroxylase the coupling constant is not resolved due to the large line width. In riboflavin-binding protein the coupling is not observable due to rapid proton exchange.46 This observation is in agreement with published data showing accessibility of N(3)H to bulk solvent. What concerns the N(5) atom is that all flavoproteins have a hydrogen bond comparable to the one in flavodoxins, except those in bacterial luciferase, Old Yellow Enzyme, andp-hydroxybenzoate hydroxylase which are very strong. Furthermore, addition of substrate to p-hydroxybenzoate hydroxylase abolishes the hydrogen bond at N(5), as expected, while in the 6-hydroxy-L-nicotine oxidase-inhibitor complex a stronger hydrogen bond is observed than in the free enzyme. This hydrogen bond is formed between N(5) of the enzyme and the inhibitor. The N(10) atom of the above flavoproteins is sp2 hybridized like in flavodoxins. A decreased sp2 hybridization is observed in bacterial luciferase,26 Old Yellow Enzyme,47 and in 6-hydroxy-L-nicotine oxidase-inhibitor complex,33 the degree of sp3 hybridization is similar with that observed in A. vinelandii flavodoxin17 (Table 2). In all reduced flavoproteins the N(l) atom exhibits 15N chemical shifts very similar to the ones of ionized reduced free flavin is aqueous, alkaline solutions (Table 2). This fact proves that all reduced flavoproteins contain an ionized prosthetic group. This observation is corroborated by 13C NMR (see below). Protonation of N(l) does not occur at pH values as low as about 5, indicating that the ionization constant of protein-bound flavin is decreased as compared with that of free flavin. It should, however, be noted that the degree of ionization, respectively, the degree of hydrogen bonding with N(l) varies among the proteins. The difference was explained either by hydrogen bonding or the interaction of the negatively charged N(l) atom with positively charged amino acid residues. The only exception is observed in riboflavin-binding protein which exhibits a pKa value of 7.5 in agreement with the greater accessibility of the pyrimidine subnucleus of the prosthetic group.46 All reduced flavoproteins, except riboflavin-binding protein forming a hydrogen bond with bulk solvent, possess a hydrogen bond between N(3)H and the protein. The strongest hydrogen bonds are formed in Old Yellow Enzyme and Anabaena 7120 flavodoxin, the

574

Chemistry and Biochemistry of Flavoenzymes

weakest in D. vulgaris flavodoxin, substrate-reduced 6-hydroxy-L-nicotine oxidase and free /?-hydroxybenzoate hydroxylase (Table 2). The coupling constants of the N(3)H group, as far as determined and observable, indicate predominantly sp2 hybridization of the N(3) atom and inaccessibility of the N(3)H group to bulk solvent (Table 3). The majority of the 15N chemical shifts due to the N(5) atom in reduced flavoproteins range from about 58 ppm to about 62 ppm indicating hydrogen bonding interactions, the strongest hydrogen bonding occurring in D. vulgaris flavodoxin (Table 2). Chemical shifts appearing upfield from that of reduced FMN in aqueous solution are observed in Old Yellow Enzyme, Anabaena 7120 flavodoxin, dithionite-reduced 6-hydroxy-L-nicotine oxidase, phydroxybenzoate hydroxylase in the presence of product, and glucose oxidase at pH 7.4. Since the upfield shifts in Old Yellow Enzyme, dithionite-reduced 6-hydroxy-L-nicotine oxidase and glucose oxidase at pH 7.4 correspond with a decreased absorption at 450 nm, it was concluded that the N(5) atom in these proteins possesses a considerable degree of sp3 hybridization; i.e., the atoms are moved out of the molecular plane. It is not yet clear if a similar situation is present in Anabaena 7120 flavodoxin, the determination of the coupling constant could clarify this point. The above presented interpretations are supported by 1 15 J[ N(5)-1H(5)] (Table 3). The observation of a N(5)H coupling constant in most reduced flavoproteins has been interpreted in the past in terms of inaccessibility of the functional group to bulk solvent. Recent data have however shown that the N(5) proton is not exchanging in even free flavin in aqueous solution and is therefore an inherent property of reduced flavin, irrespective of free or bound to proteins. Hence the earlier conclusions are no longer valid. The N(10) atom in reduced flavoproteins is highly sp2 hybridized except for bacterial luciferase. This is also true for glucose oxidase, but to a lesser degree. For free flavins in the reduced state, an equation was developed for the calculation of the endocyclic angles allowing to evaluate the conformation of the molecule.4 Due to the complex interaction between the prosthetic group and apoflavoprotein the equation cannot be used to obtain information about the conformation of protein-bound reduced flavin.

IV.

13

C NMR

As mentioned in the introduction there exists a linear relationship between the 13C chemical shift and the TT electron density on a particular carbon atom. This allows to detect subtile submolecular differences in the flavin molecule when exposed to different environments, i.e., solvents or apoflavoproteins. The detection and assignment of a particular resonance in a 13C NMR spectrum, even if the prosthetic group is selectively enriched, can be more difficult than in 15N NMR spectra. As the size of a protein increases the natural abundance 13C resonances can become predominant masking the specific resonance. This problem can, however, be overcome as demonstrated in Figure 7 by difference spectroscopy. The chemical shifts collected in Table 4 demonstrate that hydrogen bonding interaction with O(2a) and O(4a) of free oxidized flavin leads to a downfield shift of the corresponding resonances. This holds also for the reduced molecule where in addition the ionization state of N(l) is reported by the chemical shifts due to C(2) and C(10a). Based on this information the 13C NMR spectra of oxidized and reduced flavoproteins were interpreted with regard to the hydrogen binding interaction of the prosthetic group with the apoprotein. This interaction is confined to the pyrimidine subnucleus of flavin. This part of the flavin molecule, though rather polar, is buried in the protein interior and not the xylene subnucleus as one would have expected based on its apolar character. So far this is true for all flavoproteins except riboflavin-binding protein which is a carrier protein with no apparent catalytic purpose. The hydrogen bond formation to protein-bound flavin serves two purposes: (1) to support proper binding of the flavin, and (2) to tune probably also the flavin for its specific catalytic mechanism.

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575

FIGURE 7. 13C NMR spectra of various flavoproteins in the oxidized and reduced state. (A) D. vulgaris flavodoxin; (B) riboflavinbinding protein from egg white; (C) Old Yellow Enzyme; (D) bacterial luciferase; (E) glucose oxidase.

The 13C and 15N chemical shifts are complementary and allow for a detailed structural interpretation only when the two sets of data are combined. Nevertheless the 13C chemical shifts will not be discussed here in great detail but only the major findings. All oxidized flavodoxins form hydrogen bonds with the O(2a) and O(4a) atoms, the hydrogen bond to O(2a) being stronger than that to O(4a). The weakest hydrogen bond is observed in A. vinelandii flavodoxin to the O(2a) atom.

576

Chemistry and Biochemistry of Flavoenzymes TABLE 4 C NMR Chemical Shifts of Free and Protein-Bound Flavins

13

Chemical shift (ppm)a Compound

pH

C(2)

C(4)

C(4a)

C(10a)

Ref.

Oxidized TARF in CHC13 FMN FAD Flavodoxin from Megasphaera elsdenii Clostridium MP Azotobacter vinelandii Desulfovibrio vulgaris Anabaena 7120 Bacterial luciferase Old Yellow Enzyme + Ligand Glucose oxidase 6-Hydroxy-L-nicotine oxidase Free enzyme + Inhibitor p-Hydroxybenzoate hydroxylase Free enzyme + Substrate Riboflavin-binding protein from Egg yolk Egg white Egg white D-Amino acid oxidase Salicylate hydroxylase Free enzyme + Substrate Lipoamide dehydrogenase Glutathione reductase Mercuric ion reductase Butyryl-CoA dehydrogenase

7.0 8.0

155.2 159.8 161.0

159.8 163.7 164.8

135.6 136.2 137.3

149.1 152.1 153.2

26 26 51

7.0 8.0 8.0 8.0 8.0 8.0 7.5 7.0 8.5 8.0 8.0 5.8

159.4 159.8 159.8 159.3 159.6 159.7 158.6 158.5 160.6 162.1 160.2 159.8

162.6 162.4 162.3 161.7 161.7 162.4 161.6 162.6 164.2 165.8 163.5 162.5

135.4 135.6 135.5 135.3 135.7 134.3 135.3 137.4 137.1 139.0 133.8 137.8

153.1 151.5 151.4 154.8 153.5 152.3 152.2 151.3 152.9 154.7 153.1 152.5

10 17 17 10 17 43 44 26 28 7 28 45

7.0 7.0

159.9 160.0

164.9 164.8

136.9 138.3

151.5 151.7

33 33

7.0 7.0

159.5 156.6

163.2 162.8

136.4 135.7

151.6 151.1

35 35

8.5 8.5 7.0 8.0

159.6 159.7 161.2 160.7

162.0 162.1 163.6 165.0

135.9 135.9 137.6 137.3

152.4 152.3 154.0 152.5

46 46 9 51

7.0 7.0 7.0 7.0 7.0 7.0

160.2 159.5 158.3 160.4 159.6 158.2

164.2 161.4 163.2 163.3 163.6

151.4 150.3 152.2 152.3 152.5

—

136.0 134.4 137.6 135.7 137.5 138.7

—

35 35 52 52 52 52

Reduced TARF in CHC13 FMNH2 FMNHFlavodoxin from Megasphaera elsdenii Clostridium MP Azotobacter vinelandii Desulfovibrio vulgaris Bacterial luciferase Old Yellow Enzyme Glucose oxidase 6-Hydroxy-L-nicotine oxidase Dithionite-reduced Substrate-reduced

5.0 8.5

150.6 151.1 158.2

157.0 158.3 157.7

105.2 102.8 101.4

137.1 149.7 155.5

26 26 26

7.8 8.0 8.0 8.0 8.0 8.0 7.0 8.0 7.8

156.5 156.9 156.6 158.1 158.3 157.5 157.9 159.4 159.0

154.9 154.8 154.8 154.9 155.2 154.0 157.2 163.0 160.4

103.1 103.5 103.7 102.2 102.6 102.7 103.5 95.3 97.4

154.2 154.5 154.1 154.9 155.2 155.0 156.2 157.7 158.3

10 17 17 10 17 43 26 28 45

7.0 7.0

158.7 158.2

160.0 157.1

98.1 99.3

160.0 156.1

33 33

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577

TABLE 4 (continued) C NMR Chemical Shifts of Free and Protein-Bound Flavins Chemical shift (ppm)a Compound

pH

C(2)

C(4)

C(4a)

C(10a)

Ref.

Reduced p-Hydroxybenzoate hydroxylase Free enzyme + Substrate Riboflavin-binding protein from Egg white Salicylate hydroxylase Free enzyme + Substrate Lipoamide dihydrogenase EH2b EH4C Glutathione reductase EH2b EH4C Mercuric ion reductase EH2b EH4C Butyryl-CoA dehydrogenase a

b c

7.0 7.0

156.9 156.9

157.6 158.1

99.9 99.3

155.6 153.3

35 35

6.0 9.0

151.1 158.1

156.5 156.8

102.8 102.8

143.9 155.0

46 46

7.0 7.0

158.1 158.4

160.1 159.9

98.1 99.7

154.5 151.7

35 35

7.0 7.0

158.6 159.4

163.8 159.7

140.3 98.7

153.0 157.2

52 52

7.0 7.0

161.6 161.0

163.9 162.0

139.9 99.7

153.8 159.1

52 52

7.0 7,0 7.0

160.1 160.1 158.3

164.5 161.8 —

140.7 98.9 104.4

153.8 158.6 —

52 52 52

Chemical shifts are reported relative to TMS. The values of Miura et al.51 are the original ones obtained using a slightly different reference compound. Two-electron reduced form. Four-electron reduced form.

As already pointed out above, the N(10) atom in this flavodoxin possesses a considerable degree of sp3 hybridization; i.e., TT electrons from this atom are only partially delocalized onto O(4a) not allowing for a strong hydrogen bond to be formed with this position. Also Anabaena 7120 flavodoxin forms a weak hydrogen bond with O(4a), the 15N chemical shift of N(10) is identical with that of free FMN. Both values are however lower than those observed in flavodoxins from M. elsdenii, C. MP, and D. vulgaris. Therefore it could be concluded that also Anabaena 7120 flavodoxin possesses a N(10) with a decreased sp2 hybridization, though to a lower extent than that in A. vinelandii flavodoxin. This suggests that the redox properties of the two proteins are very similar. Indeed when the redox potentials of the flavodoxins are compared48 then an apparent relationship seems to exist between the degree of sp3 hybridization of the N(10) atom in oxidized flavodoxin and the El redox potential yielding the following order of increasing El potentials (more positive): A. vinelandii < Anabaena 7120 < M. elsdenii — D. vulgaris flavodoxins. It is possible that this relationship will become more obvious when the redox potentials have been redetermined to a greater accuracy. In the reduced state the flavodoxins show a rather uniform hydrogen bonding pattern as deduced from the 13C chemical shifts; i.e., hydrogen bond to O(2a), no hydrogen bond to O(4a). In addition all reduced flavodoxins possess a planar or almost planar structure. The 13C chemical shifts due to the carbon atoms of the xylene subnucleus of flavin are given in Table 5. The 13C chemical shifts of the C(6) and C(8) atoms demonstrate the polarization of the protein-bound flavin in the oxidized state whereas the 13C chemical shifts of the reduced flavodoxins show the increased IT electron density in the xylene subnucleus of the prosthetic group as compared to that of reduced free FMN.

578

Chemistry and Biochemistry of Flavoenzymes TABLE 5 C NMR Chemical Shifts of Free and Protein-Bound Flavins: Xylene Subnucleus

13

Chemical shift (ppm)a Compound

C(5a)

C(6)

C(7)

C(7a)

C(8)

C(8a)

C(9)

C(9a)

Ref.

Oxidized TARF FMN Flavodoxin from A. vinelandii D. vulgaris M, elsdenii C. MP Anabaena 7120 Bacterial luciferase Old Yellow Enzyme Plus ligand 6-Hydroxy-L-nicotine oxidase Free enzyme + Inhibitor

134,6 136.4

132.8 131.8

136.6 140.4

19.4 19.9

147.5 151.7

21.4 22.2

115.5 118.3

131.2 133.5

4 26

136.9 137.4 138.4 138.5 135.7 135.7 135.2 134.1

132.6 132.5 133.0 133.1 129.8 130.8 — —

141.2 142.0 141.4 141.4 141.9 139.0 141.2 139.4

20.4 20.5 20.6 20.5 20.3 20.2 19.3 19.1

152.2 154.0 153.0 153.0 152.8 148.6 151.8 150.1

21.9 23.3 22.1 22.0 23.0 21.9 21.9 21.9

117.9 117.2 117.5 117.4 117.8 119.5 117.7 116.8

131.5 131.9 132.9 132.9 130.8 134.6 131.7 131.3

17 43 17 17 44 26 28 28

136.9 136.3

131.6 133.0

137.7 137.4

20.7 20.8

148.3 148.1

21.6 21.5

120.3 120.0

135.5 135.2

33 33

Reduced TARFH2 FMNH2 FMNHFlavodoxin from A. vinelandii D. vulgaris M. elsdenii C. MP Bacterial luciferase Old Yellow Enzyme 6-Hydroxy-L-nicotine oxidase Dithionite-reduced Substrate-reduced a

136.0 134.4 134.2

116.1 117.1 117.3

133.6 134.3 133.0

18.9 19.0 19.0

129.0 130.4 130.3

18.9 19.2 19.4

118.0 117.4 116.8

128.2 130.4 130.9

4 26 26

135.5 134.6 136.3 136.5 135.0 133.9

113.8 114.5 112.4 112.4 116.8 —

130.4 130.7 131.1 131.3 132.7 131.8

19.8 19.4 19.6 19.6 19.4 18.3

125.5 126.7 125.9 125.6 126.2 128.5

19.3 20.3 19.2 19.1 19.7 19.5

115.2 114.7 115.3 114.8 115.6 117.0

131.2 129.1 131.8 132.1 130.7 131.8

17 43 17 17 26 28

135.2 134.8

120.8 117.8

134.5 131.8

19.9 20.5

129.7 128.5

21.4 21.4

117.9 116.9

130.1 130.6

33 33

Chemical shifts are reported relative to TMS.

One-bond 13C-13C coupling constants of the pyrimidine moiety of flavin are given in Table 6. The coupling constants observed in flavodoxins resemble more those of FMN in water than those of TARF in chloroform. These coupling constants support the proposed hydrogen bonding pattern and can also be used to estimate the s-character of the atoms making up the bond. Additional C-C and C-H coupling constants were determined in Anabaena 7120 and D. vulgaris flavodoxins.43'44 These data yielded information on the internal mobility of the bound flavin17 and the structure of flavin which shows some distortion in the C(8a), C(8), C(9) region in the protein44 as also observed in free flavin.2'49 Bacterial luciferase forms hydrogen bonds to both O(2a) and O(4a), albeit weaker than those of FMN in water (Table 4). However the C(8) and C(7) resonances indicate that the prosthetic group is not as strongly polarized as free FMN in aqueous solution (Table 5). In the reduced state a hydrogen bond is observed to O(2a) only. Using 13C NMR, it was observed that bacterial luciferase binds excess reduced flavin, probably in an aspecific manner, suggesting that bacterial luciferase has two different binding sites for reduced flavin. In the oxidized state a stoichiometric amount of flavin was bound if a highly active preparation was used. Thus 13C NMR can be used to monitor specific and aspecific interactions between

Volume III

579

TABLE 6 Some Direct 13C-13C Coupling Constants of Free and Protein-Bound Flavins in the Oxidized and Reduced States Coupling constants (Hz) Compound

*J["C(4)-13C(4a)]

1

J[13C(4a)-13C(10a)]

Ref.

76.5 75.5 75.4

53.3 53.3 55.9

10 17 10, 17

76.5 76.5 76.3 76.9 76.3 74.9

58.8 56.5 56.2 57.3 56.5 56.9

10 17 17 43 17 44

79.1 82.0 84.0

84.5 81.6 74.4

17 17 17

85.2 85.5 86.7 87.9

73.2 73.2 72.8 72.0

17 17 43 17

Oxidized TARF FMN

Flavodoxin from M. elsdenii C. MP D. vulgaris A. vinelandii Anabaena 7120

Reduced TARFH2 FMNH2 FMNH" Flavodoxin from M. elsdenii C. MP D. vulgaris A. vinelandii

an apoflavoprotein and the prosthetic group, e.g., partially unfolded conformation or other alterations influencing the proper interaction of flavin with the apoprotein. The structure of the catalytically important intermediate in the bacterial luciferase reaction, the C(4a)-hydroperoxyflavin, was elucidated by 13C NMR. 50 This was possible because the intermediate has a rather long lifetime. Old Yellow Enzyme forms in the oxidized state a hydrogen bond, comparable to that of FMN, with O(2a) and a rather strong one with O(4a) (Table 4). In the reduced state strong hydrogen bonds are formed with both carbonyl groups. The resonance of the C(4a) atom is considerably upfield shifted due to various effects.28 On complexation with phenolic compounds the chemical shifts of the N(5) and N(10) atoms are upfield shifted in a parallel manner, but not to the same extent, the largest upfield shift amounts to 14.3 ppm [N(5)] in the presence of p-chlorophenol.28 The N(3) atom is not affected very much and N(l) shows a peculiar behavior in that a downfield shift is observed which does not parallel the above mentioned upfield shifts of the N(5) and N(10) atoms. On complexation an increasing 15N chemical shift of N(l) is observed in the presence of p-nitrophenol, p-hydroxybenzaldehyde and w-nitrophenol; and 15N chemical shift decreased then in the presence of p-hydroxy-AT(rt-butyl)benzamide and /?-chlorophenol. As compared with the 13C chemical shifts of the free enzyme all resonances of the xylene moiety of protein-bound FMN are upfield shifted indicating acquisition of IT electron density upon complexation. The largest upfield shifts are however observed for C(9), C(7) and C(5a) (Table 5).28 These atoms are under the influence of N(10) indicating that N(10) releases some of its negative charge, acquired on complexation, onto these three carbon atoms. This explains the observation that the upfield

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Chemistry and Biochemistry of Flavoenzymes

shift of N(10) is less than that of N(5). Using the 15N chemical shifts no linear correlation between the chemical shifts and the pKa values of the phenols could be found for both atoms.28 Plotting the difference chemical shifts between free and complexed protein against the pKa values of the phenolic compounds gives a fair correlation for the N(10) atom but not for the N(5) atom. No correlation could be found when the 15N difference chemical shifts were plotted against the Hammatt para substituent constants (ap) of the phenolic compounds. On the other hand a linear correlation was demonstrated between the difference chemical shifts of the C(4a) atom and the crp value of the phenolic compounds.7 The discrepancy between the two sets of data is not obvious but reflects the high complexity of the system. It was also demonstrated that the phenolic compound is bound in the ionized form to the enzyme28 and that the different isoenzymes of Old Yellow Enzyme perturb the flavin binding differently.7-28 The resonances of the C(l) and C(2,6) atoms of /?~nitrophenol are shifted upfield upon complexation, indicating loss of IT electron density in the benzene ring of the phenolate. Furthermore C(2) and C(6) of the phenolate become magnetically inequivalent in the protein complex, indicating very tight binding of the phenolate at an asymmetric binding site of the protein.28 Glucose oxidase exhibits a strong hydrogen bond with O(2ot) and a weaker one with O(4a) in the oxidized state. In the reduced state the hydrogen bond with O(4a) is increased in strength.45 In 6-hydroxy-L-nicotine oxidase the xylene moiety of the prosthetic group is embedded in a hydrophobic environment as deduced from the chemical shifts of C(7) and C(8). The polarization of the isoalloxazine ring as a whole is, however, much more comparable to that of free flavin in a polar and protic environment. The polarization can be ascribed to hydrogen bonding interactions with the two carbonyl groups (Tables 4, 5).33 Binding of the competitive inhibitor, 6-hydroxy-D-nicotine, influences strongly the resonance position of N(5) and C(4a), suggesting a hydrogen bond formation between N(5) of flavin and the hydroxyl group of the inhibitor. In the reduced state the N(5) atom is fully sp2 hybridized (Table 2). This leads to a redistribution of electron density from N(5) onto C(6) and C(8) (Table 5). p-Hydroxybenzoate hydroxylase, riboflavin-binding protein, D-amino acid oxidase, salicylate hydroxylase, lipoamide dehydrogenase, glutathione reductase, mercuric ion reductase, and butyryl-CoA dehydrogenase have been studied in less detail than the above discussed flavoproteins (Table 4). Only the most salient findings will, therefore, be mentioned here. p-Hydroxybenzoate hydroxylase and salicylate hydroxylase exhibit hydrogen bonds to O(2ot), their strength is comparable to that observed in FMN. The same holds for the carbonyl group in position 4, the hydrogen bond in salicylate hydroxylase being stronger than that in p-hydroxybenzoate hydroxylase. Addition of substrate leads to the disruption of the hydrogen bond at O(2a) in both enzymes, while the hydrogen bond at O(4a) remains in place in p-hydroxybenzoate hydroxylase and is lost in salicylate hydroxylase (Table 4). An interesting result on /?-hydroxybenzoate hydroxylase is the observation that the active site structure of the enzyme-substrate complex in the oxidized state is also attained in the free, two-electron reduced state implying that reduction leads to about the same conformational change as binding of substrate to the oxidized enzyme.35 D-amino acid oxidase shows a weak hydrogen bonding to the carbonyl group at position 2 and a stronger one to the other carbonyl group. In the presence of inhibitors the hydrogen bond at O(4a) is lost and the hydrogen bond at O(2a) is strengthened. In these complexes the resonances of C(10a) and C(4a) are upfield shifted indicating an increase of TT electron density at these positions.51 However, no clear trend is visible for these complexes which often are referred to as "charge-transfer" complexes. However, an obvious difference exists between these complexes and those of Old Yellow Enzyme. The 13C chemical shifts of the remaining four flavoproteins listed in Table 4 indicate that the interactions between the apoflavoproteins and the prosthetic groups are very similar

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in the oxidized state. In the two-electron (EH2) and four-electron (EH4) reduced states some differences become apparent, notably in lipoamide dehydrogenase. A more detailed characterization of these flavoproteins by NMR techniques is under way. 52 Flavodoxin from M. elsdenii forms a red colored product if irradiated in the presence of an organic acid like acetic acid. This product has been studied by 13C and 15N NMR.52 The preliminary data indicate that the 13C and 15N chemical shift resemble much more closely those of reduced than oxidized flavodoxin, except for the 15N chemical shift of N(5) which is similar with that of oxidized flavodoxin. The NMR data indicate that a residue of the acid is attached to C(8) of flavodoxin leading to a 1,8-dihydroflavin structure.52 The 13C NMR technique has also been used to study the environment of the two thiol groups of M. elsdenii flavodoxin upon cyanylation with I3 CN~. 53 Preliminary data indicate that the thiol groups are in a different environment and that the environment of one SH group was changed when the flavin was removed from the protein. The reaction is reversible.53 In still another application of the 13C NMR spectroscopy the internal mobility of FMN bound to the apoflavodoxin from M. elsdenii was investigated by relaxation studies.54 It was shown that the protein-bound flavin possesses very little, if any at all, mobility on the nanosecond time scale.

V. 19F NMR Fluoride is an interesting nucleus for NMR studies because of the high natural abundance of 19F and its high sensitivity which is close to that of J H. 19F NMR has, however, only been used in very few cases. The synthesis of a flavin derivative possessing a fluoro instead of the methyl group in position 8 of the molecule has been described.55 In the same paper, a brief description of the 19F NMR spectrum was given and stated that the chemical shift of the flavin derivative appears at 57.80 ppm and is close to that of dinitrofluorobenzene (53.21 ppm), indicating similar deshieldings. In using fluorosubstituted molecules in 19F NMR it should be considered that fluorine is strongly electron attracting and forms easily hydrogen bonds. When the fluoroflavin was bound to riboflavin-binding protein a small upfield shift, as compared to the free molecule, was observed in the 19F NMR spectrum.56 The pH-dependence of the 19F NMR spectra of the free and protein-bound oxidized flavin revealed a pKa of 9.6 and about 10.6 for the N(3)H group, respectively. The pKa value for the free flavin is lowered by about one pH unit as compared to riboflavin. The chemical shift of the fluor atom is upfield shifted upon deprotonation of N(3)H suggesting charge redistribution from the pyrimidine onto the xylene moiety of flavin. It is interesting to note that the 13C chemical shift of C(8) of the fluoro-containing flavin has a value of 162.6 ppm. This value is more than 10 ppm downfield from that of FMN (see Table 5). This is according to expectation showing an increased partial positive charge at C(8) as compared to FMN. Therefore this increased positive charge is probably responsible for the decreased pKa value. It would be desirable to study the fluoro compound by 13C and 15N NMR to obtain more information about the TT electron density distribution in this molecule. The increased pKa of N(3)H of 8-fluoroflavin bound to the protein could, as an alternative interpretation, be due to hydrogen bonding, most likely with the protein. Macheroux et al.57 studied a number of flavoproteins containing 8-fluoroflavin. It is interesting and admirable with which great detail the spectra were interpreted, based on one single resonance. Disturbing is the fact that the published 13C and 15N NMR data by Moonen et al.4 were misread. The proteins were studied in the oxidized and reduced states in the presence and absence of ligands. The 19F chemical shifts were in the range of about 60 to 70 ppm for the oxidized proteins, and about 55 to 65 ppm for the reduced proteins.57

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Chemistry and Biochemistry of Flavoenzymes TABLE 7 F Chemical Shifts of Free and Protein-Bound 8-Fluoroflavins and Topology at Position 8

19

19

F Chemical shifts (ppm)

Compound FAD Riboflavin pH 8.5 Methanol M. elsdenii flavodoxin D-Amino acid oxidase + Benzoate p-Hydroxybenzoate Hydroxylase + Substrate Old Yellow Enzyme Riboflavin-binding protein Glucose oxidase Lactate oxidase General acyl-CoA dehydrogenase + Acetoacetyl-CoA a b c

Rate constant (min ')

Oxidized Reduced Thiophenol*

Iodoacetamideb Topology

65.6

35.7

1.84

0.46

64.9 60.0 66.1 59.3 62.5

36.0 33.3 28.5 30.9 —

—

—

— 0.0039 2.65 0

— 0.49 tV 2 ~6 h tV 2 ~6 h

65.8 67.4 65.7 64.7 69.4 63.7 68.1 66.5

32.3 31.2 38.1 33.7 38.5 35.7 36.9 —

2.24 0.47 20.8 0.0035 0 tV 2 > 1 day nd< —

1.2

-6 0.80 — tV2~12h tV 2 ~1 wk — —

Ref. 57, 60

— — Exposed Exposed Exposed

57 57 57, 58, 60 57, 58, 60 57, 58

Exposed Exposed Exposed Buried Buried Buried Buried Buried

57, 57, 57, 57, 57, 57, 57, 57,

58, 58 58, 58, 58, 58, 58 58

60 60 60 60 60

The compound used was 8-chloroflavin. The compound used was 8-mercaptoflavin. nd denotes not determined.

It is interesting to note that the line widths of the 19F NMR spectra of D-amino acid oxidase, p-hydroxybenzoate hydroxylase, general acyl-CoA dehydrogenase and lactate oxidase vary greatly. The line widths are dependent on the presence or absence of ligands. With the free enzymes no correlation is observed between the line width and the molecular weight of the protein, in contrast to the claims of the authors.57 Confusing, at least at this time, is the fact that addition of ligand causes an increase in line width in some cases and a decrease in other cases. 19 F Chemical shifts are sensitive to hydrogen bonding and microenvironment (e.g., dispersion forces from neighboring substituents, particularly if they are bulky). Therefore 8-fluoroflavin bound to apoflavoproteins should be a valuable tool to probe the accessibility of position 8 in such reconstituted flavoproteins. 8-Chloroflavin has been incorporated into a series of apoflavoproteins and the accessibility of the 8 position monitored by UV spectroscopy and nucleophilic substitution reactions.58 These data are not always easily to interpret and can lead to incorrect conclusions; it has been concluded, e.g., that flavin in reduced flavodoxin is not ionized,59 a conclusion in contradiction with 15N NMR data (see above). In addition 13C and 15N NMR data show that binding of flavin to apoflavoproteins perturbs the electronic structure of the molecule in a unique and complex manner (see Tables 2 , 4 , 5). Using flavoproteins reconstituted with modified flavins in which the substituent perturbs the electronic structure of the flavin already in the free state adds further complexity to the interpretation of the data. Therefore a detailed interpretation of 19F NMR spectra of 8-fluoro flavin-containing proteins in terms of electronic structure is not feasible without supporting 13 C and 15N NMR data. However, the 19F NMR data of the flavoproteins studied57 should in particular yield some information regarding the accessibility of position 8 in proteinbound 8-fluoroflavin. In Table 7, the I9F chemical shifts57 and the topology of the position 8 of protein-bound flavin as deduced from kinetic and spectroscopic studies are presented.58 The pseudo-first order rate constants for the reaction of thiophenol with protein-bound

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8-chloroflavin, and of iodoacetamide with protein-bound 8-mercaptoflavin are also given in Table 7.60 It is reasonable to compare the protein-bound 8-chloro- with the 8-fluoroflavin because their structural perturbation is about the same for both proteins. There are different ways to look at the data of Table 7. If the 19F chemical shifts are interpreted according to Macheroux et al.57 then one would have to conclude that the more downfield shifted the chemical shift of a particular flavoprotein appears the more the flavin ring would be polarized. Due to the electronegativity of the fluorine atom the polarization would lead to a higher polarization of the C(8)-F bond and consequently to a higher reactivity of the functional group, if accessible. For flavodoxin and Old Yellow Enzyme, for which 13C chemical shifts are available for the whole molecule (Tables 4, 5), the degree of polarization at the C(8) atom can be roughly estimated. These data indicate that flavodoxin is more polarized than Old Yellow Enzyme. This is in accord with the 19F chemical shifts of the two proteins. The large downfield shift of 19F resonances of both proteins as compared to that of free flavin in methanol indicates strong hydrogen bond formation with the fluorine atom. Thus the C(8) atom in these proteins seems to be accessible to bulk solvent and their reactivity towards nucleophiles is expected to be about the same. This is neither reflected in the rate constant of the reaction with thiophenol nor with iodoacetamide, although some other factors like charges on the reactant and the protein, and the bulkiness of the reactant must be considered. Riboflavin-binding protein, whose 19F chemical shift resembles that of Old Yellow Enzyme, exhibits about the same reactivity towards thiophenol as flavodoxin (Table 7), although the xylene ring of flavin is buried in the interior of riboflavin-binding protein. From this point of view it is very difficult to consolidate the data of Table 7. The best way to unravel this problem would be to elucidate the topology of flavoproteins around the C(8) position in more detail by 19F NMR spectra taken in H2O and D2O.61 This approach would also allow to separate the effects of hydrogen bounding with the fluorine atom from electronic effects induced on the flavin by the apoprotein.

VL 17O NMR Two 17O labeled flavin derivatives were synthesized:62 [295% isotopic purity) from Anabaena 7120. More than 10% of the expected N-H cross peaks could be resolved. The interaction between the apoprotein and the prosthetic group have already been discussed above. A few 15N resonances could be assigned to amino acid residues.23 From sequence specific ]H and 13C NMR assignments the (3-sheet structure was determined and found to ue very similar to that observed in the related flavodoxin from Anacystis nidulans by X-ray crystallography.21 The five parallel (3-sheet strands forming the p-sheet show the same connectivity as found in M. elsdenii flavodoxin; i.e., 2-1-3-4-5. A NOESY distance-constrained energy-minimized structure of the FMN binding site in oxidized Anabaena 7120 flavodoxin was calculated.44 The electronic environment of the proteinbound flavin was elucidated by 13C and 15N chemical shifts. The flavin was found to be polarized due to hydrogen bond formation with the carbonyl groups, as observed in other flavodoxins studied. In addition it was demonstrated that the xylene ring of FMN is nonplanar to a certain degree.44 The above brief discussion demonstrates, although 2D-NMR has only been applied to flavoproteins during the past few years, that an impressive progress has been achieved. The fact that the structural elucidation of the larger of the two flavodoxins studied, Anabaena 7120 flavodoxin (21 kDa), is so well underway proves that the combination of !H, 13C and 15 N NMR spectroscopy facilitates the resolution of the structure of relatively large proteins. It can therefore be expected that the above studies will further stimulate the application of 2D-NMR spectroscopy in the field of flavoproteins.

VIII. CONCLUSION The data presented above demonstrate that very detailed information can be obtained by NMR techniques regarding the electronic structure, the ionization state, the conformation/ configuration, and the hydrogen bond interaction of protein-bound flavin. These results will ultimately yield the desired information concerning the principle governing the regulation of the redox potential in flavoproteins, when more data will become available in the near future. In addition the 2D- and 3D-NMR techniques will add further information as the three-dimensional structure of other flavoproteins will be resolved. These techniques will also be very helpful to elucidate the structure of modified flavoproteins and structural changes induced in flavoproteins by site-directed mutagenesis. The NMR program in the author's laboratory was undertaken to test the proposal that specific hydrogen bonding interactions between the prosthetic group and apoflavoprotein are determining the specific catalysis of a particular flavoprotein.5 This proposal was refined later by Hemmerich and Massey.93 In this respect it is surprising how similar the hydrogen bonding pattern is in the flavoproteins so far studied by NMR techniques. Beside other atoms, the N(3)H and C(2)O groups are always involved in hydrogen bonding. These two

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functional groups probably serve to bind the flavin properly. While hydrogen bonding with N(3)H does not perturb the electronic structure of flavin to a great extent, hydrogen bonding with C(2)O does lead to the polarization of the isoalloxazine ring system. Hydrogen bonding of course also contributes favorably to the binding of flavin to apoflavoproteins. The most remarkable information obtained by 13C and 15N NMR techniques is the observation that, except in riboflavin-binding protein, flavin in all two-electron reduced flavoproteins studied is ionized and that the degree of hybridization of the N(10) and N(5) atoms can be regulated independently by the apoflavoproteins. The fact that flavin is ionized was not easily accepted but was recently supported by X-ray data.94 The ionization of flavin in reduced flavoproteins and the observation that the phosphate group of FMN bound to apoflavodoxins is also ionized led to the proposal of Moonen et al.95 that charge-charge interactions in flavoproteins and especially in flavodoxins are a means to regulate the redox potential. In addition, it was also stated that factors like dipole-dipole, dipole-monopole, and monopole-monopole interactions, and the dielectric permittivity of the isoalloxazine binding site play an important role in the redox potential regulation. Calculations indeed showed that the redox potentials of a few flavodoxins could be predicted reasonably by the theory.95 A different model, still based on charge-charge interactions, was recently presented.94'96 In this proposal it was suggested that probably Glu-60 in M. elsdenii flavodoxin, forming a hydrogen bond with N(3)H of flavin, contributes mainly to the regulation of the redox potential in the protein. The redox potential of the semiquinone/dihydroflavodoxin couple is then explained by an increase of the pKa of Glu-60 by about 3 units as compared to that of the oxidized/semiquinone couple.94*96 Interestingly Gast et al.97 have proposed that the pKa value of 3.4, determined from the pH-dependence of the flavin-apoprotein interaction in M. elsdenii flavodoxin, is due to Glu-60/61. The value of 3.4 is in good agreement with that calculated by Ludwig et al., i.e., 3.O.94 This indeed could explain the observed pHdependence of the redox potential of the semiquinone/hydroquinone couple in M. elsdenii flavodoxin. The charge-charge repulsion between N(l) of flavin and Glu-60, suggested to weaken the flavin binding interaction,94'96 is however not reflected in the 15N NMR spectra (see above). On the other hand it could be expected that protonation of Glu-60 would also be crucial for the flavin-apoprotein interaction in reduced M. elsdenii flavodoxin as observed for the oxidized protein.97 In addition, the close proximity of the two charges in the protein would suggest to lead to a very unfavorable flavin-protein interaction. Further experiments will be required to support the present theories. Recently Hall et al.98"100 have presented theoretical data which, for the first time, consolidate most biophysical data available. These molecular orbital calculations are now fit to make important contributions to the above discussed problems.

ACKNOWLEDGMENTS I am indebted to Mr. D. Schmid for his help in the preparation of the manuscript. I also wish to thank my previous co-workers for their valuable contribution to my research program, especially Drs. C. T. W. Moonen, H. J. Grande, R. Gast, C. G. van Schagen, W. J. H. van Berkel, C. P. M. Van Mierlo, P. J. M. Bonants, and J. Vervoort. I also acknowledge the pleasant cooperation on some projects with the research groups of Profs. H. Riiterjans (Goethe-University, Frankfurt a.M.), A. Bacher (Technical University, Munich), D. E. Edmondson (Emory University, Atlanta), J. Lee (University of Georgia, Athens), K. Decker (Albert-Ludwigs University, Freiburg i. Br.), S. G. Mayhew (University College, Dublin), J. LeGall (University of Georgia, Athens), and R. Kaptein (University of Utrecht, Utrecht). I also acknowledge the general financial support of my research program by The Netherlands Organization for Scientific Research (NWO) and The Netherlands Foundation for Chemical Research (SON).

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REFERENCES 1. Muller, F., Free flavins: syntheses, chemical and physical properties, in Chemistry and Biochemistry of Flavoenzymes, Vol. 1, Muller, F., Ed., CRC Press, Boca Raton, FL, 1991, 1. 2. Grande, H. J., Cast, R., van Schagen, C. G., van Berkel, W. J. H., and Muller, F., 13C NMR study on isoalloxazine and alloxazine derivates, Helv. Chim. Acta, 60, 367, 1977. 3. van Schagen, C. G. and Muller, F., A comparative !3C NMR study on various reduced flavins, Helv. Chem. Acta, 63, 2187, 1980. 4. Moonen, C. T. W., Vervoort, J., and Muller, F., A reinvestigation of the structure of oxidized and reduced flavin: carbon-13 and nitrogen-15 nuclear magnetic resonance study, Biochemistry, 23, 4859, 1984. 5. Muller, F., On the interaction of flavins with phosphine derivatives, Z. Naturforsch., 27b, 1023, 1972. 6. Ghisla, S., Macheroux, P., Riiterjans, H., and Muller, F., lonization properties of fully reduced 1,5dihydroflavin, rates of N(5)-H exchange with solvent, in Flavins andFlavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 27. 7. Miura, R., Yamano, T., and Miyake, Y., 13P- and 31C-NMR studies on the flavoprotein and flavin-ligand interactions in brewer's Yeast Old Yellow Enzyme, J. Biochem. (Tokyo), 99, 907, 1986. 8. Fran ken, H.-D., Riiterjans, H., and Muller, F., Nuclear-magnetic-resonance investigation of 15N-labeled flavins, free and bound to Megasphaera elsdenii apoflavodoxin, Eur. J. Biochem., 138, 481, 1984. 9. Miura, R., Tojo, H., Fujii, S., Yamano, T., and Miyake, Y., A 13C-NMR study on the interaction of riboflavin with egg white riboflavin binding protein, J. Biochem. (Tokyo), 96, 197, 1984. 10. van Schagen, C. G. and Muller, F., A 13C nuclear-magnetic-resonance study on free flavins and Megasphaera elsdenii and Azetobacter vinelandii flavodoxin. 13C-Enriched flavins as probes for the study of flavoprotein active sites, Eur. J. Biochem., 120, 33, 1981. 11. Yagi, K., Ohishi, N., Takai, A., Kawano, K., and Kyogoku, Y., 15N nuclear magnetic resonance of flavins, Biochemistry, 15, 2877, 1976. 12. Kawano, K., Ohishi, N., Suzuki, A. T., Kyogoku, Y., and Yagi, K., Nitrogen-15 and carbon-13 nuclear magnetic resonance of reduced flavins. Comparative study with oxidized flavins, Biochemistry, 17, 3854, 1978. 13. Muller, F. and van Berkel, W. J. H., Methods used to resolve reversibly flavoproteins into the constituents apoflavoprotein and prosthetic group, in Chemistry and Biochemistry of Flavoenzymes, Vol. 1, Muller, F., Ed., CRC Press, Boca Raton, FL, 1991, 261. 14. Gorenstein, D. G. and Debojyote", K., 31P Chemical shifts in phosphate diester monoanions. Bond angle and torsional angle effects, Biochem. Biophys. Res. Commun., 65, 1073, 1975. 15. Gorenstein, D. G., Dependence of 31P chemical shifts on oxygen-phosphorus-oxygen bond angles in phosphate esters, J, Am. Chem. Soc., 97, 898, 1975. 16. Moonen, C. T. W. and Muller, F., Structural and dynamic information on the complex of Megasphaera elsdenii apoflavodoxin and riboflavin 5'-phosphate. A phosphorus-31 nuclear magnetic resonance study, Biochemistry, 21, 408, 1982. 17. Vervoort, J., Muller, F., May hew, S. G., van den Berg, W. A. M., Moonen, C. T. W., and Bacher, A., A comparative carbon-13, nitrogen-15, and phosphorus-31 nuclear magnetic resonance study on the flavodoxins from Clostridium MP, Megasphaera elsdenii, and Azotobacter vinelandii, Biochemistry, 25, 6789, 1986. 18. Edmondson, D. E. and James, T. L., Covalently bound non-coenzyme phosphorus residues in flavoproteins.31? NMR studies on Azotobacter flavodoxin, Proc. Natl. Acad. Sci. U.S.A., 76, 3786, 1979. 19. Edmondson, D. E. and James, T. L., Physical and chemical studies on the FMN and non-flavin phosphate residues in Azotobacter flavodoxin, in Flavins and Flavoproteins, Massey, V. and Williams, C. H., Jr., Eds., Elsevier, New York, 1982, 111. 20. Live, D. H. and Edmondson, D. E., Studies of phosphorylated sites in proteins using 1H-31P twodimensional NMR: further evidence for a phosphodiester link between a seryl and a threonyl residue h Azotobacter flavodoxin, J. Am. Chem. Soc., 110, 4468, 1988. 21. Ludwig, M. L. and Luschinsky, C. L., Structure and redox properties of clostridial flavodoxin, in Chemistry and Biochemistry of Flavoenzymes, Vol. 3, Muller, F., Ed., CRC Press, Boca Raton, FL, 1991, 000. 22. Favaudon, V., Le Gall, J., and Lhoste, J.-M., Nuclear magnetic resonance of flavodoxins from sulfatereducing bacteria, in Flavins and Flavoproteins, Yagi, K. and Yamano, T., Eds., University Park Press, Tokyo, 1980, 373. 23. Stockman, B. J., Westler, W. M., Mooberry, E. S., and Markley, J. L., Flavodoxin from Anabaena 7120: uniform nitrogen-15 enrichment and hydrogen-1, nitrogen-15 and phosphorus-31 NMR investigations of the flavin mononucleotide binding site in the reduced and oxidized states, Biochemistry, 27, 136, 1988. 24 Vervoort, J., van Berkel, W. J. H., Mayhew, S. G., Muller, F., Bacher, A., Nielsen, P., and Le Gall, J., Properties of the complexes of riboflavin 3',5'-bisphosphate and the apoflavodoxins from Megasphaera elsdenii and Desulfovibrio vulgaris, Eur. J. Biochem., 161, 749, 1986.

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25. Klugkist, J., Voorberg, J., Haaker, H., and Veeger, C., Characterization of three different flavodoxins from Azotobacter vinelandii, Eur. J. Biochem., 155, 33, 1986. 26. Vervoort, J., Muller, F., O'Kane, D. J., Lee, J., and Bacher, A., Bacterial luciferase: a carbon-13, nitrogen-15 and phosphorus-31 nuclear magnetic resonance investigation, Biochemistry, 25, 8067, 1986. 27. Mayhew, S. G. and Tollin, G., General properties of flavodoxins, in Chemistry and Biochemistry of Flavoproteins, Vol. 3, Muller, F., Ed., CRC Press, Boca Raton, FL, 1991,389. 28. Beinert, W.-D., Riiterjans, H., Muller, F., and Bacher, A., Nuclear magnetic resonance studies of the Old Yellow Enzyme. 2. 13C NMR of the enzyme recombined with 13C-labeled flavin mononucleotides, Eur. J. Biochem., 152, 581, 1985. 29. Otvos, J. D., Krum, D. P., and Siler Masters, B. S., Localization of the free radical on the flavin mononucleotide of the air-stable semiquinone state of NADPH-cytochrome P-450 reductase using 31P NMR spectroscopy, Biochemistry, 25, 7220, 1986. 30. Kainosho, M. and Kyogoku, Y., High-resolution proton and phosphorus nuclear magnetic resonance spectra of flavin-adenine dinucleotide and its conformation in aqueous solution, Biochemistry, 11, 741, 1972. 31. James, T. L., Edmondson, D. £., and Husain, M., Glucose oxidase contains a disubstituted phosphorus residue. Phosphorus-31 nuclear magnetic resonance studies of the flavin and nonflavin phosphate residues, Biochemistry, 20, 617, 198L 32. Nonaka, Y., Fujii, S., and Yamano, T., Phosphorus-31 nuclear magnetic resonance and electronic spectroscopic studies of adrenodoxin reductase and its binary complex with NADP + , J. Biochem. (Tokyo), 97, 1263, 1985. 33. Pust, S., Vervoort, J., Decker, K., Bacher, A., and Muller, F., 31C, 15N and 31P NMR studies on 6hydroxy-L-nicotine oxidase from Arthrobacter oxidans, Biochemistry, 28, 516, 1989. 34. Davis, M. D., Edmondson, D. E., and Muller, F., 31P nuclear magnetic resonance and chemical studies of the phosphorus residues in bovine milk xanthine oxidase, Eur. J. Biochem., 145, 237, 1984. 35. Vervoort, J., van Berkel, W. J. H., Muller, F., and Moonen, C. T. W., NMR studies on p-hydroxybenzoate hydroxylase from Pseudomonasfluorescens and salicylate hydroxylase from Pseudomonasputida, Eur. J. Biochem., in press. 36. Muller, F. and Vervoort, J., unpublished data. 37. Williams, C. H., Jr., Lipoamide dehydrogenase, glutathione reductase, thioredoxin reductase and mercuric ion reductase — family of flavoenzyme transhydrogenases, in Chemistry and Biochemistry ofFlavoenzymes, Vol. 3, Muller, F., Ed., CRC Press, Boca Raton, FL, 121, 1991. 38. Wierenga, R. K., Drenth, J., and Schulz, G. E., Comparison of the three-dimensional protein and nucleotide structure of the FAD-binding domain of p-hydroxybenzoate hydroxylase with the FAD, as well as NADPH-binding domains of glutathione reductase, J. Mol. BioL, 167, 725, 1983. 39. van Berkel, W. J. H., van den Berg, W. A. M., and Muller, F., Large-scale preparation and reconstitution of apo-flavoproteins with special reference to butyryl-CoA dehydrogenase from Megasphaera elsdenii. Hydrophobic-interaction chromatography, Eur. J. Biochem., 178, 197, 1988. 40. Bonants, P. J. M., Muller, F., Vervoort, J., and Edmondson, D. E., A 31P-nuclear-magnetic-resonance study of NADPH-cytochrome-P-450 reductase and of the Azotobacter flavodoxin/ferredoxin-NADP+ reductase complex, Eur. J. Biochem., 190, 531, 1990. 41. Bayerl, T., Klose, G., Ruckpaul, K., and Schwarz, W., Interaction of hexane phosphonic acid diethyl ester with phospholipids in the hepatic microsomes and reconstituted liposomes as studied by 31P-NMR, Biochem. Biophys. Acta, 812, 437, 1985. 42. Eberhart, D. and Parkinson, A., Reconstitution of rat cytochrome P-450 III Al (P-450p), requires phospholipids with unsaturated fatty acids, The Toxicologist, 11, 302, 1991. 43. Vervoort, J., Muller, F., Le Gall, J., Bacher, A., and Sedlmaier, H., Carbon-13 and nitrogen-15 nuclear magnetic resonance investigation of Desulfovibrio vulgaris flavodoxin, Eur. J. Biochem., 151, 49, 1985. 44. Stockman, B. J., Krezel, A. M., Markley, J. L., Leonhardt, K. G., and Straus, N. A., Hydrogen-1, carbon-13 and nitrogen-15 NMR spectroscopy of Anabaena 7120 flavodoxin: assignment of (3-sheet and flavin-binding site resonances and analysis of protein-flavin interactions, Biochemistry, 29, 9600, 1990. 45. Sanner, C., Macheroux, P., Riiterjans, H., Muller, F., and Bacher, A., 15N- and 13C- NMR investigations of glucose oxidase from Aspergillus niger, Eur. J. Biochem., 196, 663, 1991. 46. Moonen, C, T. W., van den Berg, W. A. M., Boerjan, M., and Muller, F., Carbon-13 and nitrogen15 nuclear magnetic resonance study on the interaction between riboflavin and riboflavin-binding apoprotein, Biochemistry, 23, 4873, 1984. 47. Beinert, W.-D., Riiterjans, H., and Muller, F., Nuclear magnetic resonance studies of the Old Yellow Enzyme, 15N NMR of the enzyme recombined with 15N-labeled flavin mononucleotides, Eur. J. Biochem., 152, 573, 1985. 48. Paulsen, K. E., Stankovich, M. T., Stockman, B. J., and Markley, J. L., Redox and spectral properties of flavodoxin from Anabaena 7120, Arch. Biochem. Biophys., 280, 68, 1990.

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49. Grande, H. J., van Schagen, C. G., Jarbandhan, T., and Miiller, F., An 'H-NMR spectroscopic study of alloxazines and isoalloxazines, Helv. Chim. Acta, 60, 348, 1977. 50. Vervoort, J., Muller, F., Lee, J., van den Berg, W. A. M., and Moonen, C. T. W., Identification of the true carbon-13 nuclear magnetic resonance spectrum of the stable intermediate II in bacterial luciferase, Biochemistry, 25, 8062, 1986. 51. Miura, R. and Miyake, V., 13C NMR studies of porcine kidney D-amino acid oxidase reconstituted with 13 C-enriched flavin adenine dinucleotide. Effects of competitive inhibitors, /. Biochem. (Tokyo), 101, 581, 1987. 52. Muller, F., Vervoort, J., van Mierlo, C. P. M., Mayhew, S. G., van Berkel, W. J. H., and Bacher, A., C-13, N-15 and two-dimensional NMR techniques in flavoprotein research, in Flavins and F lav oproteins, Edmondson, D. E. and McCormick, D. B., Eds., Walter de Gruyter, Berlin, 1987, 261. 53. Malthouse, J. P. G., Matherway, R., and Mayhew, S. G., A 13C NMR investigation of the environment of the thiol group of flavodoxin, Biochem. Soc. Trans., 17, 395, 1989. 54. Moonen, C. T. W. and Muller, F., On the mobility of riboflavin 5'-phosphate in Megasphaera elsdenii foavodoxin as studied by 13C nuclear magnetic resonance relaxation, Eur. J. Biochem., 133, 463, 1983. 55. Kasai, S., Sugimoto, K., Miura, R., Yamano, T., and Matsui, K., 8-Fluon>8-demethylriboflavin as a potential active-site-directed reagent for flavoproteins, Reaction with some amino acids, /. Biochem. (Tokyo), 93, 397, 1983. 56. Miura, K., Kasai, S., Horiike, K., Sugimoto, K., Matsui, K., Yamano, T., and Miyake, Y., 8-Fluoro8-demethyIriboflavin as a 19F-probe for flavin-protein interaction. A 19F NMR study with egg white riboflavin-binding protein, Biochem. Biophys. Res. Commun., 110, 406, 1983. 57. Macheroux, P., Kojiro, C. L., Schopfer, L. M., Chakraborty, S., and Massey, V., 19F NMR studies on 8-fluoroflavins and 8-fluoro-flavoproteins, Biochemistry, 29, 2670, 1990. 58. Ghisla, S. and Massey, V., New flavins for old: artificial flavins as active site probes of flavoproteins, Biochem. J., 239, 1, 1986. 59. Massey, V., Ghisla, S., and Moore, E. G., 8-Mercaptoflavins as active site probes of flavoenzymes, /. Biol. Chem., 254, 9640, 1979. 60. Schopfer, L. M., Massey, V., and Claiborne, A., Active site probes of flavoproteins. Determination of the solvent accessibility of the flavin position 8 for a series of flavoproteins, J. Biol. Chem., 256, 7329, 1981. 61. Peerson, O. B., Pratt, E. A., Truong, H,-T. N., Ho, C., and Rule, G. S., Site-specific incorporation of 5-fluorotryptophan as a probe of the structure and function of the membrane-bound D-lactate hydrogenase of Escherichia coli: a 19F nuclear magnetic resonance study, Biochemistry, 29, 3256, 1990. 62. Muller, F. and Vervoort, J., unpublished data. 63. Delseth, C., Nguyen, T. T.-T. and Kitzinger, J.-P., Oxygen-17 and carbon-13 nuclear magnetic resonance. Chemical shifts of unsaturated carbonyl compounds and acyl derivatives, Helv. Chim. Acta, 63, 498, 1980. 64. Vervoort, J., The Interactions Between Apoflavoproteins and Their Coenzymes as Studied by Nuclear Magnetic Resonance Techniques, Ph.D. thesis, Agricultural University, Wageningen, 1986, The Netherlands. 65. McDonald, C. C. and Phillips, W. D., Proton magnetic resonance spectra of proteins in random-coil configurations, J. Am. Chem. Soc., 91, 1513, 1969. 66. Crespi, H. L., Norris, J. R., and Katz, J. J., Magnetic resonance of isotope hybrid flavoprotein 2Hflavoprotein ('H-flavin mononucleotide), Nature (London) New Biol., 236, 178, 1972. 67. Crespi, H. L., Smith, C., Gajida, L., Tisue, T., and Ameraal, R. M., Extraction and purification of 'H, 2H and isotope hybrid algal cytochrome, ferredoxin arid flavodoxin, Biochim. Biophys. Acta, 256, 611, 1972. 68. Crespi, H. L., Norris, J. R., Bays, J. P., and Katz, J. J., ESR and NMR studies with deuterated flavodoxin, Ann. N.Y. Acad. Sci., 222, 800, 1973. 69. James, T. L., Ludwig, M. L., and Cohn, M., Dependence of the proton magnetic resonance spectra on the oxidation state of flavodoxin from Clostridium MP and from Peptostreptococcus elsdenii, Proc. Natl. Acad. Sci. U.S.A., 70, 3292, 1973. 70. Favaudon, V., Le Gall, J.-M., and Lhoste, J.-M., Proton magnetic resonance of Desulfovibrio vulgaris and Desulfovibriogigasftavodoxins, in Flavins and Flavoproteins, Singer, T. P., Ed., Elsevier, Amsterdam, 1976, 434. 71. van Schagen, C. G. and Muller, F., High resolution 'H NMR study at 360 MHz on the flavodoxin from Megasphaera elsdenii, FEBS Lett., 136, 75, 1981. 72. Rigby, S. E. J., Bishop, E. O., and Smith, B. E., Nuclear-magnetic-resonance spectroscopy of flavodoxins from diazotrophs, Biochem. Soc. Trans., 14, 1275, 1986. 73. Hazzard, J. T. and Tollin, G., Proton NMR study of the cytochrome c: flavodoxin electron transfer complex, Biochem. Biophys. Res. Commun., 130, 1281, 1985. 74. Tollin, G., Brown, K., De Francesco, R., and Edmondson, D. E., Flavodoxin-cytochrome c interactions: circular dichroism and nuclear magnetic resonance studies, Biochemistry, 26, 5042, 1987.

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75. Lubas, B., Soltysik, M., Steczko, J., and Ostrowski, W., Proton NMR study of the interaction of riboflavin with the egg-yolk apoprotein, FEBS Lett., 79, 179, 1977. 76. Yagi, K., Ohishi, N., and Nishikimi, M., Nuclear magnetic resonance study on D-amino acid and oxidase reaction, Biochim. Biophys. Acta, 206, 181, 1970. 77. Labeyrie, F., Beloeil, J.-C., and Thomas, M.-N., Evidence by NMR for mobility of cytochrome domain within flavocytochrome b2, Biochim. Biophys. Acta, 953, 134, 1988. 78. Hanemaaijer, R., Vervoort, J., Westphal, A., de Kok, A., and Veeger, C., Mobile sequences in the pyruvate dehydrogenase complex, the E2 component, the catalytic domain and the 2-oxoglutarate dehydrogenase complex of Azotobacter vinelandii, as detected by 600 MHz *H NMR spectroscopy, FEBS Lett., 240, 205, 1988. 79. Moonen, C. T. W. and Miiller, F., A proton-nuclear-magnetic-resonance study at 500 MHz on Megasphaera elsdenii flavodoxin. A study on the stability, proton exchange and the assignment of some resonance lines, Eur. J. Biochem., 140, 311, 1984. 80. Moonen, C. T. W. and Miiller, F,, On the intermolecular electron transfer between different redox states of flavodoxin from Megasphaera elsdenii, A 500 MHz *H NMR study, Eur. J. Biochem., 140, 303, 1984. 81. Kaptein, R., Dijkstra, K., Miiller, F., van Schagen, C. G., and Visser, A. J. W. G., 360 MHz laserinduced photo-CIDNP in photoreactions of flavins, J. Magn. Reson., 31, 171, 1978. 82. Miiller, F., van Schagen, C. G., and Kaptein, R., Application of nuclear magnetic resonance and photochemically induced dynamic nuclear polarization to free and protein-bound flavins, Methods Enzymol., 66, 385, 1980. 83. van Schagen, C. G., Miiller, F., and Kaptein, R., Photochemically induced dynamic nuclear polarization study on flavin adenine dinucleotide and flavoproteins, Biochemistry, 21, 402, 1982. 84. Moonen, C. T. W., Hore, P. J., Miiller, F., Kaptein, R., and Mayhew, S. G., A photo-CIDNP study of the active sites of Megasphaera elsdenii and Clostridium MP flavodoxins, FEBS Lett., 149, 141, 1982. 85. Moonen, C. T. W., Scheek, R. M., Boelens, R., and Miiller, F., The use of two-dimensional difference spectra in the elucidation of the active center of Megasphaera elsdenii flavodoxin, Eur. J. Biochem,, 141, 323, 1984. 86. van Mierlo, C. P. M., Vervoort, J., Miiller, F., and Bacher, A., A two-dimensional 'H NMR study on Megasphaera elsdenii flavodoxin in the reduced state. Sequential assignments, Eur. J. Biochem., 187, 521, 1990. 87. van Mierlo, C. P. M., Miiller, F., and Vervoort, J., Secondary and tertiary structure characteristics of Megasphaera elsdenii flavodoxin in the reduced state as determined by two-dimensional 'H NMR, Eur. J. Biochem., 189, 589, 1990. 88. van Mierlo, C. P. M., Lynzaad, P., Vervoort, J., Miiller, F., Berendsen, H. J. C., and de Vlieg, J., Tertiary structure of two-electron reduced Megasphaera elsdenii flavodoxin and some implications, as determined by two-dimensional 'H-NMR and restrained molecular dynamics, Eur. J. Biochem., 194, 185, 1990. 89. van Mierlo, C. P. M., van der Sanden, B. P. J., van Woensel, P., Miiller, F. and Vervoort, J., A two-dimensional 'H-NMR study on Megasphaera elsdenii flavodoxin in the oxidized state and some comparisons with the two-electron-reduced state, Eur. J. Biochem., 194, 199, 1990. 90. Stockman, B. J., Westler, W. M., Darba, P., and Markley, J. L., Detailed analysis of carbon-13 NMR spin systems in a uniformly carbon-13 enriched protein: flavodoxin from Anabaena 7120, J. Am. Chem. Soc., 110, 4095, 1988. 91. Stockman, B. J., Reily, M. D., Westler, W. M., Ulrich, E. L., and Markley, J. L., Concerted twodimensional NMR approaches to hydrogen-1, carbon-13 and nitrogen-15 resonance assignments in proteins, Biochemistry, 28, 230, 1989. 92. van Mierlo, C. P. M., Proton NMR Studies on Megasphaera elsdenii Ravodoxin: Structure Elucidation by 2D NMR and Implications, Ph.D thesis, Agricultural University, Wageningen, 1990, The Netherlands. 93. Hemmerich, P. and Massey, V., The role of the apoprotein in directing pathways of flavin catalysis, in Oxidase and Related Redox Systems, King, T. E., Mason, H. S., and Morrison, N., Pergamon Press, Oxford, 1979, 154. 94. Ludwig, M. L., Schopfer, L. M., Metzger, A. L., Pattridge, K. A., and Massey, V., Structure and oxidation-reduction behaviour of 1-deaza-FMN flavodoxins: modulation of redox potentials in flavodoxins, Biochemistry, 29, 10364, 1990. 95. Moonen, C. T. W., Vervoort, J., and Miiller, F., Some new ideas about the possible regulation of redox potentials in flavoprotein, with special reference to flavodoxins, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 493. 96. Schopfer, L. M., Ludwig, M. L., and Massey, V., A working proposal for the role of the apoprotein in determining the redox potential of the role of the apoprotein in determining the redox potential of the flavin in M. elsdenii flavodoxin, in Flavins and Flavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 399.

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97. Cast, R., Valk, B. E., Miiller, F., Mayhew, S. G., and Verger, C., Studies on the binding of FMN by apoflavodoxin from Peptostreptococcus elsdenii. pH and NaCl concentration dependence, Biochim. Biophys. Acta, 446, 463, 1976. 98. Hall, L. H., Orchard, B. J., and Tripathy, S. K., The structure and properties of flavins: molecular orbital study based on totally optimized geometries. I. Molecular geometry investigations, Int. J. Quantum Chem., 31, 195, 1987. 99. Hall, L. H., Orchard, B. J., and Tripathy, S. K., The structure and properties of flavins: molecular orbital study based on totally optimized geometries. II. Molecular orbital structure and electron distribution, Int. J. Quantum Chem., 31, 217, 1987. 100. Hall, L. H., Bowers, M. L., and Durfor, C. N., Further consideration of flavin coenzyme biochemistry afforded by geometry-optimized molecular orbital calculations, Biochemistry, 26, 7401, 1987.

APPENDIX A major problem in the attempt to edit a comprehensive overview of a major research field is that the space is limited. This holds also for these series. In order to partially compensate for these shortcomings some flavoproteins, not treated in these series, are listed below. To ease access to the literature the most recent references or reviews are given. Protein

Remarks

Old Yellow Enzyme Cytochrome c, quinone, and cytochrome P-450 reductases Glyoxylate carboligase Oxynitrilase Glucose oxidase NADPH-sulfite reductase Nitric oxide synthase NADH-cytochrome b5 reductase NADH dehydrogenase Superoxide generating NADPH oxidase

FMN FMN, FAD

FAD FAD FAD FMN, FAD

FAD FAD FMN FAD

Redox enzyme? Redox enzymes Redox activity? Redox activity? Redox enzyme Metal containing

— — — —

Ref.

1 2 3-5 6,7 8,9 10 11 12 13,14 15-17

APPENDIX REFERENCES 1. Schopfer, L. M. and Massey, V., Old Yellow Enzyme, in A Study of Enzymes, Vol. 2, Kuby, S. A., Ed., CRC Press, Boca Raton, FL, 1990, 247. 2. Johnson, M. S. and Kuby, S. A., Cytochrome c, quinone and cytochrome P-450 reductase, in A Study of Enzymes, Vol. 2, Kuby, S. A., Ed., CRC Press, Boca Raton, FL, 1990, 285. 3. Gupta, N. K. and Vennesland, B., Glyoxylate carboligase of Escherichia coli: some properties of the enzyme, Arch. Biochem. Biophys., 113, 255, 1966. 4. Chung, S.-T., Tan, R. T. Y., and Suzuki, L, Glycoxylate carboligase of Pseudomonas oxalaticus. A possible structural role for flavine-adenine dinucleotide, Biochemistry, 10, 1205, 1971. 5. Cromartie, T. H. and Walsh, C. T., Escherichia coli glyoxalate carboligase, properties and reconstitution with 5-deaza-FAD and 1,5-dihydro-deaza-FADH, J. Biol. Chem., 251, 329, 1976. 6. Jorns, M. S., Mechanism of catalysis by the flavoenzyme oxynitrilase, J. Biol. Chem., 254, 12145, 1979. 7. Jaenicke, L. and Preun, J., Chemical modification of hydroxynitrile lyase by selective reaction of an essential cysteine-SH group with a,p-unsaturated propiophenones as pseudo-substrates, Eur. J. Biochem., 138, 319, 1984. 8. Bright, H. J. and Porter, D. J. T., Flavoprotein oxidases, The Enzymes, 12, 421, 1975. 9. Frederick, K. R., Tung, J., Emerick, R. S., Masiarz, F. R., Chamberlain, S. H., Vasada, A., Rosenberg, S., Chakraborsky, S., Schopfer, L. M M and Massey, V., Glucose oxidase from Aspergillus niger. Cloning, gene sequence, secretion from Sacharamyces cerevisiae and kinetic analysis of a yeastderived enzyme, J. Biol. Chem., 265, 3793, 1990.

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10. Barber, M. J., Rueger, D. C., Miller, B. E., Siegel, L. M., and Kredich, N. M., Characterization of the flavoprotein moieties of NADPH-sulfite reductase from Salmonella typhimurium and Escherichia coll. Physicochemical and catalytic properties, amino acid sequence deduced from DNA sequence of cysJ and comparison with NADPH-cytochrome P-450 reductase, 7. Biol. Chem., 264, 15796, 1989. 11. Stuehr, D. J., Kwon, N. S., and Nathan, C. F., FAD and GSH participate in macrophage synthesis of nitric oxide, Biochem. Biophys. Res. Commun., 168, 558, 1990. 12. Yubisui, T., Shirabe, K., Takeshita, M., Kobayashi, Y., Fukumaki, Y., Sakati, Y., and Takano, T., Structural role of resine 127 in the NADPH-binding site of human NADH-cytochrome bs reductase, /. Biol. Chem., 266, 66, 1991. 13. Albracht, S. P. J. and van Belzen, R., The pathway of electron transfer in NADH: O2 oxidoreductase, Biochim. Biophys, Acta, 974, 311, 1989. 14. Singer, T. P., Ramsay, R. R., Krueger, M. J., and Youngster, S. K., Binding of MPP+ and of its analogs to the retonone/piericidin site of NADH dehydrogenase (NADH-ubiquinone oxidoreductase), in Flavins and Flavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 873. 15. Green, T. R. and Pratt, K. L., Detection and isolation of the NADPH-binding protein of the NADPH: O2 oxidoreductase complex of human neutrophils, J. Biol. Chem., 265, 19324, 1990. 16. Boscher, B. G. J. M., Denis, S. W., Verhoeven, A. J., and Roos, D., The activity of one soluble component of the cell-free NADPH: O2 oxidoreductase of human neutrophils depends on guanosine 5'-O(3-thio)triphosphate, J. Biol. Chem., 265, 15782, 1990. 17. Shaag, D. and Pick, E., Nucleotide binding properties of cytosolic components required for expression of activity of the superoxide generating NADPH oxidase, Biochim. Biophys. Acta, 1037, 405, 1990.

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Chapter 20

ACYL-COENZYME A DEHYDROGENASES Paul C. Engel TABLE OF CONTENTS I.

Metabolic Context

598

II.

Acyl-CoA Substrate Specificity A. The Acyl Moiety B. The Thiol Moiety

599 599 600

III.

Specificity for the Electron Acceptor

602

IV.

Oxygen Reactivity

603

V.

Spectral Characteristics of Acyl-CoA Dehydrogenases A. Introduction B. Semiquinone Forms and Charge-Transfer Complexes C. Green Complexes D. Enzyme-Acetoacetyl-CoA Complexes E. Red and Blue Semiquinones F. Reduced Forms G. Oxidized Spectrum

604 604 604 608 614 614 615 615

VI.

Structure A. Quaternary Structure B. Primary Structure C. Tertiary Structure

616 616 618 621

VII.

Chemical Modification Studies A. Group-Specific Reagents 1. Introduction 2. Cysteine 3. Methionine 4. Arginine 5. Dicarboxylic Amino Acids 6. Tyrosine B. Active Site-Directed Irreversible Inhibitors 1. Introduction 2. Acetylenic Inhibitors 3. 3,4-Pentadienoyl CoA 4. Methylenecyclopropyl Acyl-CoA Compounds 5. Propionyl CoA 6. Photoaffinity Labeling

622 622 622 624 625 625 625 626 626 626 626 627 628 628 630

598

Chemistry and Biochemistry of Flavoenzymes

VIII.

Mechanism A. Introduction B. Oxidation-Reduction Properties C. Stereochemistry D. Mechanism of Hydrogen Removal E. Side Activities

630 630 631 631 633 635

IX.

Medical Aspects A. Introduction B. Jamaican Vomiting Sickness C. Inborn Errors of Metabolism D. Medium-Chain Acyl-CoA Dehydrogenase Deficiency E. Short-Chain Acyl-CoA Dehydrogenase Deficiency F. Long-Chain Acyl-CoA Dehydrogenase Deficiency G. Isovalericacidemia

636 636 636 638 639 644 644 644

X.

Biosynthesis

645

XL

Mutants

646

Acknowledgments

646

References

647

I. METABOLIC CONTEXT After their initial discovery in the 1950s17 the acyl-CoA dehydrogenases suffered two decades of neglect, bemoaned in two previous brief summaries of the field8'9 by this reviewer who labeled them the "Cinderella of the flavoprotein field". Cinderella's attractions have at last been noticed, and since 1980 there has been an enormous upsurge of interest with many vigorous new groups entering the field. A few other recent reviews have been published by leading investigators, covering particular aspects of this important family of enzymes.10"12 The acyl-CoA dehydrogenases (EC 1.2.99.2,3) catalyze the general reaction1

This introduces or reduces a trans a, p double bond in an acyl-Coenzyme A thioester. The first representatives of this family of enzymes were discovered in the 1950s in the course of the elucidation of the pathway of p-oxidation (Figure 1) of long-chain fatty acids by the

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FIGURE 1. The (3-oxidation sequence of reactions. Fatty acyl-CoA compounds in the mitochondrial matrix pass around this sequence of 4 reactions in a descending spiral, losing 2 carbons from the acyl chain as acetyl CoA with each passage.

groups of Lynen in Munich, and Lardy and Green at Madison, Wisconsin.2 7 This pathway progressively breaks down CIS and C16 fatty acyl chains, two carbon units at a time, and the first of the repeated series of four reactions that brings this about is the acyl-CoA dehydrogenase reaction. For the straight-chain acyl-CoA dehydrogenases, substituents X and Y in Equation 1 are hydrogen atoms. More recently representatives of the family (1.3.99.10) have been identified which deal with branched chains arising from the breakdown of amino acids.13-14 Their substrates have either X=CH3 or Y=CH3 in Equation 1. A separate acyl-CoA dehydrogenase (1.3.99.7) has also been described1517 for glutaryl-CoA produced in the catabolism of lysine. These enzymes are all simple flavoproteins, containing noncovalently bound FAD and no other prosthetic group. The oxidant in the forward reaction (Equation 1) is ETF, "electron transferring flavoprotein".18'20 In the mitochondrial p-oxidation system this protein in turn feeds reducing equivalents to an iron-sulfur-containing flavoprotein, ETF-dehydrogenase,21 or ETF-CoQ reductase, which is an integral part of the respiratory chain. The acyl-CoA dehydrogenases share this acceptor system with sarcosine and dimethylglycine dehydrogenases.22'23 In aerobic organisms the reverse process, the reduction of unsaturated fatty acyl chains, is carried out by a different enzyme: the reaction involves NADPH as the reductant and the acyl chain is attached to the integral phosphopantetheine SH of acyl carrier protein rather than to Coenzyme A.24 In some anaerobic organisms, however, the redox potential allows the acyl-CoA dehydrogenase reaction to proceed in the reverse direction. The acyl-CoA dehydrogenase of such organisms functions as an enoyl-CoA reductase in a mechanism for disposing of excess reducing equivalents (Figure 2) by production of short-chain fatty acids.25-26 Physiologically, therefore, Reaction 1 does not operate reversibly in any one organism, but in different redox environments the reaction may be used either forward or in reverse.

II. ACYL-CoA SUBSTRATE SPECIFICITY A. THE ACYL MOIETY The repetitive nature of the p-oxidation spiral of reactions requires that the acyl-CoA dehydrogenase reaction be carried out eight times, in, for example, the breakdown of CIS stearic acid to 2-carbon units. Each time the chemistry is identical except for the changing length of the acyl chain. At first sight a single enzyme of relaxed chain-length specificity

600

Chemistry and Biochemistry of Flavoenzymes

FIGURE 2. The role of short-chain acyl-CoA dehydrogenase in fermentative organisms that produce volatile fatty acids as a way to dispose of reducing equivalents. The saturated fatty acyl CoA is converted to the fatty acid for export by transferring the CoA to another substrate carboxylic acid, thereby conserving the free energy of hydrolysis of the thioester bond.

might seem the answer to the task, but that would require substrates of very different physical character to compete on equal terms for the active site. Otherwise conversion of the poorest substrate would become a rate-limiting step and thereby also tie up an unnecessarily large proportion of the limited Coenzyme A pool. Instead, therefore, in mammalian mitochondria at least, there are separate acyl-CoA dehydrogenases for long, medium, and short chain substrates. The "medium-chain" enzyme (EC 1.3.99.3) is also known as general acyl-CoA dehydrogenase, and the "short-chain" enzyme (EC 1.3.99.2) as butyryl-CoA dehydrogenase (BCD). There is nevertheless some degree of overlap in chain-length specificity between the three enzymes, although the extent of such overlap varies according to species and method of assay. li27 - 28 In anaerobic bacteria that produce short-chain fatty acids, the main fermentation products tend to be propionate arising from lactate, and butyrate and valerate produced, respectively, from 2 acetate units or an acetate and a propionate by condensation and reduction.29 For a long time it was assumed that a single "butyryl-CoA dehydrogenase" functioning as a broadspecificity, short-chain enoyl-CoA reductase would serve the necessary role here.25 It now appears however, that the BCD from Megasphaera elsdenii shows very low activity towards the C3 substrate.30 There is no evidence here for a separate flavoprotein propionyl-CoA dehydrogenase. Interestingly, however, Simon and colleagues,31 seeking an activity in anaerobic bacterial extracts with the ability to reduce ethyl vinyl ketone, discovered an enzyme which appears to be an iron-sulfur dependent acrylyl-CoA reductase able as a side activity to reduce the vinyl group of this ketone. The evidence for separate branched-chain acyl-CoA dehydrogenases came initially from the study of human clinical conditions,32'34 (see also Section IX) in which impaired ability to oxidize a branched short-chain acid (isovaleric or a-methylbutyric) led to specific accumulation of that acid in body fluids. Despite the suggestion35 that this might result from altered substrate specificity in a single short-chain acyl-CoA dehydrogenase, there is now abundant evidence, mainly from Tanaka's group, for separately encoded distinct activities in man and rat, one for isovaleryl-CoA and another for a-methyl substituted acyl-CoA substrates. These enzymes are immunologically distinct, they have been separately purified, and the genes have been cloned and sequenced. 13 - 14 - 36 " 41 An a-methylbutyryl-CoA dehydrogenase which functions as an enoyl-CoA reductase has also been found in the parasitic nematode Ascaris suum.26

B. THE THIOL MOIETY Although the physiological carrier of the acyl group is Coenzyme A, the majority of the CoA molecule is not obviously required for the catalytic chemistry of the reaction. The ability of the acyl-CoA dehydrogenase to use thioesters based on simpler thiols was significant in early studies because Coenzyme A was not commercially available. Although

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FIGURE 3. The Coenzyme A molecule is shown with the thiol at the extreme left and the adenosine moiety, carrying the distinctive 3'-phosphate, on the right. The fragment to the left of the imaginary cleavage point represented by arrow 1 is AT-acetyl cysteamine. The material to the left of arrow 2 is pantetheine.

TABLE 1 Binding of S-Acetoacetyl Thioesters to M. elsdenii Butyryl-CoA Dehydrogenase Thiol CoASH Pantetheine jV-Acetoacetylcysteamine JV-Acetylcysteamine

Apparent Kd (jtM)

Apparent AG° (kj mol-1)

0.12 4.25

-30.5 -30.7 -22.5 -13.9

116 3660

Note: Data from Reference 43. Note that for the 3 truncated CoA analogues the titrations clearly indicated negative cooperativity. Although this was not seen with acetoacetyl CoA, this may be because only data for 90 to 100% saturation could be reliably calculated with this ligand. In all cases the Kd reported is for the weakest binding detected — i.e., as saturation was approached. The long-wavelength maximum was at 580 nm in all cases.

the Madison group overcame the problem by heroic purification of CoA from yeast by copper precipitation,42 the Munich group resorted to CoA analogues, employing jV-acetyl cysteamine thioesters2 (Figure 3). Since then this aspect of specificity has received relatively little attention. In the case of BCD from Megasphaera elsdenii, however, a series of acetoacetyl thioesters have been studied.43 Acetoacetyl CoA forms a spectrally recognizable complex with the enzyme,44 and the dissociation constant can be examined by spectrophotometric titration. Such studies give a rough indication of the contribution of different parts of the CoA structure to the overall binding energy (Table 1). As discussed in a later section, studies of the catalytic reaction reveal that the thiol portion of the substrate also influences the oxygen reactivity of the enzyme and thus fulfills an "effector" role. In the same vein, a revealing study was carried out by Frerman and co-workers45 on the mammalian medium-chain acyl-CoA dehydrogenase (MCAD). They compared the effectiveness of several alternative substrates, all with the same octanoyl moiety but with different thiol components (Table 2). The comparison between the CoA and pantetheine thioesters reveals a 50-fold increase in KM and a 50-fold decrease in V max for the latter, leaving no doubt that the adenosine phosphate portion contributes greatly to binding and successful catalysis. The 3'-dephospho-CoA analogue on the other hand is, if anything, a slightly better substrate than octanoyl CoA, and the etheno-CoA analogue is also quite a good substrate, indicating that the enzyme will tolerate the extra bulk of the bridge to the adenine 6-NH2 position. The same workers also tested S-octyl CoA, thioether rather than a

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TABLE 2 Kinetic Parameters for Oxidation of Various S-Octanoyl Thioesters Catalyzed by Porcine Medium-Chain Acyl-CoA Dehydrogenase Thiol

K M (JJLA/)

Vmax (min ')

V max /K M (min- 1 jJiAf-1)

CoA 3' Dephospho CoA Etheno CoA Pantetheine

2.21 1.09 9.28 107

675 657 416 14.1

305 603 44.8 0.13

Note: Data from Reference 45. Rates were measured with DCPIP as terminal acceptor and 0.41 to 0.45 |xM ETF as mediator. Temperature 20°C, pH not indicated.

thioester, and found competitive inhibition, with a K 4 of 6.5 |xM, which can be compared with the Ki for product inhibition by 2-octenoyl CoA under the same conditions of 3.4 (xM. Thus the carbonyl function of the thioester is also not essential for good binding. Frerman et al.45 introduce their study by noting the remarkably poor activity of free CoA, or indeed acetyl CoA or propionyl CoA, as inhibitors, seeming to question the major contribution of the CoA moiety to the strength of binding. Their findings therefore present an apparent paradox. The conclusion must be that tight binding depends on an element of induced fit, promoted by binding of an acyl chain of the right size, bringing the groups responsible for anchoring CoA into the right orientation. Thorpe et al.46 also examined the stringency of the requirement for a thioester, again with the mammalian MCAD. This time a 17-carbon chain was used. One analogue tested was again the thioether, S-heptadecyl CoA which showed a Kj of 40 nAf or less. The second analogue was heptadecan-2-onyl-dethio-CoA, in which the sulfur of CoA is replaced by carbon, and this turned out to be a substrate, giving rates and extents of enzyme reduction similar to those obtained with palmityl-CoA. This finding was taken to be in keeping with the mechanism put forward in Section VIII, in view of the expected acidity of the a-protons in the ketone.

III. SPECIFICITY FOR THE ELECTRON ACCEPTOR The "electron-transferring flavoprotein"19 (ETF) is the subject of a separate article in this review series.20 The rather nondescript name belies the fact that this protein funnels reducing equivalents from a very small set of flavoprotein dehydrogenases, passing them on to ETF dehydrogenase (ETF CoQ reductase), an integral component of the respiratory chain.21 These client dehydrogenases in turn are remarkable in their fidelity, using no other physiological oxidant at an appreciable rate. Most important of all, they are not reoxidized by molecular oxygen. As first shown by Beinert and his colleagues,47-48 substrate-reduced acyl-CoA dehydrogenases remain reduced in the presence of oxygen for many hours. What is the physiological significance of these properties? Although the issue has not been widely considered in the literature, it seems likely that the high degree of mutual specificity extends some of the properties of a membrane-bound array to a soluble system; thus, 1. 2.

It ensures that substrate-derived reducing equivalents are faithfully delivered to oxygen via the respiratory chain so that the ATP-generating mechanism is not short-circuited. It presumably extends respiratory control beyond the components of the membranebound chain, so that if ADP is depleted, mitochondrial p-oxidation will be switched off.

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Nevertheless it allows an ETF molecule to interact with several different dehydrogenases in a way that would not be possible in a membrane-bound system. From a practical biochemical standpoint, the specificity for ETF has been an obstacle to the study of acyl-CoA dehydrogenases since it makes assays difficult. The problem was circumvented by Seubert and Lynen3 by using a reductive assay for leucosafranine as reductant for 5-crotonyl Af-acetyl cysteamine. This does not require ETF but has not been widely used. Although other physiological acceptors will not substitute for ETF in the oxidative direction, various artificial systems can be used. As terminal acceptors, the most frequently used have been dichlorophenolindophenol (DCPIP),5 tetrazolium dyes,5'49 and cytochrome c.5'50 DCPIP and tetrazolium dyes can be used as sole acceptor, but in all cases the rates are greatly enhanced by addition of a mediator dye such as pyocyanine5 or phenazine methosulfate (PMS).50-51 Such assay systems are sensitive (i.e., they give good specific activity), and the mediator overcomes problems caused by varying amounts of residual ETF, but they still have significant drawbacks. The terminal acceptors, especially DCPIP, are nonspecific, and specificity cannot be guaranteed even by assaying with and without substrate, since thioesterase activity releases CoASH, which is itself an efficient reductant. PMS is light-sensitive and the photoreaction gives rise to spurious "blanks". 5154 Various improvements have been introduced. Thus, Meldolablau55 and phenazine ethosulphate (PES)30-54 have been used as substitutes for phenazine methosulfate. In the case of phenazine ethosulfate it is a reintroduction, since Mcllwain had noted already in 1937 that PES is less liable than PMS to undergo photolytic changes.52 More recently the ferricenium ion has been found to be an efficient acceptor with fewer complications.56 Nevertheless, in the context of clinical assays, there has been a return to assays based on purified ETF, used in this case as terminal acceptor, which gives rise to a large change in fluorescence upon reduction.57-58 A different approach which has been used with considerable success in measurements with living cells (e.g., skin fibroblasts), where high sensitivity and minimal interference are important, is isotope release from specifically labeled substrates. This technique, developed in Tanaka's laboratory, played an important part in establishing the existence of a separate isovaleryl-CoA dehydrogenase59 and has subsequently been refined by the use of a specific inhibitor to determine the extent of background release not attributable to the dehydrogenase.60

IV. OXYGEN REACTIVITY The question of oxygen reactivity deserves special consideration, both because control of flavin reactivity with oxygen is one of the principal sources of the catalytic versatility of flavoproteins and because, as noted above, the elimination of oxygen reactivity is of major physiological importance for the mitochondrial acyl-CoA dehydrogenases. The enzymes themselves are able to react with oxygen quite readily upon chemical reduction, e.g., with dithionite.47'51 It is only upon reduction with substrate that an oxygenstable species results and this is clearly an oxidized-substrate:reduced-enzyme complex. Thus, for example, Beinert and Page47 found that bovine medium-chain acyl-CoA dehydrogenase, upon reduction with a small excess of octanoyl CoA, remains reduced in air for many hours. There is scope in two directions for a comparative view of this stabilization of the reduced flavoprotein. First of all, as discovered by Lazarow and de Duve,61 mammalian cells possess a second (3-oxidation system located in a different organelle, the peroxisome. Its most obvious functional distinction is that the acyl-CoA oxidation is accomplished by a flavoprotein oxidase producing H2O2, rather than by a dehydrogenase. This pathway thus evades rigid respiratory control and is presumably important when it becomes necessary to use fatty acid oxidation more as a source of C2 units for biosynthesis and less as a source

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of ATP. This is similar to the situation in fatty seeds of plants where, some years earlier, an acyl-CoA oxidase activity was found in the glyoxysomes.62 Although a three-dimensional structure is not yet available for an acyl-CoA oxidase, the gene sequences63-64 reveal detectable homology with the acyl-CoA dehydrogenases.65 This is at a level where it is obvious only in limited sections of the sequence and requires insertion of numerous gaps. Indeed the existence of such homology has been denied by Osumi.66 Clearly, if the proteins are related, the divergence is a very distant event. A second interesting comparison comes from consideration of the butyryl-CoA dehydrogenase of M. elsdenii. In this organism, which is strictly anaerobic, the enzyme functions as an enoyl-CoA reductase25 and has no obvious need for protection against oxygen, which is not likely to be present in significant amounts in the natural milieu, the rumen of sheep.29 In fact when this enzyme is assayed in the direction of acyl-CoA oxidation there is an obvious oxidase activity.51-67 This was apparent in studies of the purified enzyme in which aerobic addition of butyryl Co A led not to a stable reduced complex, as with the mammalian enzymes, but to an oxidized-enzyme:oxidized-product complex.44 Subsequently this activity has been studied more systematically. There is no evidence for superoxide production. H2O2 appears to be the main reduced product of oxygen.67-68 This reaction was also studied with substrate analogues truncated in the CoA moiety. Such truncation impedes the dehydrogenase activity, as measured in dye-coupled assays, but not the oxidase activity.67*68 The obvious inference from such studies is that the adenosine moiety of Coenzyme A plays a key role in altering the environment of the isoalloxazine ring, presumably by conformational change in the protein, thereby abolishing or minimizing oxygen reactivity.

V. SPECTRAL CHARACTERISTICS OF ACYL-CoA DEHYDROGENASES A. INTRODUCTION The distinctive spectrophotometric characteristics of the flavin ring in various states of reduction and molecular association have of course provided a powerful tool in the study of flavoprotein mechanisms generally, and the recognition of the diagnostic features is therefore crucial. In this context the acyl-CoA dehydrogenases have played a key role in providing early evidence: first, of the semiquinoid state in flavoproteins and secondly, of flavin charge-transfer complexes. The fact that these enzymes exist in both these states, and that both are characterized by long-wavelength visible absorption indeed led to some initial confusion. B. SEMIQUINONE FORMS AND CHARGE-TRANSFER COMPLEXES It is now generally recognized that one of the most important sources of versatility in flavoprotein catalysis is their ability to make use of 3 possible reduction states, fully oxidized, one-electron reduced (semiquinone), and fully (two-electron) reduced. This enables them to encompass remarkably variable oxidation-reduction spans and to act as transducers between one and two-electron oxidation-reduction systems. Nevertheless, even though the semiquinoid state of riboflavin had been recognized in a number of laboratories in the 1930s, it was not identified in flavoproteins until 1956.69-47 Beinert reported that medium-chain acyl-CoA dehydrogenase showed a transitory broad absorption band between 500 and 650 nm, both during reduction with dithionite and again during reoxidation by air. A band of the same amplitude and in the same region was also produced during reduction by substrate but was less transitory and indeed extremely stable in air. Beinert concluded at this stage that dithionite reduction produced a semiquinoid intermediate on the way to full reduction, and that substrate, if present, stabilized this state.

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By 1961, however,70-71 a problem with this interpretation had emerged. The semiquinone is a radical species and, as such, should give rise to an electron paramagnetic resonance (EPR) signal. Beinert and Sands71 reported an EPR investigation of the oxidation-reduction properties of various flavoproteins. Some of them did indeed produce a radical signal upon partial reduction by substrate, but in the specific case of the acyl-CoA dehydrogenases this did not occur. The long-wavelength absorption produced during reduction by sodium dithionite or reoxidation correlated very well with the appearance of an EPR signal, but when, in the absence of dithionite, substrate and enzyme were mixed, the brownish-green complex that was formed was EPR-silent. Over a longer time-scale (hours), a free-radical signal was detected but the overall absorbance at 570 nm gradually declined as this signal built up. The slow emergence of the radical appeared to correlate with reoxidation by air and was much delayed under anaerobic conditions. The maximal yield of radical never exceeded a few percent of the total flavin, in marked contrast to the situation during reduction by dithionite. The EPR-positive species was clearly formed far too slowly to be a catalytic intermediate, but to what then was the rapidly formed long-wavelength band due? Beinert and Sands obtained a high yield of radical, even after addition of substrate, by the subsequent introduction of dithionite, and deduced that the flavin was perhaps not fully, or even 50% reduced, in the enzyme-substrate complex. They were nevertheless unwilling to believe that a complex could exist between (initially) oxidized enzyme and reduced substrate without some degree of oxidoreduction occurring, and they considered two possible explanations: either the substrate-flavin interaction might be a charge-transfer pairing; or else perhaps interaction between two neighboring radicals might broaden the EPR signal beyond detection. Interaction between neighboring FAD radicals would have been hard to reconcile with the ready detection of EPR signals during dithionite reduction, but interaction of flavin and substrate radicals is less easy to dismiss as a hypothesis. The intellectual climate of the time was not very receptive to the idea of charge-transfer complexes in flavoproteins. Physical chemists demanded rigorous evidence and in particular wanted to see a demonstration that the energy of interaction, and hence the wavelength of the new charge-transfer absorption band, varied with the donor and acceptor potential and with polarity of the medium.72'74 These tests were not at the time readily applicable for most proteins by virtue of the exigencies of protein stability and substrate specificity. The case of the "Old Yellow Enzyme" was therefore of general importance, because this enzyme forms long-wavelength-absorbing complexes with a wide variety of aromatic and heteroaromatic aldehydes and ketones,75 and because it also proved possible to prepare apoenzymes, and replace the FMN with analogues of varying oxidation-reduction potential.76 Abramovitz and Massey demonstrated a correlation between the wave number of the long-wavelength maximum and the Hammett a function for a series of para-substituted phenols. A correlation with the oxidation-reduction potential of the flavin was also shown. The issue has been reviewed in detail by Massey and Ghisla.77 The short-chain acyl-CoA dehydrogenase of M. elsdenii is a prolific producer of longwavelength bands both in the fully oxidized and the partially reduced states44'51'78 (Table 3, Figure 4, 5) but some of the larger changes in absorbance maximum are produced by methyl substituents, and it is clear that this reflects changes in orientation rather than intrinsic donor potential. No systematic correlation with donor potential has been attempted. (It is assumed, in the oxidized complexes, that FAD is the charge donor, but in the partially reduced complexes seen during catalysis, the situation is reversed with reduced flavin as the donor and oxidized substrate as the acceptor.) At the time of these studies it had not proved possible to produce an apoprotein that could be reactivated. More recently this has been done,79 but the putative charge-transfer complexes have not as yet been reinvestigated with the enzyme reconstituted with FAD analogues. In the case of pig medium-chain acyl-CoA dehydrogenase,

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TABLE 3 Long-Wavelength Absorption Maxima of Complexes Formed by M. elsdenii ButyrylCoA Dehydrogenase with Various Acyl-CoA Compounds Oxidized Complexes Acyl-CoA compound

Pent-2-enoyl CoA 4-Methylpent-2-enoyl CoA Malonic semialdehyde CoA Methylmalonic semialdehyde CoA Acetoacetyl CoA 2-Methylacetoacetyl CoA 3-Oxopentanoyl CoA 2-Methyl-3-oxopentanoyl CoA Cyclohept-2-ene carboxylic acyl CoA

Long-wavelength maximum (nm)

810 900 500—650

575 580 500—600

560 500—600 >900

Reduced Complexes Crotonyl CoA 2,4 Pentadienoyl CoA Cyclohex-2-enoyl CoA

580 680 560

Note: Data from References 9, 44, 78, and 98; for cyclohex-2-enoyl CoA, Engel, P. C, unpublished results. This reduced complex is unusually stable for M, elsdenii BCD under aerobic conditions. In some cases the complex has been formed by direct addition of unsaturated acyl CoA (or 3-oxo-acyl CoA); in others the complex has been formed by aerobic turnover and its composition is inferred.44 As discussed in Section VIII.D, the composition of the complex formed upon addition of pent-2-enoyl CoA must now be in some doubt.

FIGURE 4. Formation of a long-wavelength absorbing complex between M. elsdenii BCD and acetoacetyl CoA. The enzyme was stripped of other CoA-containing ligands and 26.4 \LM (in terms of subunits) enzyme (dashed spectrum) was treated with a 3.3-fold molar excess of acetoacetyl CoA to give the spectrum shown by the dashed line. The buffer was 0.1 M potassium phosphate, pH 7. (Adapted from Engel, P. C., Williamson, G., and Shaw, L., in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 403.)

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FIGURE 5. Formation of a long-wavelength absorbing complex between reduced M. elsdenii BCD and penta-2, 4-dienoyl CoA. The yellow form of the enzyme was used and the starting spectrum is shown by the dashed and dotted curve (-•-•). The dotted curve (•••) shows the reoxidized spectrum a few minutes after addition of a 2.5-fold molar excess of pent-4-enoyl CoA. The other curves show intermediate reduced spectra obtained in two different ways: (1) the solid curve ( ) was obtained by addition (aerobically) of a 60-fold molar excess of pent-4-enoyl CoA and reading the spectrum on a scanning spectrophotometer; (2) the dashed line (—) with separate experimental points was obtained with a 2.5-fold excess of substrate in the stopped-flow spectrophotometer, each point (•) indicating the maximum change observed at a particular wavelength. (From Engel, P. C. and Massey, V., Biochem. J., 125, 889, 1971. With permission.)

however, a convenient method80 for producing a reconstitutable apoenzyme in high yield has allowed studies with FAD analogues.81 An active enzyme was obtained with 8-chloro, 7-bromo, and 2-thioFAD (Figure 6), and the position of maximum long-wavelength absorption was altered in the direction predicted by the increasing oxidation-reduction potential of these flavins (i.e., towards shorter wavelength). Much earlier, in a detailed study of MCAD from beef liver in Drysdale's laboratory, Murfin82 took advantage of the enzyme's activity with phenylpropionyl CoA (hydrocinnamoyl-CoA) as substrate. Various aspects of the reaction with phenylpropionyl CoA and different ring-substituted derivatives were examined, including the absorption spectrum of the putative charge-transfer complex. The /?-OH, /?-CH3O, /?-Cl, and m-NO2 derivatives were examined in addition to the unsubstituted compound, and a systematic variation of absorption maximum from about 610 nm to 720 nm was found to correlate well with the Hammett a constants (Table 4). The issue of what constitutes appropriate and adequate evidence for the charge-transfer nature of a flavoprotein-ligand complex has continued to generate lively controversy,83-84 but it is probably now generally accepted that the long-wavelength absorption band formed upon mixing acyl-CoA dehydrogenases with reduced substrate is indeed due to a chargetransfer interaction.

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 6. Long-wavelength bands formed by an acyl-CoA dehydrogenase reconstituted with an FAD analogue. Pig MCAD was reconstituted with 8-C1 FAD. The solid line ( ) shows the spectrum of 6.1 |J-A/ reconstituted MCAD. The dashed line (—) shows the result of adding acetoacetyl CoA (6-fold excess) and the dotted line (•••) the result of separately adding a 2-fold excess of octanoyl CoA. The long-wavelength bands are shifted 20 nm to the red for the oxidized complex and 20 nm to the blue for the reduced complex, in keeping with the role of the flavin as a charge-transfer acceptor in the first and a donor in the second. (From Thorpe, C. and Massey, V., Biochemistry, 22, 2972, 1983. With permission.)

TABLE 4 Position of Long-Wavelength Absorption Maximum in Complexes of Reduced Bovine Medium-Chain Acyl-CoA Dehydrogenase with Ring-Substituted Cinnamoyl-CoA Derivatives Substitution p-OH

/?-CH3O H

p-a

m-NO2

Absorption maximum 615 635 655 668 720

nm nm nm nm nm

Hammett 1(XU07'113-117

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FIGURE 12. The effect of borohydride reduction on BCD. Panel (a) shows the spectrum of bovine BCD (9.8 u,Af bound flavin) before (solid line) and after (dashed line) addition of an excess of NaBH4. The spectrum was recorded once H2 emission had ceased. Panel (b) shows the spectrum of the flavin extracted with 7.5% trichloracetic acid from an enzyme sample (22 jjiAf) treated as in (a). TCA was removed with ether after centrifuging to separate the white protein pellet. The sample was then buffered at pH 7.6 before reading the spectrum.

Early estimates1 based on flavin content, which should give a minimum value for the molecular weight, gave very high values, in some cases in excess of 100,000. It is now clear that the early preparations must have been partially depleted of flavin, a view that is borne out by high values of A^^A^ relative to those now routinely obtained. Purifications

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 13. Conversion of green BCD to a yellow form. In this experiment, carried out in a stoppered cuvette without any volume changes, the green sample of M. elsdenii BCD (solid curve), well buffered in 0.1 M phosphate, pH 7, was first reduced with a few grains of solid sodium dithionite (dashed curve). After the slow reduction, taking some minutes at room temperature, air was readmitted and the dashed spectrum was obtained without any trace of long-wavelength absorption. There is an isobestic point at 430 nm very close to the absorption maximum of the green form. (From Engel, P. C. and Massey, V., Biochem. J., 125, 879, 1971. With permission.)

published since 1960 have all taken advantage of new chromatographic media, allowing much milder conditions. Even so, Ikeda et al.13-14'114'115 report in their series of papers on the separation of five acyl-CoA dehydrogenases from rat liver that activity was in some cases enhanced 1.5 to 2.5-fold by addition of FAD. In the case of isovaleryl-CoA dehydrogenase,13 they estimated from spectrophotometric evidence that the purified enzyme (before adding back FAD) contained 0.6 mol FAD/mol subunit. They found114 that of the straight-chain acyl-CoA dehydrogenases, the short-chain enzyme bound FAD the most tightly, whereas the other two lost the prosthetic group more readily. The Kd for FAD was 2.4 (xM for the medium-chain acyl-CoA dehydrogenase. In general the molecular weights have not been pursued with great physicochemical rigor: usually the subunit molecular weight has been estimated by calibrated SDS-PAGE and the native molecular weight by calibrated gel filtration (Table 6). However, in some cases the approximate estimates of molecular weight have now been backed up by precise values from cDNA sequencing38'39-41'118'119 and the agreement is good. As to quaternary structure, in the case of butyryl-CoA dehydrogenase of M. elsdenii, Williamson98 carried out a cross-linking study with dimethylsuberimidate, and strong bands were seen for monomer, dimer, trimer, and tetramer but not for higher aggregates. For mammalian MCAD the tetramer structure is now directly confirmed by the results of X-ray crystallography.120'122 B. PRIMARY STRUCTURE Surprisingly, none of the acyl-CoA dehydrogenases have been sequenced via the protein. This is probably because the prolonged period of their neglect roughly coincides with the two decades of greatest activity in protein sequencing. Since the acyl-CoA dehydrogenases began to attract widespread interest once more in the 1980s, the method of choice has been DNA sequencing, and there are indeed now cDNA sequences available for the medium-118

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TABLE 6 Molecular Weight Estimates for Various Acyl-CoA Dehydrogenases Enzyme Subunit Mr Butyryl-CoA dehydrogenase

Medium-chain acyl-CoA dehydrogenase

Long-chain acyl-CoA dehydrogenase Isovaleryl-CoA dehydrogenase 2-Methyl branched-chain acyl-CoA dehydrogenase

Native Mr Butyryl-CoA dehydrogenase

Medium-chain acyl-CoA dehydrogenase

Long-chain acyl-CoA dehydrogenase Isovaleryl CoA dehydrogenase 2-Methyl branched-chain acyl-CoA dehydrogenase

Source

Ref.

Estimate of Mr

Method

M. elsdenii Rat liver Human liver Pig kidney Rat liver Rat liver Human liver Rat liver Rat liver Human Rat liver

51 229 40 109 229 119 40 229 13 40 14

43,400 41,000 41,000 42,000 45,000 43,700 44,000 45,000 43,000 42,000 41,500

a c c c c d c c c c c

Ascaris Suum

26

42,000

c

M. elsdenii Rat liver Human liver Pig kidney Rat liver Human liver Rat liver Rat liver Human liver Rat liver

51 229 40 109 229 40 229 13 40 14

150,000 160,000 168,000 160,000 180,000 178,000 180,000 175,000 172,000 170,000

b b b b b b b b b b

Ascaris suum

26

170,000

b

Note: Methods are (a) amino acid analysis and flavin extraction; (b) calibrated gel filtration; (c) SDS-PAGE; (d) DNA sequencing.

and short-chain123 acyl-CoA dehydrogenases from man and for the long-, medium-,39'119 and short-chain39 acyl-CoA and isovaleryl-CoA39 dehydrogenases from rat. A detailed sequence comparison (Figure 14) has been presented by Matsubara et al.39 As might be expected, the cross-species comparisons between two mammals, man and rat, show very high levels of identity — 86.5% for medium-chain and 90.3% for short-chain acyl-CoA dehydrogenases. Within a species, for the rat, comparisons show remarkably homogeneous levels of similarity across the family, ranging from 30.6% identities, up to 35.4%. Predictably, perhaps, isovaleryl-CoA dehydrogenase seerns to be closer to short-chain acyl-CoA dehydrogenase (34.1% identities) than to the others. The closest pair are the medium- and short-chain enzymes (35.4%). What is slightly surprising is that long-chain acyl-CoA dehydrogenase shares more identities with the short-chain enzyme (31.7%) and indeed isovaleryl-CoA dehydrogenase (32.0%) than the medium-chain acyl-CoA dehydrogenase (30.6%). Without seeing similar comparisons for other species, one cannot be certain whether these small percentage differences are true reflections of ancestry or merely random "noise". A search for highly conserved residues reveals 57 invariant residues and a further 94 that were found in three out of the four rat sequences. Glycines, critically important in determining the folded shape, were very highly represented in these two groups (15 out of 57 and 9 out of 94). Significantly, in terms of mechanism, neither cysteines nor histidines were highly conserved. The other noteworthy point emerging from these comparisons is that the similarity

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 14. Alignment of several acyl-CoA dehydrogenase sequences and also that for rat peroxisomal acyl-CoA oxidase. RSCAD, RMCAD, RLCAD, and RIVD denote the rate short-, medium-, and long-chain acyl-CoA dehydrogenases and isovaleryl-CoA dehydrogenases and isovaleryl-CoA dehydrogenase, respectively. HSCAD and HMCAD indicate the human short- and medium-chain enzymes; and RACOX, the rat oxidase. Dark shading indicates the same residue in all 6 acylCoA dehydrogenases. A dot over the column indicates a residue the oxidase shares with all or most of the dehydrogenase sequences. (From Matsubara, Y., Indo, Y., Naito, E., Osaza, H., Glassberg, R., Vockley, J., Ikeda, Y., Kraus, J., andTanaka, K., J. BioL Chem., 264, 16321, 1989. With permission from Wiley-Liss Inc.)

extends virtually throughout the sequence (Figure 15). Although there are patches of lesser conservation, they are uniformly spread. One would have predicted that the requirements to form a tetramer, to bind ETF, to anchor FAD, and provide a site for CoA would all tend to dictate sequence conservation. On the other hand, the differences in fatty acyl-chain specificity might have led one to anticipate a substrate-binding domain showing less similarity. This is not obvious from the comparison. A surprising find is the lack of conservation of Glu-416. (This numbering refers to the overall alignment;39 in the numbering for pig medium-chain, acyl-CoA dehydrogenase, it is Glu-401). It had been proposed (see Section VII) that this might be a critical catalytic residue, but if so, its role must be assumed by a substitute in another position in the longchain acyl-CoA and isovaleryl-CoA dehydrogenases.

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621

(continued).

None of the nonmammalian acyl-CoA dehydrogenases have been sequenced. We have, however, one fragment of unpublished sequence for M. elsdenii BCD. This sequence of 36 residues from the N-terminus was originally determined with the intention of designing oligonucleotide probes. The alignment (Figure 16) once again shows identities at the level of over 30% for this small region. This is striking (1) because the N-terminus is not the most highly conserved section; (2) because of the large evolutionary distance between the organisms, and (3) because the bacterial enzyme actually functions in the reverse direction. C. TERTIARY STRUCTURE As mentioned above, an X-ray crystallographic study has now provided a detailed structure120"122 for one member of the acyl-CoA dehydrogenase family, the medium-chain enzyme from pig kidney. The level of homology discussed above gives some confidence that this study will provide a framework for understanding structure-function relationships throughout the family. The schematic backbone drawing121 of a monomer (Figure 17) shows that the sequence folds into N-terminal (120 residues) and C-terminal (150 residues) domains, both mainly comprising antiparallel a-helices, separated in the sequence by a section of about 90 residues forming two layers of 3-sheet structure (Figure 18). The FAD is, as usual, bound in an open, extended conformation close to the surface, and the four flavins are well separated, with 30 A as the shortest distance between isoalloxazine rings. The flavin-binding scaffold is quite different from that found in other flavoprotein structures so far determined. The isoalloxazine ring is wedged between the N-terminal helical domain and the p-sheet domain; the ADP moiety lies between the C-terminal and p-sheet domains of the same subunit and the C-terminal domain of a neighboring subunit. There is no "Rossmann fold" supporting the FAD binding site.

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Chemistry and Biochemistry of Flavoenzymes

FIGURE 15. A diagonal sequence comparison of the long- and medium-chain acylCoA dehydrogenase precursor sequences from rat. Constructed with the COMPARE program, window 30 residues, stringency 18. The comparison shows that similarity extends throughout the sequences of the mature proteins. (From Matsubara, Y., Indo, Y., Naito, E., Osaza, H., Glassberg, R., Vockley, J., Ikeda, Y., Kraus, J., and Tanaka, K., J, Biol Chem., 264, 16321, 1989. With permission from Wiley-Liss Inc.)

The structure determination has now been taken to high resolution122 with structures for both the free enzyme and an enzyme-substrate complex at 2.41 A. In the vicinity of the isoalloxazine ring, the side chain of Trp-66 lies over the benzenoid ring, and the side chains of Ile-371, Tyr-375, and Tyr-133 also make close approach to the ring system. Tyr-375 moves aside to make way for the substrate. This structure was determined for enzyme cocrystallized anaerobically in the presence of octanoyl CoA and shows a hydrophobic tunnel to bind the acyl moiety. The detailed analysis of this structure and that of other complexes in the near future should prove extremely informative.

VII. CHEMICAL MODIFICATION STUDIES A. GROUP-SPECIFIC REAGENTS 1. Introduction Over the past few years information about possible essential residues has come increasingly from the use of mechanism-based inhibitors, from genetic variants, and now from the determination of the first 3-dimensional structure for an acyl-Coenzyme A dehydrogenase. Previously such information had come only from the net-casting approach of chemical modification with group-selective reagents. In general, this approach has been remarkably unsuccessful in providing a plausible picture of the catalytic machinery of the acyl-CoA dehydrogenase. One possible explanation is that investigators have not usually ensured that their enzyme preparations were free of active-site ligands. It is very likely that many such

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FIGURE 16. A comparison between the N-terminal sequence of M. elsdenii BCD and the sequences of human medium- and shortchain acyl-CoA dehydrogenases. The first 36 residues of the M. elsdenii BCD sequence (ME) were determined by automated Edman degradation at the University of Leeds SERC sequencing facility and aligned by eye with the human sequences (HM = medium chain, HS = short chain), as given in Reference 39. Over this stretch each of the mammalian sequences shows 10 identities (28%) with the bacterial sequence; these are boxed in the diagram. Over the same stretch the two human enzymes show 13 identities (36%); those that are not shared with the bacterial enzyme are marked with a black spot.

FIGURE 17. a-Carbon chain trace of pig MCAD from the medium-resolution (3A) structure determined by X-ray crystallography. (From Kim, J.-J. P., Vollmer, S. H., and Frerman, F. E., Proc. NatL Acad. Sci. U.S.A., 85, 6677, 1988. With permission.)

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FIGURE 18. Representation of the secondary structure elements in the pig MCAD monomer. a-Helices are shown as cylinders and (i-sheet strands as arrowed ribbons. (From Kim, J.-J. P., Vollmer, S. H., and Frerman, F. E., Proc. Natl. Acad. Sci. U.S.A., 85, 6677, 1988. With permission.)

experiments have been doomed to failure by the use of what were, in effect, protected enzyme samples. It was pointed out as early as 1958 for the mammalian enzymes that, after adding radioactive substrate, the enzymes remained stoichiometrically labeled unless denaturing measures were employed.48 Later studies with the M. elsdenii short-chain acylCoA dehydrogenase again showed that, whatever the spectral form of the enzyme, there was always stoichiometrically bound CoA (or CoA derivatives) unless specific measures were adopted to remove it.90'91 Chemical modification studies have not yet been repeated with "stripped" enzyme. The same is true, of course, for residues involved in binding the prosthetic group. The tightness of binding of FAD or FMN in a flavoprotein makes it much harder to modify residues in the cofactor binding site, than, say, for an N AD+-dependent dehydrogenase. Despite these comments there have been a few experiments implicating various amino acid side chains in different aspects of activity. 2. Cysteine Initial attempts51 to modify protein-SH groups sprang from the suggestion88 that green forms of BCD might involve a charge-transfer interaction between a cysteine thiolate donor and flavin as acceptor. Mahler85 had also reported inhibition of bovine BCD by mercurial. Experiments51-89'95 on the enzyme from M. elsdenii showed that various reagents (phenylmercuric acetate, A^-ethylmaleimide, DTNB) abolished the green color, but it became clear (1) that the reagents were achieving this effect by virtue of their reaction with an active-site ligand rather than the protein (see Section V.C); (2) that some of them clearly also modified protein-SH groups without any effect on catalytic activity. Although amino acid analysis originally suggested up to 5 cysteine residues in M. elsdenii butyryl-CoA dehydrogenase, later experiments with DTNB and also separation of carboxymethylated peptides124 suggest only 3 residues. Titrations with mercurial89 would tend to indicate that they can probably all be modified without loss of activity. It was also clear that none of these groups were

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accessible to iodoacetate or iodoacetamide in the native state of the enzyme. The availability of the 710 nm absorbance signal also revealed differences in kinetic accessibility of the various reactive sulfur sites. These experiments have thus ultimately provided evidence against involvement of thiol groups in the activity of butyryl-CoA dehydrogenase, and this is consistent with the fact that no invariant cysteine residues are found in sequence comparisons across the family.39 Okamura-Ikeda et al.125 have offered evidence obtained with the short-, medium-, and long-chain acyl-CoA dehydrogenases of rat liver suggesting that there is an -SH group in the vicinity of the flavin binding site. Their method of preparation of the medium- and long-chain acyl-CoA dehydrogenases yielded both holoenzyme and apoenzyme. Only the latter forms were susceptible to inactivation by Af-ethylmaleimide, pchloromercuribenzoate and iodoacetate — i.e., the flavin cofactor protects completely. The short-chain enzyme was only obtained with bound flavin, but in this case even the holoenzyme was inactivated by P-chloromercuribenzoate. Significantly, however, all three enzymes reacted also with methyl mercury halide without disturbing the flavin. It seems therefore that the -SH group in question is not essential for FAD binding, and that inactivation results only from the hindrance caused by bulky substituents. The porcine medium-chain acyl-CoA dehydrogenase shows no inactivation in response to the same reagents126 and its 5-6 cysteine residues per FAD are accessible to DTNB only after denaturation. 3. Methionine Iodoacetate inactivates porcine medium-chain acyl-CoA dehydrogenase at pH 6.6 by reacting with a single methionine residue per subunit.126 Flavin binding is unaffected. Modification affects the interaction between the acyl-CoA substrate and the enzyme: although perturbation of the flavin spectrum indicates that the reduced substrate still binds, no flavin reduction occurs. Accordingly also prior addition of substrate protects almost completely against inactivation. 4. Arginine Arginine residues have frequently been found in the binding sites for nucleotide substrates, providing electrostatic bonding for the charged pyrophosphate linkage. Jiang and Thorpe127 treated their medium-chain acyl-CoA dehydrogenase with the arginine-specific reagent cyclohexane-l,2-dione and obtained somewhat equivocal results. Modification of 1.3 arginine per FAD caused pseudo-first-order loss of 90% of activity assayed in borate buffer, but the same sample showed 55% residual activity in phosphate buffer. This activity shows a marked lag. The authors discuss whether the recovery reflects dissociation of borate from the arginine-dione adduct or of the dione from the arginine. They favor the former conclusion which implies a partially active cyclohexanedione adduct. Their argument does not appear to take full account of the influence of rapid and tight binding of substrate on a reversible equilibrium with the reagent. It still seems entirely possible that their observations reflect modification of a truly essential residue by a reagent that can be displaced by excess substrate. The argument as to whether observed activity reflects partially active modified enzyme molecules is similar to the earlier controversy over partial inactivation of various enzymes by reaction of lysyl side chains with pyridoxal phosphate.128 5. Dicarboxylic Amino Acids The side chains of aspartate and glutamate are in general modifiable by carbodiimides. These were an interesting target for chemical modification since the rather acidic acyl-CoA dehydrogenase molecules were thought to interact with the natural acceptor ETF, at least in part, through electrostatic forces.129 Frerman et al.130 employed the water soluble diimide l-ethyl-3 (3-dimethylaminopropyl)-carbodiimide (EDC) to modify pig MCAD. The modified enzyme was markedly inhibited when assayed with ETF, but activity was unaffected in

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assays mediated by PMS. In these, the KM for butyryl CoA was unaffected, and acyl CoA also failed to protect the enzyme against EDC. The authors concluded that the modification of carboxylate residues specifically impaired binding to ETF. 6. Tyrosine Nothing has been published on chemical modification of tyrosine in acyl-CoA dehydrogenase. However, Dr. Gary Williamson obtained evidence several years ago in the reviewer's laboratory for inactivation of M. elsdenii butyryl-CoA dehydrogenase by tetranitromethane, normally regarded as a reasonably specific reagent for tyrosyl residues. The reaction produced the expected spectral changes and the modified enzyme clearly bound its FAD less tightly. The nature of the modification, however, was not rigorously confirmed by reduction and isolation of aminotyrosine. It is, nevertheless, now interesting to note that in the emerging structure for the mammalian medium-chain acyl-CoA dehydrogenase121'122 two tyrosine residues, Tyr-375 and Tyr-133, appear to interact with the isoalloxazine ring system of the bound FAD. Whether homologous tyrosines are present in the bacterial enzyme is not known. B. ACTIVE-SITE-DIRECTED, IRREVERSIBLE INHIBITORS 1. Introduction In contrast to the types of reagent just discussed, active-site-directed inhibitors are designed to resemble in some way the natural substrate(s) so that, at low concentrations, they will react exclusively with residues in the vicinity of the active site. They may be further subdivided into those that are intrinsically reactive compounds and those131'132 that are activated through the catalytic activity of the target enzyme. The latter group serves a dual function in that they serve as probes of mechanism as well as being labeling agents. As discussed in Section VIII, a common element of all recent hypotheses as to how acylCoA dehydrogenases bring about the oxidation of their thioester substrate is the abstraction of a proton from the a-carbon. Several mechanism-based inhibitors have been designed to test this hypothesis. 2. Acetylenic Inhibitors Gomes et al.15 investigated the effects of 3-butynoyl pantetheine and 3-pentynoyI pantetheine on glutaryl-CoA dehydrogenase (from Pseudomonas fluorescens) and butyryl-CoA dehydrogenase (from M. elsdenii). They found that both compounds inactivated both enzymes irreversibly, following first-order kinetics in those cases where the inactivation was slow enough to monitor conveniently. The absorption spectra did not suggest that the flavin prosthetic groups were modified, and in both cases the FAD was still reducible with dithionite. Fendrich and Abeles133 analyzed the time-dependent reaction of M. elsdenii BCD with 3-pentynoyl pantetheine in more detail. No more than a twofold molar excess of inhibitor over enzyme subunits was required to give 95% loss of activity. When 14C-labeled reagent was used, the label was incorporated into the protein. It could be removed from denatured protein by incubation either at alkaline pH or in hydroxylamine. The base-catalyzed hydrolysis released 2-oxopentanoic acid, and the NH2OH treatment formed a protein hydroxamate. The authors deduced there had been an ester formed between a protein carboxylic group and enolate. In order to identify the residue involved, the putative ester was reduced with NaB3H4 to release the 14C label and convert the carboxyl to a carbinol. After hydrolysis two peaks of 3H labeling were found that were absent in the control sample. The larger was 2-amino 5-hydroxyvaleric acid, the predicted carbinol derivative of glutamate; the other peak was proline. They concluded that the carboxyl group of a glutamyl side chain in the active site had been labeled.

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Using a similar approach at about the same time, Frerman and colleagues45 investigated the effects on pig MCAD of 3-butynoyl CoA. Again irreversible inhibition was obtained, displaying pseudo first-order kinetics with an apparent rate constant that showed saturation behavior. When ETF and an acceptor dye were present they were not reduced, so that apparently there is no partitioning between productive catalysis and suicide inactivation. The flavin extracted from inactivated enzyme was indistinguishable from normal FAD. Again it appeared that the suicide inactivator was modifying the protein rather than the prosthetic group. Both groups of workers surmized that inactivation proceeded via an initial proton abstraction at the a-carbon, leading to formation of an allenic intermediate. Frerman et al.45 therefore did the further experiment of making 2, 3-octadienoyl CoA and testing its behavior; it too gave rapid inactivation. Thorpe's group investigated the interaction of the same enzyme with acyl-CoA compounds in which the 2, 3 carbon-carbon bond, rather than the 3, 4, was acetylenic. They anticipated that 2-octynoyl CoA might inactivate without further activation by the enzyme. They found134 that there is indeed rapid, stoichiometric, irreversible inactivation, but the details were unexpected. Initial binding caused an immediate 10 nm red shift and increased resolution in the main visible flavin absorbance band, but thereafter a long-wavelength band emerged rapidly, centered at 800 nm and still associated with an oxidized spectrum in the 450 nm region. The long-wavelength absorbance declined over a period of an hour or so, and this was accompanied by the release of free CoA. Inactivation and covalent modification of the protein, however, occurred before the decline of the 800 nm band. The C(5) and C(15) homologues gave similar behavior, with 800 nm bands of lower intensity. With the C(15) compound it was particularly clear that the rise of the long-wavelength band was considerably faster than the inactivation. Powell and Thorpe135 pursued these interesting findings, using 14C labeling to establish the site of reaction with the protein, and deuterium labeling to probe its mechanism. Purification and analysis of a 13-residue tryptic peptide from a digest of the enzyme inactivated by [l-MC]-2-octynoyl-CoA established a unique site of modification, at Glu-401, to give an acid-stable but base-labile modification. Dideuteration at C4 produced a six- to sevenfold slowing both of inactivation and of the formation of the 800 nm band. The process was taken, therefore, to be mechanism-based, depending on rate-limiting removal of a proton from C(4). Powell and Thorpe135 speculate that the same active-site glutamyl residue may interact with both the a and ^ positions. Glu-401 would in that case be the base involved in initiating the normal catalytic reaction. From an inspection of amino acid sequences, Powell and Thorpe also note that there is a glutamyl residue in a somewhat similar conserved sequence in acyl-CoA oxidases: IleTyr-Glu in Candida acyl-CoA oxidase64 matches the same tripeptide sequence in pig mediumchain acyl-CoA dehydrogenase in an optimal alignment. The oxidases are also inactivated by 2-octynoyl CoA. The glutamate in question corresponds to residue 416 in the alignment of Matsubara et al.39 discussed in Section VI. If this is a key residue in catalysis the failure to conserve it in some of the other acyl-CoA dehydrogenases is clearly puzzling. 3. 3,4-Pentadienoyl CoA The use of an allenic acyl CoA — 2,3-pentadienoyl CoA — has already been described. As another mechanistic probe, Wenz et al.136 explored the 3,4-dienoyl isomer. This, they predicted, was unlikely to serve as a normal substrate since the corresponding product with three adjacent olefinic bonds would be energetically disfavored. The compound, however, gave rapid and potent inactivation: in a stoichiometric reaction over 1 to 2 min the visible absorption was bleached and activity was lost. This bleached enzyme species could be slowly reactivated by addition of a large excess of substrate, but did not itself react with the usual

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electron acceptors and was therefore presumed not to contain a normal reduced flavin. Rather the allene was thought to have reacted covalently with the FAD, possibly at the N(5) position of the isoalloxazine system. Attempts to release the flavin in a modified form failed, however, all yielding normal oxidized FAD. Without interference, the presumed adduct decomposes over a few hours, giving the conjugated 2,4-dienoyl-CoA and normal active, oxidized enzyme. However, up to 20% of the activity is irreversibly lost. It appears that 3,4-pentadienoyl CoA serves as a suicide substrate, partitioning 5 or 6 to 1 in favor of isomerization. 4. Methylenecyclopropyl Acyl-CoA Compounds These compounds are derived from plant toxins and their medical and metabolic background is given in detail in Section IX. The end result, however, is suicide inhibition and it is therefore appropriate to consider the chemistry here. The first of these compounds is methylenecyclopropylacetyl CoA. In vivo it is responsible for irreversible inhibition of shortchain acyl-CoA dehydrogenase and isovaleryl-CoA dehydrogenase. The studies in vivo have not in fact concentrated on these target enzymes but on what are presumed to be analogous situations, the choice being dictated by the availability of purified enzymes. Thus Ghisla et al.137 established that MCPA CoA inactivates butyryl-CoA dehydrogenase from M. elsdenii, bleaching the flavin spectrum and producing a covalent modification of the FAD. The same authors extended their study to medium-chain acyl-CoA dehydrogenase from pig kidney.138 This enzyme also was inactivated by a small molar excess of MCPA CoA with marked protection afforded by octanoyl-CoA and complete protection by anaerobic chemical reduction of the enzyme. Release of the prosthetic group showed that several different modified flavin products resulted. The small amounts of these and their instability have hindered positive identification. Even the use of enzyme reconstituted with modified flavins,139 restricting the number of reactive positions, has not altogether resolved the uncertainties. It seems clear, however, that activation of the pseudo-substrate by removal of an a-proton facilitates opening of the cyclopropane ring and generates a highly reactive intermediate with a number of options for covalent modification of the flavin. Methylenecyclopropylformyl CoA, one carbon unit shorter than MCPA CoA also appears to inhibit some of the acyl-CoA dehydrogenases (see Section IX), but these are apparently not its main site of metabolic inhibition, and the chemistry of the interaction with those acyl-CoA dehydrogenases that are affected has not been elucidated. 5. Propionyl CoA This compound is the most unexpected of the suicide substrates studied so far with this group of enzymes, the suicide reaction having been approached neither through inventive chemistry nor through toxicological detective work. Propionyl CoA was, on the face of it, merely a surprisingly poor substrate for the short-chain, acyl-CoA dehydrogenases30'92'100 and even a poor competitive inhibitor.45-100 Specifically, with beef short-chain acyl-CoA dehydrogenase, Shaw100'110-140 found that KM values for butyryl CoA and propionyl CoA were 6 \LM and 150 \iM, approximately and that Vmax for propionyl CoA was nearly 100fold less than the figure of 2.5 s ~ l for butyryl CoA. This striking difference lends significance to the evidence of Auer and Frerman141 for a substrate-induced conformational change (in the medium-chain acyl-CoA dehydrogenase). If the same process is part of the productive catalytic cycle for the short-chain dehydrogenase also, then evidently the lack of one methylene group makes a critical difference. As a competitive inhibitor, propionyl CoA also appeared to be a striking flop! 20 \LM propionyl CoA gave no detectable diminution of the rate measured with 2 jxA/ butyryl CoA as substrate. This might have been the end of the story had Shaw140 not noticed the marked curvature of reaction time-courses with propionyl CoA as substrate. With such low rates, this curvatun

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FIGURE 19. Absorption spectrum of flavin released from propionyl-CoA treated bovine butyryl-CoA dehydrogenase. The enzyme (13.4 \^M bound FAD) was treated with a fivefold molar excess of propionyl CoA for 20 min. The flavin was released with guanidinium hydrochloride at a final concentration of 5.25 M.

could hardly be attributed to an approach to equilibrium, and a second dose of enzyme gave an additive (rather than doubled) rate, establishing clearly that the first dose had largely been inactivated. In a more systematic investigation of this inactivation, 6.7 (jiM enzyme (subunit molarity) was incubated with 200 \LM and 500 \LM propionyl CoA at 25°C and sampled periodically for assay. A pseudo first-order decline of activity to an equilibrium value of 22% residual activity was seen with kapp of 7.2 x 10^ 3 s^ 1 for 500 jjiAf propionyl CoA. Lower concentrations of enzyme and substrate gave lower extents of inactivation; for example, 0.3 (o-Af enzyme with 0.6 (jiM substrate gave 57% residual activity at equilibrium. Tritiated CoA #as used to label the substrate and after inactivation the protein was precipitated, and repeatedly washed, with ammonium sulfate. The radioactivity incorporated in the precipitate was proportional to inactivation and indicated a 1:1 stoichiometry. Butyryl CoA protected very effectively. The effect of propionyl CoA on the absorption spectrum was quite unlike that of a normal substrate. There was extensive bleaching and no long-wavelength absorption, but instead a new band was seen at 335 nm. Successive spectra gave a good isosbestic point, indicating only two significant species present. In long-term incubations there was a slow reversion towards a normal flavoprotein spectrum. The reversion could be accelerated by using a high concentration of an avid ligand,100 namely acetoacetyl CoA. This is in keeping with the equilibrium nature of the inactivation, and accordingly the assays also show a distinct upward curvature — i.e., inactivated enzyme gradually reactivates in the presence of a large excess of good substrate. Kinetic analysis of the spectral changes gave rate constants in excellent agreement with those from activity assays, both indicating a partitioning between productive catalysis and a suicide reaction in the ratio 4.3:1. It seemed likely that a covalent modification of the flavin was occurring, possibly by formation of an N(5) adduct. Flavin was therefore released with guanidinium hydrochloride, detergent, or trichloracetic acid; in each case giving a pink solution with a spectrum (Figure 19) reminiscent of that of a neutral flavin semiquinone. EPR measurements, initially in the laboratory of Professor N. M. Atherton and subsequently with Dr. R. C. Bray, confirmed that this was a stable radical, and integration of the signal at g=2.0037 indicated about 90% conversion of modified flavin

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into free radical. This radical species proved to be sufficiently stable to purify by HPLC, allowing confirmation that the radical was associated with the radioactivity derived from the labeled propionyl Co A. The obvious analogous models for this system are those provided by the reoxidation of reduced yV-5-alkyl-lumiflavins, 142144 also generating air-stable radicals. With the propionylCoA treated enzyme, no radical signal is seen until the flavin is released, and no radical appears if the release is under anaerobic conditions. In order to assign the structure of the radical, a detailed study was carried out by George et al.145 involving the use of propionyl CoA specifically deuterated at position 2 or position 3 and also of propionyl CoA enriched with 13C at C(2) and separately at C(3). Four principal possibilities were envisaged: namely, propionyl-CoA or acrylyl-CoA adducts attached to the N(5) atom of the flavin via either C(2) or C(3). The detailed arguments are beyond the scope of the present review, but the conclusion was drawn that the adduct contained an acrylylCoA moiety attached via C(2) of the acyl chain. The mechanism of its formation is not yet entirely clear. 6. Photoaffinity Labeling It is well established that medium-chain acyl-CoA dehydrogenase will tolerate bulky substituents on carbon 3 of the acyl chain. Thus phenylpropionyl CoA82 and |3-(2-furyl) propionyl CoA146 are both good substrates. Frerman and Turnbull147 have taken advantage of this property in designing two photoaffinity ligands, /?-azidophenylpropionyl CoA and 4Af-(4-azido-2-nitrophenyl) aminobutyryl CoA. The first of these is a good substrate, following the typical pattern of reaction with the enzyme seen with straight-chain substrates. The unsaturated product forms a charge-transfer band with maximal absorbance at 700 nm, in keeping with the electron-withdrawing properties of the azidophenyl group. The product itself has an absorbance band at 340 nm and irradiation at this wavelength results in firstorder inactivation which can be prevented by adding octanoyl CoA. Irradiation alone, without the photoaffinity ligand, has little effect. Work to identify the modified peptide is in progress.

VIII. MECHANISM A. INTRODUCTION As with most flavoproteins, the acyl-CoA dehydrogenase reactions can be divided into separate partial reactions, a half-reaction in which acyl-CoA substrate reduces enzyme-bound FAD, and a half-reaction in which the oxidized flavin is regenerated. The second phase, as we have seen in earlier sections, is brought about by another flavoprotein, ETF. As ETF has a chapter to itself elsewhere in this review series,20 the present discussion will not concern itself with the reoxidation. The acyl-CoA dehydrogenases have been part of the known biochemical world for nearly 40 years. In retrospect, in considering the mechanism of their action, it is remarkable how deep the first workers dug. The extensive series of papers from the laboratories of David Green and Helmut Beinert1'19 exposed most of the main questions and made a remarkable start on providing the answers. As mentioned earlier, the 1980s saw a major revival of interest in this group of enzymes. In terms of furthering our understanding of their mechanism in some depth, one contribution stands out among many. The group of Colin Thorpe at Delaware has made elegant and systematic use of synthetic chemistry to create a series of probes of different aspects of the mechanism of the medium-chain acyl-CoA dehydrogenase of pig kidney, clearing up previous confusion and producing a satisfying and consistent picture of the mechanism. Before tackling the heart of the reaction mechanism, however, we turn first to two other related matters, oxidation-reduction thermodynamics and stereochemistry.

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B. OXIDATION-REDUCTION PROPERTIES As discussed in Section I, different acyl-CoA dehydrogenases have to function either in the direction of saturated acyl-CoA oxidation or in the direction of enoyl-CoA reduction. It is clearly essential for efficient oxidation-reduction catalysis that the potential of the enzyme-bound flavin be suitably poised relative to that of the substrate. The EO for the butyryl CoA/crotonyl-CoA couple was first estimated by Hauge50 as - 15 mV by establishing equilibrium with pyocyanine. Subsequently Gustafson et al.148 obtained a much more negative value, — 126 mV. Redetermination by spectroelectrochemical methods149 in Marion Stankovich's laboratory gave a value of - 19 mV, essentially confirming that of Hauge. Fink et al.93 have determined the midpoint potential for butyryl-CoA dehydrogenase from M. elsdenii as — 79 mV. This enzyme has to reduce crotonyl Co A in vivo,25 and therefore, the value of Gustafson et al. for the potential of the substrate couple might imply an uphill task. The higher values remove this apparent difficulty. Addressing the problem, nonetheless, Fink et al.93 also note that binding of the competitive inhibitor acetoacetyl CoA44 lowers the oxidation-reduction potential of the enzyme flavin to -180 mV. They speculate, therefore, that the structurally similar substrate crotonyl CoA may modulate the potential of the enzyme in the appropriate direction to ensure efficient transfer of reducing equivalents. The mitochondrial p-oxidation enzymes function, of course, in the opposite direction and the opposite problem exists. For the medium-chain acyl-CoA dehydrogenase, Lenn et al.150 report an EQ of -135 mV, in good agreement with Gustafson et al.,144 whereas for the C(4), C(8), and C(16) substrate couples, they give values of -39, -36, and -36 mV, respectively, determined with the medium-chain enzyme. The potential was remeasured for the enzyme itself in the presence of an excess of a mixture of substrate and product; this time values of -33, -33, and -68 mV were obtained in the presence of C(4), C(8), and C(16) acyl-CoA ligands. In other words, as with the bacterial enzyme, the substrate again appears to alter the poise of the oxidation-reduction equilibrium, this time in the opposite sense, so that the oxidation-reduction potential of the enzyme is well matched to that of the substrate couple. One other interesting point in relation to the regulation of oxidation-reduction potential emerges from the work of Madden et al.151 They note that the variation in E'0 values for various flavoproteins must reflect differences in the tightness of binding of the reduced and oxidized forms of the flavin by the apoprotein. In the case of medium-chain, acyl-CoA dehydrogenase, the enzyme (even without substrate bound) is a stronger oxidant than free FAD (-208 mV152) by about 80 mV, and therefore the enzyme should stabilize reduced FAD, relative to oxidized FAD. Madden et al. point out that the converse must also be true, and they duly demonstrate dramatic stabilization of the protein against heat or urea denaturation. In 7.3 M urea at 25°C the oxidized enzyme lost virtually all activity within 4 h. In the presence of an excess of substrate over 60% of the activity was retained, but simple reduction with dithionite resulted in retention of 97% activity in 7.3 M urea after 4 h. In summary, this family of enzymes shows clearly how the protein can modify the oxidation-reduction properties of flavin and how this ability can be further modulated by substrate or substrate-like molecules. C. STEREOCHEMISTRY The substrates for the straight-chain, acyl-CoA dehydrogenases have methylene groups at the a and (3 positions and thus there are two choices at either position for the hydrogen to be removed. The first published work on the stereochemistry was that of Biellmann and Hirth who in separate experiments labeled the (3-carbon153 and the a-carbon154 stereospecifically with tritium. Tritium at the S position on the p-carbon was mostly retained, whereas tritium at the R position was largely released into the water when the labeled substrates

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were used with the short-chain acyl-CoA dehydrogenase from pig liver. The result reported was not entirely clean, but apparent leakage from the S position was attributed to partial racemization during the chemical synthesis from butyric acid. The experiments with butyryl CoA labeled at the a-position again indicated that it is the pro-R hydrogen which is removed. Cleaner labeling patterns were obtained some years earlier by Drysdale155 using an exclusively enzymatic route in labeling the substrate. The enoyl CoA reductase of yeast was used either with tritiated NADPH or unlabeled NADPH in tritiated water to label at C(3) or C(2), respectively. When the tritiated butyryl CoA was used as a substrate for bovine short-chain acyl-CoA dehydrogenase, the label at either position was released. This indicated a stereospecific reaction involving the same hydrogen positions as those involved in the enoyl-CoA reductase reaction, but the actual positions were not identified. In further experiments, however, it was found that only the S-isomer of 2-methylbutyryl CoA would serve as a substrate for the short- or medium-chain, acyl-CoA dehydrogenase. This leaves no doubt that the enzyme removes the pro-R hydrogen at C(2). For the medium-chain acylCoA dehydrogenase the stereochemistry at C(3) was also established by using the two C(3) tritiated isomers of nonanoyl CoA. Again pro-R removal was found. In the case of the M. elsdenii short-chain, acyl-CoA dehydrogenase results consistent with this stereochemistry at least for the 3-carbon were obtained by G. Williamson in my laboratory156 by examining the specificity for the two enantiomers of p-hydroxybutyryl CoA which is slowly oxidized to acetoacetyl CoA by the flavoprotein.44 More recently, detailed stereochemical studies have been carried out for the branchedchain, acyl-CoA dehydrogenases. Aberhart and Tann157 took advantage of the exclusive oxidation of isovaleryl CoA by its own dehydrogenase to perform a stereochemical experiment in vivo. Stereospecifically C(2) tritiated isovaleric acid was administered to biotindeficient rats and the resulting hydroxyisovaleric acid — following oxidation, hydration, and hydrolysis — was analyzed to show removal of the pro-R hydrogen. Nonstereospecific hydration precluded a similar approach at C(3), and the determination of stereochemistry at this position was approached with purified enzyme;158 this time using stereospecific 13C labeling of one of the methyl groups as the reporter for the stereochemistry of the product. This showed that the dehydrogenase carries out anti-elimination of the hydrogen at C(2) and C(3) in keeping with the pattern seem with the straight-chain, acyl-CoA dehydrogenases. In similar studies with the 2-methyl branched-chain acyl-CoA dehydrogenase,15913C labeling was again used; this time coupled with mass spectrometric analysis of the products. In isobutyryl CoA, the substrate used, the hydrogens on each methyl group are equivalent but the methyl groups themselves are not. The study showed removal of the hydrogens from the a-methine ()CH) and the (pro-2S)-methyl. A consistent picture of stereochemistry emerges, irrespective of substrate chain length or biological species, as might reasonably be expected for a homologous family of enzymes. Finally it must be mentioned that pig MCAD is one of a small group of flavoproteins for which the absolute stereospecificity of reduction of the flavin ring system has been determined.160 For NAD(P)+-dependent dehydrogenases such experiments were performed a quarter of a century earlier, but the lability of N-H bonds in reduced flavin molecules appeared to make this impossible for flavoproteins. However, the use of 8-demethyl-8hydroxy-5-deaza-5-carba flavin analogues has overcome this problem for flavoproteins that can be resolved to give apoprotein and then reconstituted with the appropriate FMN or FAD analogue. The carbon at position 5 gives a cofactor which, like NAD + , does not exchange the hydrogen introduced by enzymatic or chemical reduction. The absolute reference point is glutathione reductase, in which it can be clearly seen from the crystal structure that the exposed re face of the flavin is used for reduction. Pig MCAD reconstituted in this way was reduced with NaB3H4 and the stereoselectivity reduced flavin released and used to reconstitute the apoenzyme of glutathione reductase. Nearly all the label was released upon oxidation by NAD + , proving that the two enzymes use the same face of the flavin ring.

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D. MECHANISM OF HYDROGEN REMOVAL The reaction catalyzed by acyl-CoA dehydrogenase involves the breaking of two-carbon hydrogen bonds to create a double bond, and one of the central questions has been how these kinetically stable bonds are broken and in what form the hydrogens are removed. The free carboxylic acids are chemically rather inert, and this problem is overcome by the attachment in thioester linkage to Coenzyme A. One consequence of this linkage is enhanced acidity of the a-protons. Accordingly, in the first specific proposal for a chemical mechanism for the acyl-CoA dehydrogenases, following upon the detailed studies of Beinert and Page,47 Cornforth161 suggested that a base provided by the protein might abstract the first hydrogen as a proton from the a-position. Thereafter the flavin has to receive the equivalent of a hydride ion. Various routes can be envisaged, including a transitory covalent addition to form a flavin adduct,162 a radical-pair mechanism as proposed by Cornforth, or direct transfer of the hydride ion. As we have seen in Section V, however, there is no good evidence for a radical species participating in catalysis. There is a basic philosophical problem in assessing much of the evidence. Many authors have sought to simplify the overall picture in various ways to solve the simultaneous equations, as it were, by eliminating some of the variables, usually by employing a modified substrate. One then has to decide, however, whether the results and conclusions reached from the use of each substrates are really representative of what happens with a good substrate or whether they represent' 'derailment' * mechanisms. For example, if we take the hypothesis that catalysis involves a covalent intermediate,162 the experiments of Shaw and Engel140 with propionyl Co A, a very poor substrate for beef liver butyryl-CoA dehydrogenase, clearly provide evidence for the formation of a covalent adduct. This adduct, moreover, ultimately breaks down to regenerate an unchanged enzyme plus a product. Does this represent a greatly slowed down version of the efficient catalysis seen with a good substrate or is it the disastrous consequence of failure to achieve the normal catalytic route? The current consensus would certainly favor the latter view; i.e., that the activated intermediate in catalysis can collapse in various directions if denied the normal productive route. There appears to be universal agreement over the involvement of a carbanion formed by abstraction of an a-proton. The experiments reported in Section VII with acetylenic and allenic substrate analogues were all designed to test this hypothesis and they provide evidence in its support. Moreover, the protein chemical studies of Fendrich and Abeles133 and of Thorpe et al.135 focus attention on a glutamyl side chain which could serve as the base in such a process. The only puzzling feature here, as noted in Section VI, is the lack of conservation or even conservative substitution at this position in at least two members of the enzyme family which presumably share a very similar mechanism. The role of the glutamate could be taken over by another residue, but this issue has not yet been resolved. Negatively charged acyl CoA species or analogues in the active site of acyl-CoA dehydrogenase tend to give charge-transfer bands, archetypes being the enzyme acetoacetylCoA complex and the green CoA-persulfide complex, both discussed in Section V. In studies with the medium-chain acyl-CoA dehydrogenase from pig kidney, Thorpe's group has provided a couple of striking examples of such charge-transfer complexes. The first, reported by Powell et al.,163 is an interaction with trans-3-octenoy\ CoA, which produces an intense absorbance band centered on 820 nm. The authors show by nuclear magnetic resonance (NMR) that there is indeed rapid exchange of a proton at the a-position with D2O, and conclude that the likely charge-transfer donor is the delocalized carbanion.

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FIGURE 20. The complex formed by pig medium-chain acyl-CoA dehydrogenase with 3-thiaoctanoyl CoA. The solid line ( ) is the spectrum of the uncomplexed enzyme (11 \^M) and the broken line (—) is the spectrum of the complex formed upon addition of a twofold excess of the substrate analogue. (From Lau, S.-M., Brantley, R. K., and Thorpe, C., Biochemistry, 27, 5089, 1988. With permission.)

They note the strong similarity with a spectrum reported in our own studies for a complex between butyryl-CoA dehydrogenase and fra/w-2-pentenoyI-CoA44 and speculate, almost certainly correctly, that our results followed upon an isomerization interconverting 2-pentenoyl CoA and 3-pentenoyl CoA. In later, unpublished experiments we indeed obtained an identical spectrum with 3-pentenoyl CoA but attributed the result to a possible impurity rather than isomerization! There is no secondary reduction step following the formation of these complexes and the absorbance in the 350 to 450 nm region is that of an oxidized, if slightly blue-shifted, flavin. The second example comes in an account by Lau et aL164 of the effects of various octanoyl-CoA analogues in which carbon atoms in the acyl chain have been replaced by oxygen or sulfur. The 3-thia analogue gives a stable absorbance band with an extinction coefficient at 804 nm of 8.7 mM" 1 cm" 1 on binding to the enzyme (Figure 20). Again the a proton is exchangeable with deuterium allowing the possibility that an enolate with the negative charge on the thioester carbonyl function is the charge transfer donor. This analogue is ideally suited for testing events at the a-carbon atom in a situation where there is no possibility of reduction since the sulfur at position 3 has no hydrogens to be removed. In fact analogous oxidized charge-transfer complexes do not seem to be a major contributor to the spectral species seen in rapid reaction studies of enzyme reduction by substrate and subsequent turnover.165-166 The mechanism put forward by Schopfer et al.166 in a closely reasoned and detailed paper to account for the rapid-reaction findings (Scheme 1) includes a Michaelis complex between oxidized enzyme and reduced substrate, but this initial complex is assumed to have no long-wavelength absorption, and the next species in the mechanism is already in the reduced-flavin state. Their interpretation is in line with several studies82-167-168 indicating that deuteration at both carbon 2 and 3 produces very large isotope effects on rates both of reduction and of formation of long-wavelength bands. Each position separately gives a substantial isotope effect, and the fact that the combined effect is more or less multiplicative argues strongly that the removal of hydrogen from the two positions is a concerted process.

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SCHEME 1. GAD denotes '' general acyl CoA dehydrogenase" (MCAD). Subscripts0 and r indicate "oxidized" and "reduced", respectively. B-CoA is butyryl CoA and C-CoA is the oxidized product crotonyl CoA. MC, and MC2 are Michaelis complexes. (From Schopfer, L., Massey, V., Ghisla, S., and Thorpe, C., Biochemistry, 27, 6599, 1988. With permission.)

This general conclusion should perhaps be qualified somewhat in view of Murfin's study82 of reduction of beef medium-chain acyl-CoA dehydrogenase by phenylpropionyl CoA which includes a careful survey of pH effects. He reports multiplicative effects of deuteration at the two carbon positions at high pH (9.5) but finds an isotope effect only at carbon 3 at low pH (6). The conclusion, for this substrate at least, is that at low pH the proton abstraction is rapid compared to hydride transfer and separated from it; whereas at high pH they are simultaneous steps. It is thus important to bear in mind the dangers of a too dogmatic generalization about the mechanism. There may well be significant differences in the rate-determining step, and therefore kinetically significant observable species, depending on the source and substratespecificity of the enzyme, the choice of substrate, and the precise experimental conditions. Nevertheless very considerable progress has now been made in unraveling the mechanism of the reductive half-reaction. Probably the most direct evidence so far bearing on the mechanism of hydrogen removal comes from Ghisla et al.169>17° Following up the study of Gomes et al.15 on exchange at the a and (3 carbons of butyryl CoA catalyzed by the BCD of M. elsdenii, they established the stoichiometry of exchange in 3H2O as 1.94 per substrate molecule. This was kinetically monophasic, implying a concerted process. NMR and mass spectrometric measurements following exchange in D2O confirmed that one atom was exchanged at the a-position and one at the p-position. With the pig medium-chain enzyme, on the other hand, only the exposition exchanged. Direct transfer of hydrogen between the substrate and the flavin was demonstrated by making use of 5-deazaFAD. This analogue, in contrast to normal FAD, does not exchange hydrogen at position 5 of the ring system since the nitrogen is replaced by carbon (see Section VIII.C above). Ghisla et al. examined transfer of reducing equivalents from flavin to substrate in two ways; first of all, free 5-deazaFAD was reduced with borotritide and used to reconstitute MCAD which was then mixed with crotonyl CoA. Roughly half the counts ended up in the product butyryl CoA and half in the enzyme, since only one of the two positions at C(5) could transfer tritium to the substrate. On the other hand, when the flavin analogue was reduced with NaB3H4 on the surface of the enzyme after reconstitution, and crotonyl CoA was then added, nearly all the label (91%) went into the butyryl CoA. It was also established that this label was on the p carbon and that it could be removed by the M. elsdenii enzyme, proving that they share the same stereospecificity. The rate of reduction of crotonyl CoA by the reduced 5-deazaFAD enzyme was shown to be similar to that for the native enzyme, making it unlikely that the view of the mechanism provided by the analogue is distorted by major changes in rate-limiting steps or in the catalytic behavior of the prosthetic group. E. SIDE ACTIVITIES Some of the ' * side activities'' of acyl-Co A dehydrogenases have already been discussed. Thus we have seen (Section IV) that butyryl-CoA dehydrogenase from M. elsdenii can

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function as an oxidase. McFarland et al.146 found the same with furyl propionyl CoA as a substrate for medium-chain, acyl-CoA dehydrogenase from pig liver. Likewise we have referred to the fact that the short-chain dehydrogenase catalyzes an isomerization of enoyl CoA. It is able, for example, to interconvert crotonyl CoA and vinyl acetyl CoA.44 This activity seems to be diminished or missing in the mammalian medium-chain, acyl-CoA dehydrogenase.161 It is also worth recalling that the M. elsdenii butyryl-CoA dehydrogenase catalyzes a slow oxidation of p-hydroxybutyryl CoA in the absence of NAD + to produce acetoacetyl CoA.44 In our studies with acetoacetyl thioesters of various thiols we noted a conversion of acetoacetyl ethanethiol plus Coenzyme A to acetoacetyl CoA plus ethanethiol. In view of Steyn-Parve and Beinert's long-standing observation of labilization of the thioester linkage by acyl-CoA dehydrogenase,48 it seemed possible that the enzyme catalyzed thiol-thioester exchange. Patricia Ellison in my laboratory pursued this171 however, and showed that the nonenzymic process is sufficiently rapid to account for the observed exchange. There is thus no firm evidence to suggest that this is a feature of the enzyme. Another possible side activity to emerge from early studies44 was enoyl-CoA hydratase. Such activity had to be present in order to explain how butyryl CoA could gradually be converted to acetoacetyl CoA in the presence of high concentrations of bacterial butyrylCoA dehydrogenase. However, low levels of contamination by a separate hydratase would have been sufficient to account for this activity. Such contamination is usually present but it can be destroyed by taking advantage of differential susceptibility to denaturation by urea.172 There then remains an irreducible level of endogenous hydratase activity associated with the dehydrogenase. Photooxidation which inactivates the dehydrogenase173 gives parallel inactivation of the residual hydratase activity.172 The belief that this is an intrinsic property is strongly reinforced by the findings of Lau et al.174 who found this activity in the pig kidney medium-chain acyl-CoA dehydrogenase. In their study, hydratase activity was destroyed by removal of FAD and by mechanism-based inhibitors of the dehydrogenase. Interestingly (and fortunately from the standpoint of binding studies) the hydratase activity is seen with a C(4) substrate but not with frans-2-octenoyl CoA. Lau et al.174 propose that the same base responsible for proton abstraction in the dehydrogenase reaction may also catalyze the hydration of bound enoyl CoA.

IX. MEDICAL ASPECTS A. INTRODUCTION Since (3-oxidation provides one of the two major energy sources for man and other mammals, its inhibition or interruption is not a trivial metabolic problem. The repetitive nature of the pathway, however, and the matching multiplicity of the enzymes that catalyze the 4 chemical conversions that recur down the spiral, provide a degree of insurance. Paradoxically, it is precisely for this reason that there is a voluminous literature on clinical disturbances of (3-oxidation. If, for example, inborn errors affecting this pathway were invariably fatal in utero, they would not have attracted attention. Instead, because the overlap in chain-length specificity provides partial "cover", genetic defects cause illness of varying severity, in some cases episodic with long intervening periods of apparent normality. These conditions have attracted increasing attention in recent years175 as their prevalence has been revealed. Insight into the consequences of genetic enzyme deficiency is also coming from the study of the analogous effects of specific inhibition by toxins. B. JAMAICAN VOMITING SICKNESS Consumption of the unripe fruit of the ackee (Blighia sapidd) causes a serious clinical

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FIGURE 21. Comparison of the molecules of the plant toxin hypoglycin A and leucine, and of their respective metabolic products, methylenecyclopropyl acetyl CoA and isovaleryl CoA. In both cases the conversion proceeds via transamination to give the 2-oxo acid which is then oxidatively decarboxylated by the branchedchain oxoacid dehydrogenase complex.

condition involving violent vomiting, coma, and occasionally death.176"178 Biochemical symptoms include a dramatically depressed blood glucose level, and, in the liver, depleted glycogen and fatty infiltration. The active principle of the ackee fruit was isolated and identified by Hassall and Reyle179 as hypoglycin A, an a-amino acid with a methylenesubstituted cyclopropyl group in its side chain (Figure 21). The hypoglycemic properties of this amino acid attracted the attention of pharmaceutical companies searching for effective antidiabetic agents, but it gradually became clear that the basis of its action made it entirely unsuitable for such use. This compound is so similar to leucine (Figure 21) in its size and physical properties that the two are difficult to separate,180'181 and the metabolic systems of the body experience the same difficulty. Hypoglycin A is processed as if it were a normal branched-chain amino acid, first by transamination to give the oxoacid and then, by oxidative decarboxylation, to give methylenecyclopropylacetyl CoA (MCPA CoA).182 These metabolites are the true inhibitors.183 This is a good example of what Peters184 termed "lethal synthesis1', in which a compound, innocuous in itself, is metabolized to give a potent inhibitor. Hypoglycin A poisoning strongly inhibits fatty acid oxidation,185'186 and while some of the hypoglycin metabolites may also inhibit gluconeogenesis, a major reason for the severe hypoglycemia is that by blocking fat oxidation, the inhibitor forces all tissues to draw upon glucose that would otherwise be spared for those tissues, e.g., the brain, which depend exclusively upon it. von Holt and von Holt185 showed that riboflavin phosphate alleviated the symptoms of hypoglycin poisoning in rats, and in an inspired guess suggested that the acyl-CoA dehydrogenases might be the site of inhibition. Kean186 reported that acyl-CoA dehydrogenases isolated from hypoglycin-treated rabbits showed specific inhibition of the short-chain dehydrogenase. A possible mechanism for such an effect emerged with the developing understanding of suicide inhibitors.131'132 MCPA CoA was tested in in vitro studies with both pig medium-chain acyl-CoA dehydrogenase137 and M. elsdenii short-chain acyl-CoA dehydrogenase.138 In both cases "irreversible" inhibition resulted — in the sense that it was not counteracted by dialysis. However, the detailed analysis of the inactivated pig enzyme revealed that is primarily the FAD rather than the protein that has been covalently modified, thus rationalizing the previously puzzling obser-

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vations on riboflavin feeding. Several different modified flavin molecules were tentatively identified. As pointed out, however, by Sherratt187189 in a series of reviews on this subject, over two decades,187"189 it has always seemed likely that hypoglycin A metabolites have several distinct sites of action. It was noted already in 1967 by Posner and Raben190 that oxidation of leucine is inhibited by hypoglycin A and, in an ensuing series of papers, Tanaka and colleagues33-34 maintained that MCPA CoA inhibited specific branched short-chain acylCoA dehydrogenases, especially isovaleryl-CoA dehydrogenase. They proposed indeed that Jamaican vomiting sickness served to some extent as a model for the genetic defect known as isovaleric acidemia. At that stage only indirect evidence existed for separated branchedchain, acyl-CoA dehydrogenases, as noted35 by this reviewer. Nevertheless, Tanaka's group have gradually accumulated overwhelming evidence in favor of their hypothesis and there is now no doubt that MCPA CoA covalently inactivates isovaleryl-CoA dehydrogenase as well as butyryl-CoA dehydrogenase. Finocchiaro et al.40 have purified short-chain and medium-chain acyl-CoA dehydrogenases and also isovaleryl-CoA dehydrogenase from human liver and have shown that in vitro, all three are severely inhibited by MCPA CoA. In addition to Hypoglycin A (methylenecyclopropylalanine) the ackee fruit produces smaller quantities of a homologue shorter by one methylene group, methylenecyclopropylglycine. This is also toxic191-192 and is present in much higher concentrations in the kernel of lychees. There is no corresponding ''Chinese vomiting sickness" because these kernels are not eaten and the edible soft parts of the lychee fruit are nontoxic. The hypoglycemic effects of methylenecyclopropylglycine have been demonstrated in both mice191 and rats.193 Enzyme assays show no inhibition of general acyl-CoA dehydrogenase in liver mitochondria from rats treated with this compound but there was marked inhibition of the a-methyl branched-chain acyl-CoA dehydrogenase. Also, in vitro, the presumed toxic metabolite, methylenecyclopropylformyl CoA, inactivates enoyl-CoA hydratase. This clearly would be an alternative way of halting p-oxidation. There has, however, been a gap in the chain of evidence until very recently. Even though MCPA CoA inhibits medium-chain acyl-CoA dehydrogenase in vitro, the failure of methylenecyclopropylglycine to do so in vivo is not strictly relevant since this is not the site of in vivo inhibition for MCPA CoA either. Now, however, a new detailed metabolic study has been reported on the effects of methylenecyclopropylglycine administered intraperitoneally to fasted rats.194 Activities of 12 different enzymes were measured, and it emerges that, apart from a slight inhibition (25%) of the short-chain acyl-CoA dehydrogenase, there is no significant inhibition of either the acyl-CoA dehydrogenase or enoyl-CoA hydratase steps of p-oxidation. p-Oxidation is nevertheless totally blocked, and this appears to be mainly due to severe inhibition of the thiolytic cleavage step. Inhibition of the branched-chain acyl-CoA dehydrogenases was also confirmed. The contrast between the final outcomes in these two related examples of lethal synthesis is striking and instructive. C. INBORN ERRORS OF METABOLISM The field of inborn errors of fatty acid metabolism175-195 is made obscure for the beginner by the nomenclature, based on secondary consequences rather than immediate causes. The causes inevitably are the last piece of the jigsaw. Disease conditions have been traditionally classified according to the analytical methods used by chemical pathologists. These tend to be measurements in blood and urine of the levels of glucose, ketone bodies, amino acids, and other organic acids. Conditions are thus described as nonketotic hypoglycemia, ethylmalonicaciduria, glutaric aciduria Type II, etc. In general, the markers picked up by the clinicians are alternative metabolites produced as a result of a metabolic blockage and also conjugation products of these metabolites with carnitine and glycine. Thus, blockage of the main p-oxidation pathway leads to an overspill into the pathways for co-oxidation (hence, adipic acid and other dicarboxylic acids) and for propionate metabolism (hence, ethylmalonic acid).

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FIGURE 22. A graph of the reports of fatty acid oxidation defects. The diagram indicates the increasing pace of discovery of such disorders and shows that all three straight-chain acyl-CoA dehydrogenases have been implicated. (From Matsubara, Y., Indo, Y., Naito, E., Osaza, H., Glassberg, R., Vockley, J., Ikeda, Y., Kraus, J., and Tanaka, K., /. Biol. Chem., 264, 16321, 1989. With permission from Wiley-Liss Inc.)

These traditional methods of diagnosis are of course indirect and often ambiguous as compared with direct enzyme assay or DNA-based methods, for example. The first clearly documented case of a (^-oxidation defect was wrongly attributed as a result of this ambiguity. The infant in question was diagnosed with a condition vividly described as the "odor-ofsweaty-feet syndrome".195 The apparent identification of large amounts of butyric acid led to the suggestion that this was due to a defect in the short-chain acyl-CoA dehydrogenase, The organic acid identification was, however, based only on GLC evidence, and members of different homologous series can give overlapping peaks, in this case butyric and isovaleric acids. A reinvestigation of the same patient several years later197 with the additional aid of mass spectrometry proved that this was after all a case of isovaleric acidemia similar to that previously described by Tanaka and colleagues.32 Subsequently, as a result both of greater awareness and of better instrumentation, other cases have come to light, and as shown in Figure 22, disease conditions have been associated with a number of different acyl-CoA dehydrogenases and also ETF. There are a number of other diagnosed cases affected elsewhere in the oxidative machinery, either mitochondrial or peroxisomal,175 but they fall outside our present scope. D. MEDIUM-CHAIN ACYL-CoA DEHYDROGENASE DEFICIENCY A major focus of interest in recent years has been medium-chain acyl-CoA dehydrogenase deficiency. There are several reasons for this. First of all it appears that of the 3 acyl-CoA dehydrogenases involved in |3-oxidation of straight-chain fatty acids, the medium-chain is the commonest site of inborn errors. This is presumably a consequence of the pattern of overlap in substrate specificities. All even-chain fatty acids have to pass through the C(4) stage, and the medium-chain enzyme cannot deal with C(4). Even though, under some circumstances of assay in vitro with the C(4) substrate alone, the medium-chain enzyme can contribute 50% of the observed activity;198 this is most unlikely to be the case in vivo in the

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face of competition from the physiological substrates. Hence, a defect in the short-chain acyl-CoA dehydrogenase is likely to be severe in its consequences. It will therefore also tend to be rare. Long-chain fatty acids are the major input into the fatty acid oxidation system in man and the medium-chain enzyme will not deal with C(16)-C(20). Peroxisomal oxidation may help out, but even so, deficiency of the long-chain acyl-CoA dehydrogenase has a high mortality rate. On the other hand, the fatty acids that the medium-chain, acylCoA dehydrogenase is adapted to handling can probably at a pinch be dealt with by one of the other two enzymes. It follows that deficiency of the medium-chain enzyme is milder and therefore more common. In fact this deficiency may remain undiscovered and asymptomatic for long periods. Affected individuals may be thrown into a metabolic crisis by unaccustomed fasting or by a minor infection, and medium-chain acyl-CoA dehydrogenase deficiency is therefore a significant cause of "cot death" or "sudden infant death syndrome" (SIDS). It is important to bear in mind that these clinical labels do not describe a single disease; they are merely administrative pigeon-holes for unexplained death. The cot-death correlation emerged by chance: Dr. Lee Shaw, carrying out assays in my laboratory for a suspected case of the short-chain acyl-CoA dehydrogenase deficiency in a collaboration with our local chemical pathologists,199'200 used as controls a series of cotdeath liver samples (not truly normal, but at least not homogeneously abnormal) previously studied in the Sheffield Cot-Death Project. Medium-chain acyl-CoA dehydrogenase was assayed as an internal reference, and one of the "control" samples turned out to be totally devoid of this activity. This was a case in which histology had shown severe fatty infiltration of the liver, quite commonly seen at post-mortem examination of cot-death cases. The ensuring retrospective study202'203 produced tentative evidence that medium-chain acyl-CoA dehydrogenase might be responsible for 5 to 10% of the cot-death cases in the Sheffield study. However, the study could be questioned on various counts: of the total sample, only a few had sufficient liver kept for enzyme analysis; the samples had been stored frozen for varying times up to 5 years post-mortem; little was known of the time interval and conditions between death and the freezing of the tissue. This retrospective study nevertheless triggered a prospective survey.204 Although this does not so far appear to bear out the high level of incidence implied by the initial study, subsequent reports from a number of laboratories in the countries do suggest a very significant contribution of MCAD-deficiency cases to their total SIDS numbers.205'207 This study led also to a successful prenatal diagnosis208 of an affected baby in a case where there had been a previous cot death in the family. The mother elected against termination. The diagnosis was confirmed on the newborn baby, but with simple dietary management (frequent feeding with a high-carbohydrate, low-fat diet), the child has remained healthy. In view of the fact that this condition (1) can be easily managed if recognized; (2) can otherwise be fatal; simple methods of detection and diagnosis are particularly important. Methods are required both for post-mortem diagnosis (to guide genetic counseling) and for diagnosis of live offspring either at birth or in utero. There are a number of metabolites in blood and urine that are regarded as indicative of MCAD deficiency, but this type of diagnosis has serious drawbacks. Notably, there is little agreement about what constitutes the range of variability in normal individuals, and there can certainly be large fluctuations in single individuals depending on their physiological and nutritional state. Measurements of fatty acid oxidation are also used, but these alone do not pinpoint a defect. Ideally, it is clearly preferable to have primary evidence of the presence or absence and functional state, if present, of the enzyme protein or its encoding gene. Starting with the protein, one may apply two types of criteria: immunological and enzymological. With a monospecific antibody it is possible to carry out a Western blot analysis of SDS gels of a crude cell extract. This reveals not only whether the protein is

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present but also whether the subunits are of the correct molecular weight. In fact, in those cases that have been examined to-date it appears that defects are due to point mutations. Thus, Tanaka and colleagues209 examined material from 13 patients. Cells were labeled with [35S] methionine and the MCAD was immunoprecipitated and run on an SDS gel. In all cases both the mature enzyme and its precursors were of the correct lengths. This suggests that, in general, polyclonal antibodies are not likely to be of much help in diagnosing these conditions. Nevertheless, Strauss et al.210 report an exceptional case. In their studies a different labeling procedure was employed. Western blots of SDS gels of cell lysates were treated with antibody and then with protein A, labeled with radioactive iodine. The majority of cases conform to the pattern reported by Ikeda et al.,209 but in one case, previously reported by Duran et al.,206 no MCAD protein was detected in the liver although Northern blots revealed normal amounts of mRNA of the right size. Cloning and sequencing of DNA fragments from the MCAD genes of the patient suggested that one allele was normal at the N-terminus and the 5' end of the mRNA. The other, however, revealed 2 different mutations affecting the transit peptide, an insertion and a deletion. These could be convincingly explained by aberrant splicing of precursor mRNA containing a mutation at an exon-intron boundary. The consequence of these mutations was a protein which was not efficiently transported into the mitochondria (see Section X) and was therefore degraded in the cytosol. It is not entirely clear, however, why such a mutation in one allele should also result in degradation of the other, if indeed the second is normal at the 5' end. An alternative to immunological methods that is more immediately relevant to the clinical condition is the direct measurement of enzyme activity. The main problem here is shortage of material from biopsy or necropsy and especially from antenatal sampling procedures. Culturing cells from skin biopsies, aminocentesis, or chorionic villi imposes a considerable delay (which could be critical in prenatal diagnosis). With the usual assay procedures there are also serious problems of nonspecific interference in crude extracts, which give very high blank rates. Samples therefore have to be worked up first (e.g., by ammonium sulfate fractionation)199-200 and this inevitably entails a loss in quantitative accuracy. In order to alleviate some of these problems, various modified assay procedures have been introduced. One approach relies on the ability of the dehydrogenases to exchange hydrogen, and therefore also tritium, from the a-carbon with solvent protons.211 Another popular procedure has been the use of purified ETF as terminal acceptor,212 since ETF unlike DCPIP, etc. is not reduced by the various other potential reductants in a crude extract. Recent developments in DNA methodology have, however, raised the possibility of obtaining clearcut diagnostic information on much smaller initial samples of patient material. The prerequisite for such studies is a specific probe for the gene in question. This has been provided by the cloning activities of two groups in the United States, those of Arnold Strauss and Kay Tanaka, who independently reported the gene sequence of medium-chain acyl-CoA dehydrogenase in 1987.118>119 Strauss has made probes available to other groups, including our own, making possible an international collaboration on the molecular biology of the enzyme deficiency in man. At Sheffield we have investigated the usefulness of restriction fragment length polymorphism (RFLP) analysis. Studies of the normal human population revealed useful levels of polymorphism with several restriction enzymes213"215 (Figure 23) when the labeled MCAD cDNA was used to probe Southern blots of digests of human genomic DNA. This means in each case that somewhere in the vicinity of the MCAD gene, but not necessarily within it, is a cut site for the restriction enzyme which is present in some individuals but not in others. When a genetic disease is associated with a single mutation (i.e., the same codon is invariably the site of mutation, as in sickle-cell anemia) such methods can provide a direct and unequivocal diagnosis in an individual for whom no other information is available. Even

FIGURE 23. An example of an RFLP obtained with labeled MCAD cDNA as the probe. In this case the human DNA samples were digested with Bam HI. Bands at 2.7, 7, and approximately 12 kb are seen in all individuals examined. The 5.5-kb band is from allele 1, track 1 is from a homozygote. The 10-kb band is from allele 2, tracks 3 and 5 are from homozygotes for this allele. Tracks 2 and 4 are from heterozygotes and show both the 5.5-kb and 10-kb bands.

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when this is not the case, however, RFLP analysis can be of considerable value in a family study. Given a sufficient panel of restriction enzymes that show polymorphisms, it should be possible to establish for a newborn child or a fetus which of the 4 parental copies of the gene make up the set of two in that child. This is true for normal individuals, but it can be equally well applied to cases with the genetic defect. Since MCAD deficiency is an autosomal recessive condition, both gene copies must be defective in a diagnosed case. Thus, if one child is identified within a family as being affected, all subsequent births can be screened to establish whether they are (1) also affected, (2) carriers, or (3) free of the defective gene(s). In collaboration with the laboratories of Gregersen and Kolvraa in Denmark, we have applied this approach to families in which there is a known case of MCAD deficiency. Results have shown unexpectedly that one haplotype predominates in affected cases.215 This haplotype exhibits allele 1 with Pst I; allele 1 with Sst I; and allele 2 with Taq I. The mapping of the human genomic DNA for the region encoding human medium-chain acyl-CoA dehydrogenase by Strauss and Zhiang233 has made it possible to extend the restriction analysis by using subprobes obtained by restriction enzyme digestion of the cDNA.216 Pst I cuts within exon 3 and gives rise to a 1.7 kb fragment comprising the central and 3' sections of the cDNA; Eco RI cuts within exon 8 and gives a 0.7 kb fragment comprising the central and 5' sections of the cDNA. These two probes thus overlap, and if they are both used to probe Southern blots, any band recognized by both probes must lie between exons 3 and 8; any band detected only by the EcoR I probe must be at the 5' end; and any band hybridizing only with the Pst I probe has to be at the 3' end. The subprobes have been used on restriction digests produced with each of the enzymes that gives rise to an RFLP pattern.216 The polymorphic sites giving rise to the Taq I and Sst I RFLPs are both in the central region of the gene and those giving rise to all the other RFLPs so far identified, including Pst I, are in the 3' region. No RFLPs have, therefore, yet been found in the 5' region (exons 1—3). It has now also emerged that a single base transition is by far the commonest cause of MCAD deficiency.217 It is the so-called "985 mutation" in which the replacement of adenine by guanine at position 985 in the cDNA causes the substitution of Lys-329 by Glu. From the enzymological standpoint this is rather unexpected in that there are in theory so many ways in which one might envisage different mutations leading to defective or inactive enzyme. Two obvious explanations present themselves. One is the "founder effect", implying that all affected cases are ultimately descended from a single affected individual. This might tie in with the fact that the disease appears to be largely confined to North-West Europeans and white North Americans. The second possibility involves a "hot spot", for whatever reason, in the DNA/RNA, so that the same mutation is likely to occur repeatedly. A third possibility is that, like the sickle-cell mutation, the 985-mutation confers some selective advantage. At this stage it is difficult to envisage what such an advantage could be or why it should be confined to one site within the protein/ gene. Over the past year it has emerged that the 985 mutation is not the only cause of MCAD deficiency. Nevertheless it seems that about 85% of defective MCAD alleles carry the 985 mutation. In 410 consecutive births in the Trent Health Region (U.K.) 6 G985 heterozygotes were found, an incidence of 1 in 68 (no homozygotes).218 This analysis was carried out on DNA from stored Guthrie spots, i.e., on a single drop of dried blood for each infant. Assuming no selection in utero, this frequency translates to an affected homozygote frequency of approximately 1 in 18,500(68 x 68 x 4). The contribution of other mutant alleles raises the overall incidence of the enzyme deficiency to about 1 in 13,400, making it one of the commonest genetically determined metabolic defects. Implicit in these various findings is a correlation between the G985 mutation and the '*!, 1, 2" haplotype. This correlation is absolute so far in MCAD patients, although it has

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not yet been tested in asymptomatic G985 heterozygotes. Such a correlation may be a piece of evidence for the relatedness of individuals carrying the G985 mutation. Alternatively, one or more of the polymorphisms may not be phenotypically neutral. Changes at these sites could perhaps interact with and modify the consequences of the G985 mutation. In addition to the possibility of rapidly characterizing mutant MCAD genes at the DNA sequence level, there is now also the possibility of attempting to interpret mutations in terms of the three-dimensional structure of the protein. It remains to be seen whether the majority of functional lesions can be explained without going to the lengths of expressing, purifying, and characterizing the mutant protein, although a system for expression in E. coli has been described. In the case of the G985 mutation, however, the likely effect of the amino acid change on enzyme function is not immediately obvious. E. SHORT-CHAIN ACYL-CoA DEHYDROGENASE DEFICIENCY In contrast to the medium-chain deficiency, defects affecting the short-chain acyl-CoA dehydrogenase are rare. In some of the cases that have been described219"221 it appears that the error may be not so much a fault in the structural gene but rather a failure in tissuespecific expression. These cases display predominantly muscle-specific symptoms and yet there is no other current evidence to suggest tissue-specific isoenzymes. Of the three clearcut cases of short-chain acyl-CoA dehydrogenase deficiency in man reported by Amendt et al.222 and Coates et aL, 221 one was fatal in the neonatal period. Enzyme assays with butyryl-CoA were carried out on fibroblast extracts after precipitating MCAD with specific antibody: activity was no more than 11% of controls in any of the cases. Two cases fell into a pattern primarily characterized by acidosis and ethylmalonicaciduria, distinctly different from the pattern already mentioned in which the most obvious symptoms affect skeletal muscle with progressive weakness and carnitine deficiency. Naito et al.223 describe tests on fibroblasts in three cases, including the last of the three mentionedabove, which show that the mRNA is of normal size, and that SCAD protein of the right size is present in normal amounts. All of these cases therefore appear to be due to point mutations. An interesting model for human SCAD deficiency has been found in the inbred strains of BALB/c mice reared for experimental purposes.224 The mutant subline BALB/cByJ is totally devoid of short-chain activity in liver and skeletal muscle. Western blots, moreover, show no detectable SCAD protein. The urine of these mice showed very high levels of butyrylglycine and ethylmalomc and methylsuccinic acids (the latter two arise by taking butyric acid down the normal metabolic pathway for propionate). The mice nevertheless appeared normal clinically even when challenged with medium-chain triglyceride. It seems that more efficient use of glycine conjugation in the mice may spare the carnitine pool (which is drawn upon for conjugation in man) thus causing fewer overt symptoms. Fasting, however, does result in marked fatty infiltration of tissues, as seen in the human acyl-CoA dehydrogenase deficiencies. F. LONG-CHAIN ACYL-CoA DEHYDROGENASE DEFICIENCY Three patients were described in detail by Hale et al.225 A further eleven have been added to this list and all fourteen have now been reviewed by Hale et al.226 Six of the fourteen had died at the time of the report, five of these before one year of age. Apart from a tendency to hypoglycemia and fasting coma, the detailed clinical descriptions are far from uniform. Long-chain acyl-CoA dehydrogenase activity was measured on fibroblasts in ten cases and also in leukocytes in four cases. The main activities were about eightfold down on controls. Molecular biological investigations have not been reported as yet. G. ISOVALERICACIDEMIA The genetic deficiency of isovaleryl-CoA dehydrogenase has already been touched upon

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TABLE 7 Subunit Molecular Weights of Precursor and Mature Forms of Rat Acyl-CoA Dehydrogenases Enzyme Short-chain acyl-CoA dehydrogenase Medium-chain acyl-CoA dehydrogenase Long-chain acyl-CoA dehydrogenase Isovaleryl CoA dehydrogenase

Precursor

Mature form

Leader

45,000 49,000 48,000 45,000

41,000 45,000 45,000 43,000

4,000 4,000 3,000 2,000

Note: Data from Reference 229. Mr values are based on SDS-PAGE analysis of immunoprecipitates.

in earlier sections. It has been fully reviewed recently by Tanaka.227 A number of mutants producing subunits of altered size have been described, as well as those which appear to be point mutations. One mutant has been shown to involve a frameshift leading to premature termination. Another, producing no cross-reacting material (precursor or mature), seems likely to be due to a frameshift in the 5'-region.

X. BIOSYNTHESIS In eukaryotic organisms the substrates of the acyl-CoA dehydrogenases are major fuels for the ATP-generating apparatus of the respiratory chain, and accordingly the enzymes themselves are found in the mitochondrial matrix. They are, however, encoded, like most other mitochondrial proteins, by nuclear genes. Most imported mitochondrial proteins are synthesized in a larger precursor form carrying a positively charged "leader" sequence which is cleaved off in the course of passage across the mitochondrial membrane.228 The acyl-CoA dehydrogenase are no exception. Several groups have added to our knowledge in this area of acyl-CoA dehydrogenase research, but it is appropriate to make particular mention of the major and sustained contribution of Tanaka's group whose efforts have been exceptional in their breadth. In the study of biosynthesis and processing of these enzymes, antibodies have been essential tools. Tanaka and his colleagues have painstakingly separated all the known members of the acylCoA dehydrogenase family from one another and used them to raise monospecific antibodies.13-1 15 Rat liver polysomal RNA was isolated and translated using the rabbit reticulocyte lysate system supplied with 35S methionine.229 Monospecific antibodies allow immunoprecipitation of the newly synthesized acyl-CoA dehydrogenases, and the analysis of immunoprecipitates on SDS gels, with fluorographic detection of the 35S-labeled bands, reveals that in all cases subunits in the acyl-CoA dehydrogenase precursors are 2-4 kDa larger than in the mature processed enzymes (Table 7). The cell-free translation products were also used to examine import into the mitochondria. After incubation with intact mitochondria at 30°C for varying times up to 1 h, the preparations were centrifuged, and the supernatant and mitochondrial pellet fractions were separately analyzed by immunoprecipitation and SDS-PAGE. Precursors were found in diminishing amounts in the supernatant and also in the pellet. Mature protein was found, in increasing amounts, only in the pellet. Trypsin treatment destroyed the precursor in both fractions, proving that precursor associated with the mitochondrial pellet was external. Gel filtration analysis of these various fractions showed no sign of tetrameric, assembled enzyme in the supernatant. Only precursor monomer and much larger (circa 400 kDa) aggregates were seen. Of the processed protein from the mitochondrial fractions, a small proportion was in a high molecular weight fraction but most ran as tetramer or monomer. The authors surmise that most likely the monomer is imported in an unfolded state and that therefore insertion of the FAD cofactor is likely to occur in the mitochondrial matrix.

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In the case of the medium-chain acyl-CoA dehydrogenase a similar study on the formation and processing of the precursor has been carried out by Inagaki et al.230 Strauss and colleagues, who cloned and sequenced the human MCAD gene, have used their probes mainly in order to study tissue-specific expression and developmental regulation. Northern dot-blot analysis231 with 32P-labeled cDNA shows MCAD mRNA to be present in lung, heart, intestine, skeletal muscle, liver, and kidney (all the tissues tested), but it is at considerably higher levels in the heart and kidney, in keeping with the importance of fat oxidation in those tissues. In the developmental studies MCAD mRNA was detected at all stages in the heart and liver with a peak in both tissues around birth. In the heart there is a second peak around the time of weaning (day 14), coinciding with a minimum in the liver.

XI. MUTANTS Over the past 4 years a situation has emerged in which examples of most members of the mammalian acyl-CoA dehydrogenase family have been cloned and sequenced, and detailed three-dimensional information is now available for one member of the family. This makes it possible to attempt to interpret the properties of mutant acyl-CoA dehydrogenases or at least of those where the mutation falls within the mature, processed protein. Mutants may be either those that occur naturally or those that have been created by design. As we have seen in Section IX, naturally occurring mutants are being found in increasing numbers, and the advent of the polymerase chain reaction will make the pinpointing of genetic lesions more rapid than hitherto. One can envisage two distinct approaches to a given mutation. The first is to turn to modeling and molecular graphics, making use of the existing high-resolution structure of the medium-chain acyl-CoA dehydrogenase as a framework for explaining the effects of the mutation. The second is to examine the mutant enzyme directly. This imposes a requirement for an expression system, since the amounts of enzyme available from cultured fibroblasts or small amounts of tissue are not sufficient for much useful enzymology. Fortunately a system for expression of the medium-chain acyl-CoA dehydrogenase in E. coli has been reported by Bross et al.232 This is equally the key to protein engineering of acyl-CoA dehydrogenases and a start has been made. The first sitedirected mutation has been a Glu -> Gin change at the position identified (Section VII) as the possible catalytic base.232 This mutation resulted, as predicted, in inactive enzyme. Curiously, this mutant enzyme is green (see Section V.C)! Whether this reflects tighterthan-normal binding of CoA-persulfide or a protein-flavin interaction is not yet clear. A number of deletions at the N-terminus have also been constructed, of which several result either in weakened substrate binding or total inactivity. 12 Clearly this is only a small beginning, but it is enough to show that we are on the threshold of an explosive growth in our understanding of structure-function relationships in this enzyme family and also of a much deeper understanding of the nature of acyl-CoA dehydrogenase mutations in man.

ACKNOWLEDGMENTS I am deeply indebted to Janet Jones, Gary Williamson, Lee Shaw, Paul Brown, Trish Ellison, and Alex Blakemore who have over the years made possible most of my occasional contributions to this area of research; to the Science and Engineering Research Council, the Wellcome Trust, and the Foundation for the Study of Infant Death who have supported the work; to Vince Massey and Steve Mayhew who first introduced me to the more colorful areas of biochemistry; to various collaborators and especially Drs. Colin Thorpe, Yasuzo Nishina, and Kiyoshi Shiga, Bob Bray, and Graham George, Michael Bennett, Diana Curtis, Arnold Strauss, Niels Gregersen, and Steen Kolvraa; and to Mrs. Janette Butler for patiently putting up with all the revisions in the typing.

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29. Elsden, S. R. and Lewis, D., Production of fatty acids by a Gram-negative coccus, Biochem. J., 55, 183, 1953. 30. Williamson, G. and Engel, P. C., Butyryl CoA dehydrogenase from Megasphaera elsdenii. Specificity of the catalytic reaction, Biochem. J . , 218, 521, 1988. 31. Sedlmaier, H., Buhler, M., Feicht, R., Bader, J., and Simon, H., Some observations on an acroylCoA reductase from Ctostridium kluyveri and an NADH-dependent fumarate reductase from Enterobacter agghmerans, in Flavins andFlavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 467. 32. Tanaka, K., Budd, M. A., Efron, M. L., and Isselbacher, K. J., Isovalericacidemia: a new genetic defect of leucine metabolism, Proc. Natl. Acad. Sci. U.S.A., 56, 236, 1966. 33. Tanaka, K., Miller, E. M., and Isselbacher, K. J., Hypoglycin A: a specific inhibitor of isovaleryl CoA dehydrogenase, Proc. Natl. Acad. Sci. U.S.A., 68, 20, 1971. 34. Tanaka, K., Isselbacher, K. J., and Shih, V., Isovaleric and a-methylbutyricacidemias induced by Hypoglycin A: mechanism of Jamaican Vomiting Sickness, Science, 175, 69, 1972. 35. Engel, P. C., Possibility of inborn defect in isovalericacidemia involving altered enzyme specificity rather than total inactivity, Nature, 248, 140, 1974. 36. Sherratt, H. S. A., Holland, P. C., Osmundsen, H., and Senior, A. E., On the mechanism of inhibition of fatty acid oxidation by hypoglycin and by pent-4-enoic acid, in Hypoglycin, Kean, E. A., Ed., Academic Press, New York, 1975, 127. 37. Noda, C., Rhead, W. J., and Tanaka, K., Isovaleryl CoA dehydrogenase: demonstration in rat liver mitochondria by ion exchange chromatography and isoelectric focusing, Proc. Natl. Acad. Sci. U.S.A., 77, 2646, 1980. 38. Krans, J. P., Matsubara, Y., Barton, D., Yang-Feng, T. L., Glassberg, R., Ito, M., Ikeda, Y., Mole, J., Francke, IL, and Tanaka, K., Isolation of cDNA clones coding for rat isovaleryl-CoA dehydrogenase and assignment of the gene to human chromosome 15, Genomics, 1, 264, 1987. 39. Matsubara, Y., Indo, Y., Naito, E., Ozasa, H., Glassberg, R., Vockley, J., Ikeda, Y., Kraus, J., and Tanaka, K., Molecular cloning and nucleotide sequence of cDNAs encoding the precursors of rat long chain acyl-coenzyme A, short-chain acyl-coenzyme A and isovaleryl-coenzyme A dehydrogenases: sequence homology of four enzymes of the acyl-CoA dehydrogenase family, J. Biol Chem., 264, 16321, 1989. 40. Finocchiaro, G., Ito, M., and Tanaka, K., Purification and properties of short-chain acyl-CoA, medium chain acyl-CoA and isovaleryl-CoA dehydrogenases from human liver, /. Biol. Chem., 262, 7982, 1987. 41. Matsubara, Y., Ito, M., Glassberg, R., Ikeda, Y., and Tanaka, K., cDNA cloning and nucleotide sequence of human isovaleryl-CoA dehydrogenase (IVS) and its expression in isovalericacidemia fibroblast cell lines, Pediatr. Res., 23, 332A, 1988. 42. Beinert, H., von Korff, R. W., Green, D. E., Buysske, D. A., Handschuhmacher, R. E., Higgins, H., and Strong, F. M., A method for the purification of coenzyme A from yeast, J. Biol. Chem., 200, 385, 1953. 43. Engel, P. C., Williamson, G., and Shaw, L., Butyryl-CoA dehydrogenase: aspects of acceptor and substrate specificity in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 403. 44. Engel, P. C. and Massey, V., Green butyryl-Coenzyme A dehydrogenase, an enzyme-acyl-Coenzyme A complex, Biochem. J., 125, 889, 1971. 45. Frerman, F. E., Miziorko, H. M., and Beckmann, J. D., Enzyme-activated inhibitors, alternate substrates, and a dead end inhibitor of the general acyl-CoA dehydrogenase, J. Biol. Chem., 255, 1U92, 1980. 46. Thorpe, C., Ciardelli, T. L., Stewart, C. J., and Wieland, T., Interaction of long-chain acyl-CoA analogs with pig kidney general acyl-CoA dehydrogenase, Eur. J. Biochem., 118, 279, 1981. 47. Beinert, H. and Page, E., On the mechanism of dehydrogenation of fatty acyl derivatives of Coenzyme A. V. Oxidation-reductions of the flavoproteins, J. Biol. Chem., 225, 479, 1957. 48. Steyn-Parve, E. P. and Beinert, H., On the mechanism of dehydrogenation of fatty acyl derivatives of Coenzyme A. VI. Isolation and properties of stable enzyme-substrate complexes, J. Biol. Chem., 233, 843, 1958. 49. Mii, S. and Green, D. E., Studies on the fatty acid oxidising system of animal tissues. VII. Reconstruction of fatty acid oxidising system with triphenyltetrazolium as electron acceptor, Biochim. Biophys. Acta, 13, 425, 1954. 50. Hauge, J. G., On the mechanism of dehydrogenation of fatty acyl derivatives of Coenzyme A. IV. Kinetic studies, J. Amer. Chem. Soc.t 78, 5266, 1956. 51. Engel, P. C. and Massey, V., The purification and properties of butyryl-Coenzyme A dehydrogenase from Peptostreptococcus elsdenii, Biochem. J., 125, 879, 1971. 52. Mcllwain, H., The phenazine series. Part VI. Reaction of alkyIphenazoniurn salts; the phenazyls, J. Chem. Soc., 1704, 1937.

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53. Engel, P. C., Butyryl-CoA dehydrogenase from Megasphaera elsdenii, in Methods in Enzymology, Vol. 71, Lowenstein, J. M., Ed., Academic Press, New York, 1981, 359. 54. Ghosh, R. and Quayle, J. R., Phenazine ethosulfate as a preferred electron acceptor to phenazine methosulfate in dye-linked enzyme assays, Anal. Biochem., 99, 112, 1979. 55. Dommes, V. and Kunau, W.-H., A convenient assay for acyl-CoA dehydrogenases, Anal. Biochem., 71, 571, 1976. 56. Lehman, T. C., Hale, D. E., Bhala, A., and Thorpe, C., An acyl-CoA dehydrogenase assay utilising the ferricenium ion, Anal Biochem., 186, 280, 1990. 57 Stanley, C. A., Hale, D. E., Coates, P. M., Hall, C. L., Corkey, B. E., Yang, W., Kelley, R. I., Gonzales, E. L., Williamson, J. R., and Baker, L., Medium-chain acyl-CoA dehydrogenase deficiency in children with non-ketotic hypoglycemia and low carnitine levels, Pediatr. Res., 17, 877, 1983. 58. Coates, P. M., Hale, D. E., Stanley, C. A., Corkey, B. E., and Cortner, J. 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79. van Berkel, W. J. H., van den Berg, W. A. M., and Muller, FM Large-scale preparation and reconstitution of apoflavoproteins with special reference to butyryl-CoA dehydrogenase from Megasphaera elsdenii. Hydrophobic-interaction chromatography, Eur. J. Biochem., 178, 197, 1988. 80. Mayer, E. J. and Thorpe, C. A method for resolution of general acyl-Coenzyme A dehydrogenase apoprotein, Anal. Biochem., 116, 227, 1981. 81. Thorpe, C. and Massey, V., Flavin analogue studies of pig kidney general acyl CoA dehydrogenase, Biochemistry, 22, 2972, 1983. 82. Murfin, W., Mechanism of the Flavin Reduction Step in Acyl-CoA Dehydrogenases, Ph.D. thesis, Washington University, University Microfilms International, Ann Arbor, MI, 1974. 83. Eweg, J. K., Muller, F., van Berkel, W. J. H., and Hesper, B., On the enigma of Old Yellow Enzyme's spectral properties, in Flavins and Flavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walter de Gruyter, Berlin, 1984, 183. 84. 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106. Miiller, F., Massey, V., Heizmann, G., Hemmerich, P., Lhoste, J. M., and Gould, D. C., The reduction of flavins by borohydride: 3, 4-dihydroflavin structure, absorption and luminescence, Eur. J. Biochem., 9, 392, 1969. 107. Massey, V., Curti, B., Miiller, F., and Mayhew, S. G., On the reaction of borohydride with D- and Lamino acid oxidases, J. Biol. Chem., 243, 1329, 1968. 108. Jiang, Z.-Y. and Thorpe, C., Acyl-CoA oxidase from Candida tropicalis, Biochemistry, 22, 3752, 1983. 109. Thorpe, C., Matthews, R. G., and Williams, C. H., Acyl-Coenzyme A dehydrogenase from kidney. Purification and properties, Biochemistry, 18, 331, 1979. 110. Shaw, L., A Study on the Short-Chain Acyl-CoA Dehydrogenase From Ox-Liver, Ph.D. thesis, University of Sheffield, 1985. 111. Whitby, L. G., A new method for preparing flavin-adenine dinucleotide, Biochem. J., 54, 437, 1953. 112. Mayhew, S. G. and Massey, V., Purification and characterization of flavodoxin from Peptostreptococcus elsdenii, J. Biol. Chem., 244, 794, 1969. 113. Frerman, F. E., Kim, J.-J. P., Huhta, K., and McKean, M. C., Properties of the general acyl-CoA dehydrogenase from pig liver, J. Biol. Chem., 255, 2195, 1980. 114. Ikeda, Y., Dabrowski, C., and Tanaka, K., Separation and properties of five distinct acyl-CoA dehydrogenases from rat liver mitochondria. Identification of a new 2-methyl branched chain acyl-CoA dehydrogenase, /. Biol. Chem., 258, 1066, 1983. 115. Ikeda, Y., Okamura-Ikeda, K., and Tanaka, K., Purification of characterization of short-chain, mediumchain and long-chain acyl-CoA dehydrogenases from rat liver mitochondria. Isolation of the holo- and apoenzymes and conversion of the apoenzyme to the holoenzyme, J. Biol. Chem., 260, 1311, 1985. 116. Hall, C. L. and Kamin, H., The purification and some properties of electron transfer flavoprotein and general fatty acyl Coenzyme A dehydrogenase from pig liver mitochondria, J. Biol. Chem., 250, 3476, 1975. 117. Hall, C. L., Heykenskjold, L., Bartfai, T., Ernster, L., and Kamin, H., Acyl Coenzyme A dehydrogenases and electron-transferring flavoprotein from beef heart mitochondria, Arch. Biochem. Biophys., Ill, 402, 1976. 118. Kelly, D. P., Kim, J.-J., Billadello, J. J., Hainline, B. E., Chu, T. W., and Strauss, A. W., Nucleotide sequence of medium-chain acyl-CoA dehydrogenase mRNA and its expression in enzyme-deficient human tissue, Proc. Natl. Acad. Sci. U.S.A., 84, 4068, 1987. 119. Matsubara, Y., Kraus, J. P., Ozasa, H., Glassberg, R., Finocchiaro, G., Ikeda, Y., Mole, J., Rosenberg, L. E., and Tanaka, K., Molecular cloning and nucleotide sequence of cDNA encoding the entire precursor of rat liver medium-chain acyl Coenzyme A dehydrogenase, J. Biol. Chem., 262, 10104, 1987. 120. Kim, J.-J. P., Vollmer, S. H., and Frerman, F. E., Crystallisation and preliminary X-ray data for the general acyl-CoA dehydrogenase, /. Biol. Chem., 259, 3318, 1984. 121. Kim, J.-J. P. and Wu, J., Structure of the medium-chain acyl-CoA dehydrogenase from pig liver mitochondria at 3 A resolution, Proc. Natl. Acad. Sci. U.S.A., 85, 6677, 1988. 122. Kim, J.-J. P., Flavins andFlavoproteins, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1990. 123. Naito, E., Ozasa, H., Ikeda, Y., and Tanaka, K., Molecular cloning and nucleotide sequence of complementary DNAs encoding human short-chain acyl coenzyme A dehydrogenase and the study of the molecular basis of human short-chain acyl-coenzyme A dehydrogenase deficiency, J. Clin. Invest., 83, 1605, 1613. 124. Engel, P. C., Thorpe, C., and Williams, C. H., Jr., unpublished experiments. 125. Ikamura-Ikeda, K., Ikeda, Y., and Tanaka, K., An essential cysteine residue located in the vicinity of the FAD-binding site in short-chain, medium-chain, and long-chain acyl-CoA dehydrogenases from rat liver mitochondria, J. Biol. Chem., 260, 1338, 1985. 126. Mizzer, J. P. and Thorpe, C., An essential methionine in pig kidney general acyl-CoA dehydrogenase, Biochemistry, 19, 5500, 1980. 127. Jiang, Z.-Y. and Thorpe, C., Modification of an arginine residue in pig kidney general acyl-coenzyme A by cyclohexane- 1, 2-dione, Biochem. J., 207, 415, 1982. 128. Chen, S.-S. and Engel, P. C., The equilibrium position of the reaction of bovine liver glutamate dehydrogenase with pyridoxal 5'-phosphate. A demonstration that covalent modification with this reagent completely abolishes catalytic activity, Biochem. J., 147, 351, 1975. 129. Beckmann, J. D. and Frerman, F. E., The effects of pH, ionic strength, and chemical modifications on the reaction of electron transfer flavoprotein with an acyl-CoA dehydrogenase, J. Biol. Chem., 258, 7563, 1983. 130. Frerman, F. E., Mielke, D., and Huhta, K., The functional role of carboxyl residues in an acyl-CoA hydrogenase, /. Biol. Chem., 255, 2199, 1980. 131. Abeles, R. H., Suicide enzyme inactivators, in Enzyme-Activated Irreversible Inhibitors, Seiler, N., Jung, M. J. and Koch-Weser, J., Eds., Elsevier/North Holland, Amsterdam, 1978, 1.

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159. Tanaka, K., O'Shea, J. J., Finocchiaro, G., Ikeda, Y., Aberhart, D. J., and Ghoshal, P. K., Substrate stereochemistry of 2-methyl-branched acyl-CoA dehydrogenase: elimination of one hydrogen each from (pro-2S)-methyl and a-methine of isobutyryl CoA, Biochim. Biophys. Acta, 873, 308, 1986. 160. Manstein, D. J., Pai, E. F., Schopfer, L. M., and Massey, V., Absolute stereochemistry of flavins in enzyme-catalysed reactions, Biochemistry, 25, 6807, 1986. 161. Cornforth, J. W., Biosynthesis of fatty acids and cholesterol considered as chemical processes, J. Lipid Res., 1, 3, 1959. 162. Hamilton, G. A., The proton in biological redox reactions, in Progress in Bio-Organic Chemistry, Kaiser, E. T. and Kezdy, F. J., Eds., John Wiley & Sons, New York, 1971, 1, 83. 163. Powell, P. J., Lau, S.-M., Killian, D., and Thorpe, C., Interaction of acyl Coenzyme A substrates and analogues with pig kidney medium-chain acyl-CoA dehydrogenase, Biochemistry, 26, 3704, 1987. 164. Lau, S.-M., Brantley, R. K., and Thorpe, C., The reductive half-reaction in acyl-CoA dehydrogenase from pig kidney: studies with thiaoctanoyl-CoA and oxaoctanoyl-CoA analogues, Biochemistry, 27, 5089, 1988. 165. Reinsch, J., Rojas, C., and McFarland, J. T., Kinetic methods for the study of the enzyme systems of (3-oxidation, Arch. Biochem. Biophys., 227, 21, 1983. 166. Schopfer, L., Massey, V., Ghisla, S., and Thorpe, C., Oxidation-reduction of general acyl-CoA dehydrogenase by the butyryl-CoA/crotonyl-CoA couple. A new investigation of the rapid reaction kinetics, Biochemistry, 27, 6599, 1988. 167. Raichle, T., Untersuchungen zum Reaktionsmechanismus der Acyl-CoA Dehydrogenasen, Ph.D. thesis, University of Konstanz, 1981. 168. Pohl, B., Raichle, T., and Ghisla, S., Studies on the reaction mechanism of general acyl-CoA dehydrogenase. Determination of selective isotope effects in the dehydrogenation of butyryl CoA, Eur. J. Biochem., 160, 109, 1986. 169. Ghisla, S., Thorpe, C., and Massey, V., Mechanistic studies with general acyl-CoA dehydrogenase and butyryl-CoA dehydrogenase: evidence for the transfer of the fi-hydrogen to the flavin N(5)-position as a hydride, Biochemistry, 23, 3154, 1984. 170. Ghisla, S., Mechanism of ot, p-dehydrogenation of fatty acid CoA derivatives by flavin enzymes, in Flavins andFlavoproteins, Bray, R. C., Engel, P. C., and Mayhew, S. G., Eds., Walterde Gruyter, Berlin, 1984, 385. 171. Ellison, P. A. and Engel, P. C., Does butyryl-CoA dehydrogenase catalyse thiol-thioester exchange?, Biochem. Soc. Trans., 15, 236, 1987. 172. Ellison, P. A. and Engel, P. C., Crotonase activity in preparations of butyryl-CoA dehydrogenase from Megasphaera elsdenii, Biochem. Soc. Trans., 14, 158, 1986. 173. Waters, B. W., Engel, P. C., and Williamson, G., Photo-inactivation of butyryl-CoA dehydrogenase, Biochem. Soc. Trans., 14, 138, 1986. 174. Lau, S.-M., Powell, P., Buttner, H., Ghisla, S., and Thorpe, C., Medium-chain acyl Coenzyme A dehydrogenase from pig kidney has intrinsic enoyl Coenzyme A hydratase activity, Biochemistry, 25, 4184, 1986. 175. Tanaka, K. and Coates, P. M., Eds,, Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Alan R. Liss, New York, 1990. 176. Hill, K. R., The vomiting sickness of Jamaica: a review, W. Ind. Med. J., 1, 243, 1952. 177. Hill, K. R., Hypoglycemia and fatty metamorphosis of the liver in the vomiting sickness of Jamaica, J. Pathol. Bacteriol., 66, 334, 1953. 178. Jelliffe, D. B. and Stuart, K. L., Acute toxic hypoglycemia in the vomiting sickness of Jamaica, Brit. Med. /., 1, 175, 1954. 179. Hassall, C. H. and Reyle, K., Hypoglycin A and B, two biologically active polypeptides from Blighia sapida, Biochem. J., 60, 334, 1955. 180. Patrick, S. J., Effects of Hypoglycin A on the metabolism of glucose by isolated tissues, Can. J. Biochem. PhysioL, 40, 1195, 1962. 181. Manchester, J. E. and Manchester, K. L., Separation of Hypoglycin A from leucine and other amino acids on Sephadex G-10, J. Chromatog., 193, 148, 1980. 182. von Holt, C., Methylenecyclopropaneacetic acid, a metabolite of hypoglycin, Biochim. Biophys. Acta, 125, 1, 1966. 183. von Holt, C., von Holt, M., and Bohm, H., Metabolic effects of hypoglycin and methylenecyclopropaneacetic acid, Biochem. Biophys. Acta, 125, 11, 1966. 184. Peters, R. A., Lethal synthesis. Croonian Lecture, Proc. Roy. Soc. Ser. B, 139, 143, 1952. 185. von Holt, C. and von Holt, L., Zur Biochemie des Hypoglycins A, Naturwissenschaften, 45, 546, 1958. 186. Kean, E. A., Selective inhibition of acyl-CoA dehydrogenases by a metabolite of hypoglycin, Biochim. Biophys. Acta, 422, 8, 1976. 187. Sherratt, H. S. A., Hypoglycin and related hypoglycaemic compounds, Brit. Med. Bull, 25, 250, 1969.

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214. Blakemore, A. I. F., Engel, P. C., and Curtis, D., Bam HI and Mspl RFLPs in strong linkage disequilibrium at the medium-chain acyl-Coenzyme A dehydrogenase locus (ACADM chromosome 1), Nucleic Acids. Res., 18, 2838, 1990. 215. Blakemore, A. I. F., Kolvraa, S., Gregersen, N., Engel, P. C., and Curtis, D., Characterisation and localisation of RFLPs of the medium-chain acyl CoA dehydrogenase gene, Human Genet., 86 , 537,1991. 216. Blakemore, A. I. F., Curtis, D., Engel, P. C., Kolvraa, S., and Gregersen, N., Restriction analysis of the human medium-chain acyl-CoA dehydrogenase (ACADM) genomic region, in Flavins and Flavoproteins 1990, Curti, B., Ronchi, S., and Zanetti, G., Eds., Walter de Gruyter, Berlin, 1991, 901. 217. Gregersen, N., Andresen, B. S., Bross, P., Winter, V., Rudiger, N., Engst, S., Ghisla, S., Christensen, E., Kelly, D., Strauss, A. W., Kolvraa, S., Bolund, L., Blakemore, A. I. F., Curtis, D., and Engel, P. C., Characterisation of a disease-causing Lys-329 to Glu mutation in 16 patients with medium-chain acyl-CoA dehydrogenase deficiency, J. Inker. Metab. Dis., 14, 314 . 1991. 218. Blakemore, A. I. F., Singleton, H., Pollitt, R. J., Engel, P. C., Kolvraa, S., Gregersen, N., and Curtis, D., Frequency of the G985 MCAD mutation in the general population, Lancet, 337, 298, 1991. 219. Turnbull, D. M., Bartlett, K., Stevens, D. L., Alberti, K. G. M. M., Gibson, G. J., Johnson, M. A., McCulloch, A. J., and Sherratt, H. S. A., Short-chain acyl-CoA dehydrogenase deficiency associated with a lipid-storage myopathy and secondary carnitine deficiency, N. Engl. J. Med., 311, 1232, 1984. 220. Di Donato, S., Cornelio, F., Gellera, C., Peluchetti, D., Rimola, M., and Taroni, F., Short-chain acyl-CoA dehydrogenase-deficiency myopathy, with secondary carnitine deficiency, Muscle Nerve, 9, 178, 1986. 221. Coates, P. M., Hale, D. E., Finocchiaro, G., Tanaka, K., and Winter, S. C., Genetic deficiency of short-chain acyl-Coenzyme A dehydrogenase in cultured fibroblasts from a patient with muscle carnitine deficiency and severe skeletal muscle weakness, J. Clin. Invest., 81, 171, 1988. 222. Amendt, B. A., Green, C., Sweetman, L., Cloherty, J., Shih, V., Moon, A., Teel, L., and Rhead, W. J., Short-chain acyl-coenzyme A dehydrogenase deficiency. Clinical and biochemical studies in two patients, J. Clin. Invest., 79, 1303, 1987. 223. Naito, E., Ozasa, H., Ikeda, Y., and Tanaka, K., Molecular cloning and nucleotide sequence of cDNAs encoding human short chain acyl CoA dehydrogenase and study of the molecular basis of human short chain acyl-CoA dehydrogenase deficiency, in Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Tanaka, K. and Coates, P. M., Eds., Alan R. Liss, New York, 1990, 652. 224. Wood, P. A., Amendt, B. A., Rhead, W. J., Armstrong, D., Millington, D. S., and Inoue, F., A murine model for short-chain acyl-CoA dehydrogenase deficiency, in Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Tanaka, K. and Coates, P. M., Eds., Alan R. Liss, New York, 1990, 427. 225. Hale, D. E., Batshaw, M. L., Coates, P. M., Frerman, F. E., Goodman, S. I., Singh, L, and Stanley, C. A., Long-chain acyl coenzyme A dehydrogenase deficiency: an inherited cause of nonketotic hypoglycemia, Pediatr. Res., 19, 666, 1985. 226. Hale, D. E., Stanley, C. A., and Coates, P. M., The long-chain acyl-CoA dehydrogenase deficiency, in Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Tanaka, K. and Coates, P. M., Eds., Alan R. Liss, New York, 1990, 303. 227. Tanaka, K., Isovaleric acidemia: personal history, clinical survey, and study of the molecular basis, in Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Tanaka, K. and Coates, P. M., Eds., Alan R. Liss, New York, 1990, 273. 228. Hay, R., Bohni, P., and Gasser, S., How mitochondria import proteins, Biochim. Biophys. Acta, 779, 7192, 1984. 229. Ikeda, Y., Keese, S. M., Fenton, W. A., and Tanaka, K., Biosynthesis of four rat liver mitochondrial acyl-CoA dehydrogenases: in vitro synthesis, import into mitochondria, and processing of their precursors in a cell-free system and in cultured cells, Arch. Biochem. Biophys., 252, 662, 1987. 230 Inagaki, T., Tsukagoshi, N., Ichihara, C., Ohishi, N., Udaka, S., Ghisla, S., and Yagi, K., In vitro synthesis of pig kidney general acyl CoA dehydrogenase, Biochem. Biophys. Res. Commun., 137, 1049, 1986. 231. Kelly, D. P. and Strauss, A. W., The tissue specific and developmental regulation of expression of rat medium-chain acyl-CoA dehydrogenase mRNA, in Fatty Acid Oxidation; Clinical, Biochemical, and Molecular Aspects, Tanaka, K. and Coates, P. M., Eds., Alan R. Liss, New York, 1990, 599. 232. Bross, P., Engst, S., Strauss, A. W., Kelly, D. P., Rasched, L, and Ghisla, S., Characterisation of wild-type and an active site mutant of human medium-chain acyl-CoA dehydrogenase after expression in E. coli, J. Biol. Chem., 256, 7116, 1990. 233. Strauss, A. and Zhiang, Z., personal communication.

Volume W

657

INDEX A AAD, see 3-Aminopyridine adenine dinucleotide AAO, see Amino acid oxidases Accessory enzymes, 510—511, see also specific types Acetate, 538 Acetoacetyl-CoA, 601, 609, 614 Acetohydroxyacid synthase, 532 a-Aceto-a-hydroxybutyrate, 532 Acetoin, 532 Acetolactate, 532 Acetolactate synthase, 364, 531—541 ALSII and, 533—538 flavin cofactor and, 538—539 inhibitors of, 539—541 sources of, 532—533 structural properties of, 532—533 Acetyl-CoA, 135, 391, 394 yV-Acetyl-cysteamine thiolesters, 601 Acetylenic inhibitors, 626—627 /V-Acetyl-phosphopyridoxyMysine, 7 Acid-base chemistry, 145—147, 161—164 Acrylyl-CoA reductase, 600 Active-site disulfides, 180—181 Active site studies of acyl-CoA dehydrogenases, 626—630 of o-amino acid oxidases, 78—83 of L-auiino acid oxidases, 87 of bioluminescence, 494—506 of glutamate synthase, 313 of glycolate oxidase, 376—377, 382—383 of lipoamide dehydrogenase, 145, 146 of luciferase, 494—506 of mercuric ion reductase, 180—185 of nitrate reductase, 324—325 of pyridine nucleotide-disulfide oxidoreductases, 128, 169—171 of thioredoxin reductase, 169—171 Active-site thiols, 180—185, 249 Acyl carrier protein, 599 Acyl-CoA dehydrogenases, 597—646, see also specific types biosynthesis of, 645—646 branched-chain, 600, 632 chemical modification of, 622—630 acetylenic inhibitors and, 626—630 group-specific reagents and, 622—626 inhibitors and, 626—630 electron acceptors and, 602—603 hydrogen removal and, 633—635 long-chain, 644 mechanisms of action of, 630—636 medical aspects of, 636—645 medium-chain, see Medium-chain acyl-CoA dehydrogenase (MCAD)

metabolic function of, 598—599 mutants of, 646 oxidation-reduction properties of, 631 oxygen reactivity and, 603—604 primary structure of, 618—621 protein engineering of, 646 quaternary structure of, 616—618 reactivity of, 630—636 short-chain, 605, 628, 632, 644 side activities of, 635—636 spectral properties of, 604—616 acetoacetyl-CoA and, 614 blue semiquinone and, 614—615 charge-transfer complexes and, 604—607 green semiquinone and, 608—614 oxidized, 616 red semiquinone and, 614—615 reduced forms, 615 semiquinone forms, 604—607 stereochemistry of, 631—632 structure of, 616—622 substrate specificities and, 599—602 tertiary structure of, 621—622 thiols and, 600—602 Acyl-CoA oxidases, 604, 616, 627 Acyl-CoA thiol esters, 300 Acyl moiety, 599 Acyl protein reductase, 510 Acyl protein synthase, 510 Adenosine 3'-phosphate-5'-phosphosulfate reductase, 166 Adenosylmethionine, 395 Adrenodoxin, 415 Adrenodoxin reductase, 415, 565 Affinity labeling, 533 (3-Alanine, 609 Alcohol oxidase, see Methanol oxidase Aldehyde oxidase, 22, 58—61 Aldehydes, 469, 479, see also specific types binding of, 481—482, 506 biosynthesis of, 510 Alkyl alkanethiolsulfonates, 475 Alkylation, 142, 155, 174 Amino acid oxidases (AAO), 616, see also specific types D-Amino acid oxidases (DAO), 70—84, 567, 580, 585, see also specific types active-site studies of, 78—83 chemical modification of, 80 cloning and, 83 DNA sequencing of, 83 gene expression and, 83 kinetics of, 76—78 metabolic function of, 88 optimum pH for, 74, 76 physicochemical properties of, 71 —76

658

Chemistry and Biochemistry of Flavoenzymes

physiological role of, 84 pig kidney, 71—74, 76, 83 purification of, 71, 75 redox properties of, 72 reduction of, 72, 75 Rhodotorula gracilis, 75—76, 79 stereospecificity of, 70, 78 structural properties of, 78—83 L-Amino acid oxidases, 74, 84—88, see also specific types inhibitors of, 86—88 kinetics of, 87—88 metabolic function of, 88 physicochemical properties of, 84—86 physiological role of, 88 purification of, 84 sources of, 84 stereospecificity of, 70 sulfite and, 85 Amino acids, see also specific types bioluminescence and, 471 branched-chain, 532 dicarboxylic, 625—626 in luciferase, 488—494 in pyridoxine-5'-P oxidase, 13 2-Amino-4-hydroxy-6-formylpteridine, 26, 58 3-Ammopyridine adenine dinucleotide (AAD), 55, 143

Ammonia assimilation, 310 Ammonium sulfate, 6 Anchor peptides, 238—239, 266—270, see also specific types Anionic species, 452—453, see also specific types Anisotropy, 12, 27 Anoxygenic phototropic bacteria, 342 Anthranylate synthase, 313 Antibodies, 315, 325, 645, see also specific types Antimalarial drugs, 164, see also specific types Antineoplastic drugs, 164, see also specific types AOX-gene, 109 Apoflavodoxins, 401, 402, 404, see also specific types Apoflavoproteins, 558, see also specific types Apo-glutamate synthase, 312 Apo-riboflavin-binding protein, 585 APS kinase, 334 APS reductase, 333—352 from anoxygenic photoropic bacteria, 342 assays for, 338 assimilatory sulfate reduction and, 334—336 chemical mechanisms of, 349—352 from Desulfovibrio spp., 346—349 dissimilatory sulfate reduction and, 336—337 flavin-sulfite adduct and, 337 mechanisms of, 349—352 metabolic functions of, 338 physicochemical properties of, 343 spectral properties of, 342—346 spectroscopy of, 342—346 structural properties of, 342—346 from sulfate-reducing bacteria, 339—341

from Thiobacillus spp., 341—342 Arginine, 13, 625 Arginyl residues, 14 Arrhenius plots, 74 Arsenite, 45, 46, 139 Ascitic cells, 18 Asparagunic acid, 191 Asparagusate dehydrogenase, 191 —192 Aspartate aminotransferase, 16 Assays, see also specific types of APS reductase, 338 enzyme linked immunosorbent (ELISA), 476 of fumarate reductase, 239—241 of luciferase, 469 of nitrate reductase, 43 of pyridoxine-5'-P oxidase, 3 of succinate dehyddrogenase, 239—241 Assimilatory nitrate reductase, see Nitrate reductase Assimilatory sulfate reduction, 334—336 Association constants, 402, 432 Association energies, 428 Autoinduction, 471, 475, 515—520 Autooxidation, 481 Auxiliary thiols, 181 —185, see also specific types Azide complex of methanol oxidase, 100—102

B Bacterial bioluminescence, see Bioluminescence Bacterial luciferase, see Luciferase Baeyer-Villiger reaction, 486 Bathchromatic shift, 12 BCD, see Butyryl-CoA dehydrogenase BCNU, see l,3-Bis(2-chloroethyl)-l-nitrosourea Benzene dioxygenase, 544, 546, 548, 551, 553, 554 Benzoate, 71, 74, 76 Benzoate dioxygenase, 544, 548, 553 Benzoate dioxygenase reductase, 553 Benzoate oxygenase, 546 Biological role of phthalate dioxygenase reductase, 544—545 Bioluminescence, 467—523, see also Luciferase active-site studies of, 494—506 applications of, 476—478 chemical mechanisms of, 485 in vitro, 469—470 in vivo, 510—515 luciferase structural properties and, see Luciferase, structural properties of lux gene regulation and, 515—521 nonfluorescent flavoprotein and, 513—515 yellow fluorescent protein and, 511—512 Biopterin cofactors, 22 Biotechnology, 468 l,3-Bw(2-chloroethyl)-l-nitrosourea (BCNU), 164 ^w-'y-glutamylcystine reductase, 191 fi/s-phosphopyridoxal, 11 Blue semiquinone, 614—615 Borohydride, 72, 75

Volume HI Branched-chain acyl-CoA dehydrogenases, 600, 632, see also specific types Branched-chain ammo acids, 532, see also specific types Bromopyruvate, 533 jV-Bromosuccinimide, 10 2,3-Butadione, 14 3-Butynoyl CoA, 627 Butyryl CoA/crotonyl-CoA, 631 Butyryl-CoA dehydrogenase (BCD), 580, 600, 608, 609, 624, 626

c Calorimetry, 404 Carbodiimides, 625, see also specific types Carbohydrates, 85, see also specific types Carbon-NMR, 106, 558, 574—581, 588 Carboxanilides, 278—279, see also specific types Carboxins, 241, 278, see also specific types 8-Carboxylic acid riboflavin, 404 Catalytic properties of fumarate reductase, 239—242 of glutamate synthase, 315 of glutathione reductase, 161 —164 of glycolate oxidase, 378—380 of lipoamide dehydrogenase, 145—147 of methanol oxidase, 107—109, 115 of succinate dehyddrogenase, 239—242 of thioredoxin reductase, 175—178 CD, see Circular dichroism Cell-free translation, 645 Chagas' disease, 189, 190 Charge relay system, 226 Charge-transfer complexes, 552, 604—607 Chemical composition of flavodoxins, 395—396 Chemical mechanisms of APS reductase, 349—352 of bioluminescence, 485 of glutamate synthase, 314—316 Chemical modification of acyl-CoA dehydrogenases, 622—630 group-specific reagents and, 622—626 inhibitors and, 626—630 of D-amino acid oxidases, 80 bioluminescence and, 487, 494, 495, 503, 504 of glutathione reductase, 164—165 of methanol oxidase, 104—107 of nitrate reductase, 324 of pyridoxine-5'-P oxidase, 12—14 of thiols, 503 Chemical properties, see Physicochemical properties l,3-£/'s(2-Chloroethyl)-l-nitrosourea (BCNU), 164 /?-Chloromercuribenzoate, 625 4-Chlorophenylacetate dioxygenase, 551 4-Chlorophenylacetate 3,4-dioxygenase, 545, 546, 553 Chorismate synthase, 538 Chromatography, 52, 395, 639, see also specific types Chromophores, 113

659

Circular dichroism (CD), 52, 138, 249, 250, 254, 258, 325 flavodoxins and, 403 magnetic, 249, 250, 254, 257, 258, 260 Cloning, 83, 468, 471, 478, 488, 646 Clostridial flavodoxins, 427—460, see also specific types as anionic species, 452—453 conformation change in, 450—452 crystals in, 428 electrostatic interactions and, 453—456, 460 FMN-binding site and, 437—447 proteins and, 437—440 residue conformation and, 440—442 isoalloxazine binding site and, 458—460 isoalloxazine ring and, 446—447 oxidation-reduction properties of, 428 oxidized-semiquinone equilibrium and, 448—452 redox properties of, 430, 453 ring bending and, 456—457 semiquinone-reduced equilibrium and, 452—457 structure of, 428, 433—435 structure-function studies of, 430—433 three-dimensional structure of, 428 X-ray analysis of, 433—437 4-CI-phenylacetate dioxygenase, 550 p-CMB, see p-Mercuribenzoate Coenzyme A, 600, see also Acyl-CoA dehydrogenases persulfide of, 610, 612—614, 646 purification of, 60] Coenzymes, see also specific types for D-amino acid oxidases, 82 for flavodoxins, 395 for glycolate oxidase, 369 for pyridoxine-5'-P oxidase, 8—9 Colorimetry, 3 Complex II, see Succinate-ubiquinone oxidoreductase Conformation change, 450—452 Cooperativity of lipoamide dehydrogenase, 144 Copper, 142 Coupling constants, 572, 578, 579 Covalent inactivators, 313 Cross-linking, 618 Crotonyl-CoA, 631 Crystals in Clostridial flavodoxins, 428, 434 in flavodoxins, 397, 586 in glutathione reductase, 214 in glycolate oxidase, 369 in lipoamide dehydrogenase, 143 in medium-chain acyl-CoA dehydrogenase, 300 in methanol oxidase, 99—100 in thioredoxin reductase, 178 Cyanide, 327, 613 6-Cyanopurine, 59 Cyclohexane-l,2-dione, 625 Cysteine, 107, 313, 334, 335, 624—625 Cysteine residues, 313 Cysteine sulfenic acid, 193

660

Chemistry and Biochemistry of Flavoenzymes

Cysteinyl residues, 265 Cystine reductase, 191 Cytochrome b, 230, 236, 266, 269 Cytochrome c, 38, 61, 338, 350, 410—411, 415 acyl-CoA dehydrogenases and, 603 nuclear magnetic resonance and, 585 Cytochrome c peroxidase, 39, 410, 415 Cytochrome P, 415 Cytochrome P reductase, 415 Cytochrome P-450 reductase, 567 Cytochromes, 412, 414, see also specific types

D DAO, see D-Amino acid oxidases DCPIP, see Dichlorophenolindophenol 5-Deaza-5-carba-8-demethyl-8-hydroxy-flavins, 360—362 5-Deaza-5-carba-7,8-didernethyl-8-hydroxy-flavins, 359—360 5-Deaza-5-carba-FAD, 538 5-Deaza-5-carba-flavins, 359 5-Deaza-flavins, 398, 406 1-Deaza-FMN, 406 1-Deaza-FMN-flavodoxin, 453 n-Decanal, 469, 471, 481, 482, 498, 503—505, 518 Decarboxylase, 136 8-Demethyl-8-chloro-FAD, 538 8-Demethyl-8-hydroxy-5-deaza-5-carba-FAD, 538 3'-Dephospho-CoA, 601 Developmental regulation, 646 Dexaflavin photochemical reduction, 313 Dialysis, 540 Diaphorase, 142, 403 l-Diazo-2-oxoundecane, 505 Dicarboxylic amino acids, 625—626 Dichlorophenolindophenol (DCPIP), 142, 368, 603 2,6-Dichlorophenolindophenol, 165, 239 Dichroism circular, 52, 138, 249, 250, 254, 258, 325 magnetic circular, 249, 250, 254, 257, 258, 260 2,4-Dienoyl-CoA, 628 Diethylamioethyl cellulose, 395 Diethylpyrocarbonate, 13 c/s-Dihydrodiols, 544, see also specific types 1,8-Dihydroflavin, 581 3,4-Dihydroflavin, 616 Dihydrolipoamide, 141, 145 Dihydrolipoic acid, 135 Dihydroxyacetone synthase, 98 Dimer interface, 218—220 Dimethylbenzene ring of FMN, 407—408 6,7-(2,3-Dimethylbutano)-A^(8)-ribityllumazine-5'phosphate, 406 7,8-Dimethy 1-10-formylmethylisoalloxazine, 105 Dimethylhydrazine, 18 6,7-(2,3-Dimethyl)-yV(8)-ribityllumazine-5'phosphate, 406 Dimethylsulfoxide reductase, 51 2,4-Dinitrofluorobenzene, 499

Dioxetanes, 486 Dioxirane, 486 Dioxomolybdenum, 49 Dioxygenases, 544, see also specific types Dispersive X-ray analysis, 42 Dissimilatory sulfate reduction, 334, 336—337 Dissociation constants, 71, 402 Distal (interchange) thiols, 124 Distribution of flavodoxins, 390—395 of glutathione reductase, 154—155 of mercuric ion reductase, 178—179 of oamide dehydrogenase, 137 of thioredoxin reductase, 165—168 Disulfides, see also specific types active site, 180—181 dithiol interchange with, 134, 171, 178 mixed, 134 thiol interchange with, 123, 163, 168 5,5'-Dithiobis-(2-nitrobenzoic acid) (DTNB), 611, 624, 625 Dithiol-disulfide interchange, 134 Dithiols, 143—145, 171, 178, see also specific types Dithionite, 398, 399, 613, 615, 616 Dithionite-reduced dehydrogenase, 54 Dithionite-reduced xanthine dehydrogenase, 54 Dithionite reduction, 313 DNA sequencing of D-amino acid oxidases, 83 n-Dodecanal, 469, 482, 503 Dot-blot analysis, 646 DTNB, see 5,5'-Dithiobis-(2-nitrobenzoic acid)

E EDTA, 33, 72, 401 Effectors, 144—145, 327, see also specific types EHR, see Monoalkylated glutathione reductase Electron acceptors, 142, 338, 368, 391, 602—603 Electron density map, 301 Electron exchange kinetics, 585 Electron exchange rates, 451 Electron nuclear double resonance (ENDOR), 42, 44, 103, 254, 545 Electron paramagnetic resonance (EPR) acyl-CoA dehydrogenases and, 605, 629 aldehyde oxidase and, 59, 60 anchor peptides and, 266, 267 APS reductase and, 342, 344, 346, 347, 350, 352 fumarate reductase and, 255, 257, 260, 261, 263, 265 catalytic properties and, 241 intensity of, 254 MCDvs.,249, 250 redox properties and, 253 intensity of, 254 methanol oxidase and, 103 nitrate reductase and, 325—328 phthalate dioxygenase reductase and, 545, 550 succinate dehydrogenase and, 255, 257, 260, 261, 263, 265 catalytic properties and, 241

Volume III intensity of, 254 MCD vs., 249, 250 redox properties and, 253 thioredoxin reductase and, 174 Very Rapid, 51, 53 xanthine dehydrogenase and, 52, 53 xanthine oxidase and, 26, 28, 32, 41, 46, 47 liquid-helium, 29 O-labeled enzymes and, 49, 50 Very Rapid, 51 Electron self-exchange reaction, 585 Electron spin echo envelope modulation (ESEEM), 254 Electron transfer flavodoxins and, see under Flavodoxins glutamate synthase and, 315 intramolecular, 34 kinetics in, 407—410 lipoamide dehydrogenase and, 147—151 at low potential, 428 Marcus theory of, 399, 410 mercuric ion reductase and, 186—189 between nascent thiols, 143—145 phthalate dioxygenase reductase and, 544—548, 551—552 protein-protein, 415—417 pure, 250, 358 in pyridine nucleotide-disulfide oxidoreductases, 123 pyridoxine-5'-P oxidase and, 16 rate constants for, 410—411 steric effects on rates of, 411—415 thioredoxin reductase and, 172 within xanthine dehydrogenase, 33 xanthine oxidase and, 28—37 Electron transfer protein (ETP), 275, 300, 305, 599, 602, 603 acyl-CoA dehydrogenases and, 639, 641 binding of, 620 as terminal acceptor, 641 Electron transfer protein-CoQ reductase, 599, 602 Electron transfer protein-dehydrogenase, 599, 602 Electrophoresis, 4 Electrostatic interactions, 453—456, 460 Electrostatic modulation of redox kinetics, 408—410 ELISA, see Enzyme linked immunosorbent assays Emission anisotropy, 12 ENDOR, see Electron nuclear double resonance Energies of association, 428 Energy transfer, 113 Enoyl-CoA, 636 Enoyl-CoA hydratase, 636 Enoyl-CoA reductase, 599, 600, 604 Enterobacteria, 334 Enzyme linked immunosorbent assays (ELISA), 475 Enzymes, see also specific types accessory, 510—511 FAD-dependent, 565—569 lipid-activated, 540 Old Yellow, 565, 573, 579, 605

661

Enzymology of luciferase, 478—487 intermediates in, 483—484 postulated reaction mechanisms and, 486—487 reaction scheme of, 479—483 stoichiometry of, 478—479 EPR, see Electron paramagnetic resonance Equilibrium dialysis, 540 ESEEM, see Electron spin echo envelope modulation ETF, see Electron transfer protein Etheno-CoA, 601 AT-Ethylmaleimide, 482, 500, 501, 503, 624 Ethylmalonic acid, 638 EXAFS, see X-ray absorption fine structure Exchange kinetics, 585 Exciplex formation, 113 Excitation spectrum of methanol oxidase, 114 Extinction coefficients, 616

F FAD, see Flavin adenine dinucleotide Fatty acid reductase complex, 510 Fatty acids, 638, see also specific types Ferredoxin-dependent glutamate synthase, 311 Ferredoxin-NADP reductase, 415, 417, 567 Ferredoxin reductase, 548 Ferredoxins, 26, 415, 428, 544, see also specific types Ferredoxin-thioredoxin reductase, 166 Ferricenium, 603 Ferricyanide, 142, 165, 338, 350 Ferricyanide reductase, 239 Flash photolysis, 33, 35, 407 Flavin in apolar environment, 137 fluorescence of, 12, 153 fumarate reductase and, 243—246 incorporation of, 41 methanol oxidase and, 113 one-electron reduction of, 32 protein interactions with, 400—406 reduction of, 16, 77 reoxidation of, 78 site of, 37—41 succinate dehydrogenase and, 243—246 sulfite and, 337 thioredoxin reductase and, 168 xanthine oxidase and, 22, 24, 30, 33, 37—41 Flavin adenine dinucleotide (FAD), 8 aldehyde oxidase and, 58, 61 D-amino acid oxidases and, 71 binding of, 220—222, 271, 304—306 excess, 54 FADH and, 35, 53, 55, 56 flavodoxins and, 404 fumarate reductase and, 245 glutathione reductase and, 220—222 lipoamide dehydrogenase and, 143—145 medium-chain acyl-CoA dehydrogenase and, 300, 304—306

662

Chemistry and Biochemistry of Flavoenzymes

methanol oxidase and, 104—107 modified, 104—107 nascent dithiol and, 143—145 nitrate reductase and, 320 redox properties of, 130, 135 site of, 48 stoichiometry of, 107 succinate dehydrogenase and, 245 xanthine dehydrogenases and, 53, 55, 56 xanthine oxidase and, 22, 29, 35, 40, 48 Flavin adenine dinucleotide (FAD)-dependent enzymes, 565—569, see also specific types Flavin-binding proteins, 9, see also specific types Flavin cofactor, 538—539 Flavin half-potentials, 32 Flavin mononucleotide (FMN), 16 binding of, 404, 437—447 glycolate oxidase and, 374—376, 380—385 proteins and, 437—440 pyridoxine-5'-P oxidase and, 7—12 residue conformation and, 440—442 1-deazo-, 406 deoxy-, 402 dimethylbenzene ring of, 407—408 in electron transfer, 407—408 8-mercapto-, 405 mobility of, 581 oxidation-reduction properties of, 398 protein interactions with, 437—440, 446 4-thio-, 406 Flavin mononucleotide (FMN) analogues, 7—10, see also specific types Flavin mononucleotide (FMN)-dependent enzymes, 560—565, see also specific types Flavin mononucleotide (FMN)-dependent pyridoxine-5'-P oxidase, 2, 6 Flavin mononucleotide (FMN) phosphate, 437 Flavin reductase, 511 Flavin-sulfite adduct, 337 Flavocytochrome b, 363, 379, 381—385, 585 Flavodoxins, 389—417, see also specific types chemical composition of, 395—396 Clostridial, see Clostridial flavodoxins comparisons of, 457—460 crystals in, 586 cytochromes and, 414 defined, 395 distribution of, 390—395 electron transfer and, 406—417 electrostatic modulation and, 408—410 kinetics and, 406—408 protein-protein complex and, 415—417 rate constants and, 410—411 redox properties and, 408—411 steric effects on, 411—415 flavin-protein interactions and, 400—406 functions of, 390—395 molecular weight of, 395—396 nitrogen-NMR and, 569 nuclear magnetic resonance and, 586 oxidation-reduction properties of, 398—401, 428

oxidized, 575 purification of, 395—396 redox properties of, 392—393, 401, 405, 408—411, 430 reduced, 577 structural properties of, 396—398, 428, 447_448 structure-function studies of, 430—433 three-dimensional structure of, 428 Flavoprotein oxidases, 75, 85 Flavosemiquinone, 103—104 Fluorescence, 9, 10—11, 247 bioluminescence and, 483, 487 Clostridial flavodoxins and, 456 flavin, 12, 153 flavodoxins and, 403, 406 lipoamide dehydrogenase and, 138, 148 mercuric ion reductase and, 185 methanol oxidase and, 109—115 protein, 10 quenching of, 12 time-resolved, 406 Fluoride, 581 Fluorine-NMR, 581—583 8-Fluoroflavins, 582 FMN, see Flavin mononucleotide Formaldehyde, 102—103 Formaldehyde dehydrogenase, 102 FRD, see Fumarate reductase Fructose-l,6-bisphosphatase, 166 Fulfite, 337 Fumarate, 246 Fumarate reductase (FRD), 229—284 active site studies of, 248, 249 assays of, 239—241 assembly of complex of, 270—273 carboxanilides and, 278—279 catalytic properties of, 239—242 flavin moiety of, 243—246 gene expression in, 281—284 gene organization of, 279—281 genetics of, 279—284 iron-sulfur clusters and, 249—266 isolation of, 231—232 membrane-bound types of, 230—232 molecular biology of, 279—284 particulate forms of, 241—242 purification of, 231—238 quinone interaction with, 273—278 redox properties of, 253 soluble forms of, 241—242 structural properties of, 279—281 substrate-binding site and, 246—249 subunit processing and, 270 thenoyltrifluoroacetone and, 241, 278—279 Functional domains of nitrate reductase, 322—323 Functional size of nitrate reductase, 321—322 p-(2-Furyl)-propionyl CoA, 630

G Gas liquid chromotography, 639

Volume III GDH, see L-Glutamate dehydrogenase Gene duplication, 493—494 Gene expression in D-amino acid oxidases, 83 in fumarate reductase, 281—284 in glutamate synthase, 316 in succinate dehydrogenase, 281—284 Gene organization, 279—281 Genetic engineering, 468 Genetic recombination studies, 185 Genetics of fumarate reductase, 279—284 of succinate dehydrogenase, 279—284 of xanthine dehydrogenase, 57—58 Glucocorticoid receptor, 166 Glucolate oxidase in spinach, 363 Glucose oxidase, 565, 573, 580 Glutamate, 626 L-Glutamate, 310 L-GIutamate amidotransferases, 310 Glutamate decarboxylase, 16 L-Glutamate dehydrogenase, 310 Glutamate synthase, 309—316 active-site studies of, 313 catalytic properties of, 315 chemical mechanism of, 314—316 ferredoxin-dependent, 311 inactivation of, 313 inhibitors of, 312 kinetics of, 311—312 optimum pH for, 311 physicochemical properties of, 314—316 sources of, 310 spectral properties of, 313—314 substrate specificities and, 311 Glutaminase, 312 Glutamine, 310 Glutamine analogues, 313 L-Glutamine-dependent reaction, 312 Glutamyl, 633 -y-Glutamyl, 312 -y-Glutamylcysteinylglycine, 154 ^-Glutamylcystine reductase, 191 Glutamyl residue, 627 Glutaredoxin, 155, 166, 167 Glutaryl-CoA dehydrogenase, 626 Glutathione, 134, 161, 166, 189 Glutathione disulflde, 134, 154, 163, 189, 214 Glutathione peroxidase, 154, 166 Glutathione reductase, 130, 153—165, 174, 178, 180, 363, 567, 580 acid-base chemistry of, 161 —164 active center of, 222—226 amino acid sequence homology in, 127—134 catalytic properties of, 161—64 chemical modification of, 164—165 crystals of, 214 description of, 214 dimer interface and, 218—220 distribution of, 154—155 inactivation of, 159—161

663

inhibitors of, 164—165 kinetics of, 156—157, 162—164 lipoamide dehydrogenase compared with, 168 metabolic function of, 154—155 monoalkylated (EHR), 143, 144, 147, 155 oxidation-reduction potentials of, 170 polypeptide structure of, 215—218 presteady-state kinetics of, 156—157 redox properties of, 154 residues in, 126—127 sources of, 153 spectral properties of, 156, 161—162 steady-state kinetics of, 157—159 structural properties of, 178, 213—226 active center and, 222—226 dimer interface and, 218—220 FAD-binding site and, 220—222 materials in study of, 214—215 methods in study of, 214—215 polypeptide, 215—218 thioredoxin reductase compared with, 168 trypanothione reductase compared to, 189, 190 Glutathione transferases, 166 Glyceraldehyde-3-phosphate dehydrogenase, 134, 161 Glycine, 135—137, 168 Glycine cleavage system, 136 Glycine reductase, 136 Glycolate oxidase, 367—385 active-site studies of, 376—377, 382—383 catalytic properties of, 378—380 coenzyme for, 369 comparison to other enzymes, 380—385 flavocytochrome b compared to, 381—385 FMN binding and, 374—376 inhibitors of, 368—369 solvent accessibility and, 373 structural properties of, 369—380 catalytic mechanism and, 378—380 crystal, 369 flexibility of, 373 FMN binding and, 374—376 overall, 370 reliability of model of, 369 secondary, 370—373 solvent accessibility and, 373 substrate pocket and, 376—377 subunit packing and, 374 water and, 373—374 substrate specificities and, 368—369 trimethylamine dehydrogenase compared to, 380 Glyoxylate carboligase, 538 Green semiquinone, 608—614 Group-specific reagents, 622—626 GSH, see Glutathione GSSG, see Glutathione disulfide

H Hammett constants, 607 Hammett plot, 48

664

Chemistry and Biochemistry of Flavoenzymes

HDNO, see Hydroxy-D-nicotine oxidase Heme, 266—270, 320 Hemiacetal, 106 Hepatoma ascitic cells, 18 Hepatomas, 18, see also specific types Herbicide resistance, 533 High-performance liquid chromatography (HPLC), 52 Histidine, 13, 107 Histidinyl residue, 499 Histidyl residues, 14 Holoenzyme, 10 HPLC, see High-performance liquid chromatography Hybridization, 573, 574 Hydride transfer, 552 Hydrocinnamoyl-CoA, 607 Hydrogen bonding, 134, 224—226, 439, 558, 572, 575 Hydrogen exchange, 559 Hydrogen-NMR, 106, 558, 584—588 Hydrogen peroxide, 102—103, 192, 604 Hydrogen removal, 633—635 Hydrolysis, 44, 334 4a,5-Dihydro-4a-hydroperoxyflavin, 469 Hydrophobic chromatography, 395 Hydroquinone, 29, 31, 32, 398 a-Hydroxy acids, 364 3-Hydroxyanthranilate, 2 p-Hydroxybenzoate hydroxylase, 567, 573, 580 yV-p-HydroxybutyryOhomoserine lactone, 515, 516 Hydroxyethyl-thiamin pyrophosphate, 533, 537, 540 6-Hydroxy-FMN, 405 8-Hydroxy-FMN, 405 3-Hydroxykynurenine, 2 Hydroxylamine, 103 6-Hydroxy-L-nicotine oxidase, 565, 573, 580 6-Hydroxy-D-nicotine oxidase, 245 2-Hydroxypyrimidine, 59

i Imidazolinone, 540 Immunochemical methods, 86 Immunological properties of nitrate reductase, 325 Inactivation of acyl-CoA dehydrogenases, 627 of glutamate synthase, 313 of glutathione reductase, 159—161 of nitrate reductase, 322 suicide, 627 Inhibitors, see also specific types of acetolactate synthase, 539—541 acetylenic, 626—627 active-site-directed, irreversible, 626—630 of L-amino acid oxidases, 86—88 of glutamate synthase, 312 of glutathione reductase, 164—165 of glycolate oxidase, 368—369 suicide, 628 Interchange (distal) thiols, 124

Intramolecular electron transfer, 34 lodoacetamide, 503, 625 lodoacetate, 625 Iron, 390 Iron deficiency, 271 Iron-sulfur, 230 fumarate reductase and, 249—266 reduction of, 29 succinate dehydrogenase and, 249—266 xanthine dehydrogenase and, 54 xanthine oxidase and, 30, 32 Isoalloxazine, 404, 538 Isoalloxazine binding site, 458—460 Isoalloxazine ring, 446—447 Isoelectric points, 71, 75 Isoenzymes, 85, see also specific types Isolation, see also specific methods of anchor peptides, 266 of fumarate reductase, 231—232 of phthalate dioxygenase reductase, 544 of succinate dehydrogenase, 231—232 Isoleucine, 532 Isomerization, 636 Isovaleric acidemia, 639, 644—645 Isovaleryl-CoA dehydrogenase, 628 Issoalloxazine, 438

J Jamaican vomiting sickness, 636—638

K ot-Ketoacid dehydrogenase, 136 a-Ketoacids, 135, 136, 532, see also specific types a-Ketobutyrate, 532 Af-(p-Ketocaproyl)homoserine lactone, 515 a-Ketoglutarate, 135 a-Ketovalerate, 537 Kinetic isotope effects (KIE), 158, 163, 194 Kinetics, 537 of aldehyde oxidase, 60—61 of D-amino acid oxidases, 76—78 of L-amino acid oxidases, 87—88 of bioluminescence, 480, 482, 483 of conformation change, 450—452 electron exchange, 585 electron transfer, 407—410 electrostatic modulation of, 408—410 exchange, 585 of glutamate synthase, 311—312 of glutathione reductase, 156—157, 162—164 of lipoamide dehydrogenase, 123, 139—142, 151 — 152 of luciferase, 484 of mercuric ion reductase, 179—180 of nitrate reductase, 326 of phthalate dioxygenase reductase, 544, 552—553 ping-pong, 123, 194, 311 presteady-state, 139—141, 156—157, 172, 179

Volume HI of pyridine nucleotide-disulfide oxidoreductases, 123 of pyridoxine-5'-P oxidase, 14—16 redox, 408—410 of reduction, 451 steady-state, 24, 141, 157—159, 175, 180, 194, 311 of thioredoxin reductase, 172 of thioredoxin reduction, 172 of xamhine dehydrogenase, 53—55 of xanthine oxidase, 24, 28—37

L LAAO, see L-Amino acid oxidases Lactyl-thiamin pyrophosphate, 537 Laser flash photolysis, 407 Leucine, 532 L-Leucine, 74 Leucosafranine, 603 Ligand-luciferase complex, 477 Ligands, 179, see also specific types Light absorption spectrum of methanol oxidase, 106 Lipid-activated enzymes, 540 Lipid-activated oxidation, 538 Lipoamide, 136 Lipoamide dehydrogenase, 135—153, 174, 178, 180, 186, 191, 193, 195 amino acid sequence homology in, 127—134 arsenite and, 139 atypically small, 168 catalytic properties of, 145—147 cooperativity of, 144 distribution of, 137 fluorescence of, 138 glutathione reductase compared with, 168 kinetics of, 123, 139—142, 151 — 152 metabolic function of, 135—137 NAD as effector in reaction of, 144—145 NMR and, 567, 580 oxidation-reduction potentials of, 139, 170 physicochemical properties of, 137—138 presteady-state kinetics of, 139—141 reactivity of, 144 redox properties of, 149 residues in, 126—127 spectral properties of, 138—139, 147—153 Lipoic acid, 191 Liquid-helium EPR, 29 Liquid-nitrogen temperature, 29 Liver dehydrogenase, 57 Liver oxidase, 2 Long-chain acyl-CoA dehydrogenase deficiency, 644

Low resolution study, 300—301 Luciferase, 468, 565, 573, 578, see also Bioluminescence active-site studies of, 494—506 amino acids in, 488—494 assays of, 469 description of, 469

665

enzymology of, 478—487 intermediates in, 483—484 postulated reaction mechanisms and, 486—487 reaction scheme of, 479—483 stoichiometry of, 478—487 genetically fused subunits in, 506 heterodimers of, 506—508 hybrid, 504, 507 kinetics of, 484 mutant forms of, 502—503, 507—510 properties of, 506 structural properties of, 487—510 amino acids and, 488—494 architecture and, 488 gene duplication and, 493—494 sequence conservation and, 492—493 subunits of, 506—508 taxonomic relationships among bacteria producing, 494 temperature sensitivity and, 508—510 Luciferase-ligand complex, 477 Lumazine, 486, 487 Lumazine protein, 512—513 Lumiflavin, 411 Lumiflavin geometry, 214 Luminescence, 111 Lux genes, 471—475, 494, 506, 510, 512, 513, 515—521 Lysine, 13, 14, 42

M Magnetic circular dichroism (MCD), 249, 250, 254, 257, 258, 260 Mai ate dehydrogenase, 166 Malate oxidation, 247 Maleimide, 142 Malelonitrile lyase, 538 Marcus theory, 399, 410 Mass spectrometry, 639 MCAD, see Medium-chain acyl-CoA dehydrogenase MCD, see Magnetic circular dichroism Medium-chain acyl-CoA dehydrogenase (MCAD), 299—306, 363, 646, see also Acyl-CoA dehydrogenases; specific types chemical modification of, 627, 628 deficiency of, 639—644 oxidation-reduction properties of, 631 spectral properties of, 605 stereochemistry of, 632, 635 structural properties of, 301—304 thiols and, 601 Melanomas, 18 Meldolablau, 603 Menaquino] oxidase, 269 Menaquinol oxidation, 276 Menaquinone, 236 6-Mercapto-FMN, 405 8-Mercapto-FMN, 405 /?-Mercuribenzoate, 45 Mercuric ion reductase, 178—189, 363, 567, 580

666

Chemistry and Biochemistry of Flavoenzymes

active-site studies of, 180—185 amino acid sequence homology in, 127—134 distribution of, 178—179 electron transfer and, \26—189 kinetics of, 179—180 metabolic function of, 178—179 presteady-state kinetics of, 179 reactivity of, 186—189 redox properties of, 181, f!85 reduction of, 179 residues in, 126—127 steady-state kinetics of, 180 Metabolism acyl-CoA dehydrogenases and, 598—599 D-amino acid oxidases and, 88 L-amino acid oxidases and, 88 APS reductase and, 338 of fatty acids, 638 glutathione reductase and, 154—155 inborn errors of, 638—639 lipoamide dehydrogenase and, 135—137 mercuric ion reductase and, ]78—179 thioredoxin reductase and, 165—168 Methanol oxidase, 95—115 apoprotein of, 104 azide complex of, 100—102 catalytic properties of, 107—109, 115 chemical modification of, 104—107 crystals of, 99—100 flavosemiquinone of, 103—104 fluorescence of, 109—115 formaldehyde interaction with, 102—103 hydrogen peroxide interaction with, 102—103 luminescence of, 111 molecular biology of, 109 optimum pH for, 108 purification of, 98—99 sources of, 98—99 spectral properties of, 106 synthesis of, 96 Methanol-regulated genes, 109 Methanol-utilizing yeast, 96 Methionine, 625 L-Methionine, 74 Methionine sulfoxide reductase, 166 Methionine synthetase, 391 4-Methoxybenzoate demethylase, 545, 550, 551 4-Methoxybenzoate monooxygenase, 548, 553 2-Metnylbutyryl CoA, 632 a-Methylbutyryl-CoA dehydrogenase, 600 Methylenecyclopropyl acyl-CoA compounds, 628, see also specific types Methylenecyclopropylglycine, 638 3-Methylflavin, 12 3-Methyl-FMN binding, 12 3-Methylhypoxanthine, 59 JV-Methylnicotinamide, 58, 61 Michaelis constants, 15 Microheterogeneity, 85 Microsomal glutathione-protein disulfide oxidoreductase, 155

Microsomal protein disulfide isomerase, 166 Milk xanthine oxidase, 567 Mitochondria, 249 Mitomycin-C, 165 Molecular biology of fumarate reductase, 279—284 of methanol oxidase, 109 of succinate dehydrogenase, 279—284 of xanthine dehydrogenase, 57—58 Molybdenum center, 41—51 Molybdenum enzymes, 22, see also specific types Molybdenum hydroxylases, 22, 49, see also specific types Monoalkylated glutathione reductase (EHR), 143, 144, 147, 155 Monoclonal antibodies, 325 Monooxygenases, 469, 544, see also specific types Morris hepatoma, 18 Multidentate ligands, 179, see also specific types Mutagenesis acetolactate synthase and, 533 acyl-CoA dehydrogenases and, 646 D-amino acid oxidases and, 82 flavodoxins and, 402 fumarate reductase and, 246 lipoamide dehydrogenase and, 152—153 luciferase and, 502—503, 507—510 mechanistic effects and, 484 point, 641, 644 site-directed, 402 succinate dehyddrogenase and, 246 Myristic aldehyde, 481

N Naphthalene dioxygenase, 544, 546, 548, 553, 554 Nascent thiols, 128, 142—145, 155—156, 174—175, see also specific types Nemst equation, 31 Nifurtimox, 189 Nitrate assimilation, 320 Nitrate reductase, 319—328 active-site studies of, 324—325 amino acid sequences of, 323—324 assay, 43 chemical modification of, 324 effectors and, 327 functional domains of, 322—323 functional size of, 321—322 immunological properties of, 325 kinetics of, 326 oxidation-reduction potentials for, 327 partial activities of, 320—321 prosthetic groups and, 320 spectral properties of, 325—326 spectroscopy of, 325—326 structural properties of, 321—322 thermodynamics of, 327—328 5-Nitrofurans, 165 Nitrogenase, 391 Nitrogen-fixing bacteria, 310

Volume HI Nitrogen-NMR, 558, 569—574, 588 NMR, see Nuclear magnetic resonance Northern dot-blot analysis, 646 L-Norvaline, 74 Nuclear magnetic resonance (NMR), 557—589 acetolactate synthase and, 539 carbon, see Carbon-NMR Clostridial flavodoxins and, 428, 430, 446, 452, 456 flavodoxins and, 390, 400, 403, 428 fluorine, 581—583 hydrogen, 106, 558, 584—588 methanol oxidase and, 97, 106 multidimensional, 428 nitrogen, 558, 569—574, 588 oxygen, 583—584 phosphorus, see Phosphorus-NMR protein-FMN interactions and, 446 two-dimensional, 558, 586—588 xanthine oxidase and, 42, 44 Nucleotide binding domain, 305

o n-Octanal, 469, 482, 503 2-Octenoyl CoA, 602 3-Octenoyl CoA, 633 2-Octynoyl CoA, 627 Old Yellow Enzyme, 565, 573, 579, 605 Oligonucleotide probes, 471, 511 One-electron reduction of flavin, 32 Optimum pH for D-amino acid oxidases, 74, 76 for glutamate synthase, 311 for methanol oxidase, 108 for pyridoxine-5'-P oxidase, 3 Organomercury lyase, 179 Oxaloacetate, 240, 247 Oxidation, 14, 22, see also Oxidation-reduction; specific types acyl-CoA thiol ester, 300 D-amino acid oxidase, 77 dithionite-reduced dehydrogenase, 54 lipid-activated, 538 malate, 247 menaquinol, 276 methanol, 96 phosphopyridoxine, 12 of phospyridoxamine, 12 rate of, 539 p-Oxidation, 598, 599, 636, 638 w-Oxidation, 638 Oxidation-reduction equilibrium, 30 Oxidation-reduction properties, see also Oxidation; Reduction of acyl-CoA dehydrogenases, 631 of clostridial flavodoxins, 428 of flavodoxins, 398—401, 428 of lipoamide dehydrogenase, 139 of nitrate reductase, 327

667

of pyridine nucleotide-disulfide oxidoreductases, 170 of thioredoxin reductase, 170 of xanthine dehydrogenase, 57 of xanthine oxidase, 31, 34 Oxidative half-reaction, 37—41, 61 Oxidative phosphorylation, 230 Oxidative radiolysis, 36 Oxidized-semiquinone equilibrium, 448—452 Oxidoreductase, 470, 471, 544 2-Oxoglutarate amidotransferase, 310 N-(3-Oxo-hexanoyl)-homoserine lactone, 516 Oxygenase reactions, 537 Oxygen-NMR, 583—584 Oxygen reactivity, 603—604 Oxynitrilase, 538, 567

p Pantetheine, 601 Papain, 161 PAPS pathway, 334, 336 PCR, see Polymerase chain reaction PDR, see Phthalate dioxygenase reductase 2,3-Pentadienoyl CoA, 627 3,4-Pentadienoyl CoA, 627—628 2,4-Pentadione, 14 Pent-2-enoyl CoA, 609 3-Pentynoyl pantetheine, 626 Peptides, 238—239, 266—270, see also Polypeptides; specific types Peracetate, 538 Peroxisomes, 70, 96, 603 Persulfide of Coenzyme A, 610, 612—614, 646 Phenazine ethosulphate (PES), 603 Phenazine methosulfate (PMS), 603 Phenylalanine, 16 Phenylglycoxal, 14 Phenylmercuric acetate (PMA), 610, 624 Phenylpropionyl CoA, 607, 630 pH-jump method, 35 pH optimum, see Optimum pH Phosphate binding, 438, 457—458 Phosphates, 560, see also specific types Phosphoinositide-specific phospholipase C, 166 Phospholipase C, 166 Phaphopyridoxamine, 12 fc/s-Phosphopyridoxal, 11 Phosphopyridoxaloxime, 10, 11 Phosphopyridoxine, 12 Phosphorus-NMR, 558, 560—569 chemical shifts and, 563—564 FAD-dependent enzymes and, 565—569 FMN-dependent enzymes and, 560—565 Phosphorylation, 2, 230 Photoaffinity labeling, 630 Photolysis, 33, 35,407 Photooxidation, 636 Photoreduction of thioredoxin reductase, 174 Photosynthesis, 276 Phthalate dioxygenase, 544

668

Chemistry and Biochemistry of Flavoenzymes

Phthalate dioxygenase reductase (PDR), 543—554 biological role of, 544—545 electron transfer and, 545—548, 551—552 isolation of, 544 kinetics of, 544, 552—553 redox properties of, 550 spectral properties of, 548—551 spectroscopy of, 548—551 structural properties of, 544, 548 X-ray analysis of, 544, 551 Physical properties, see Physicochemical properties Physicochemical properties of aldehyde oxidase, 59—60 of D-amino acid oxidases, 71—76 of L-amino acid oxidases, 84—86 of anchor peptides, 266 of APS reductase, 343 of glutamate synthase, 314—316 of lipoamide dehydrogenase, 137—138 of xanthine dehydrogenase, 52—53 of xanthine oxidase, 23—28 Physiological roles, 84, 88 Ping-pong kinetics, 123, 194, 311 PL, see Pyridoxal PLP, see Pyridoxal-P PM, see Pyridoxamine PMA, see Phenylmercuric acetate PMP, see Pyridoxamine 5'-P PMS, see Phenazine methosulfate PN, see Pyridoxine PNP, see Pyridoxine-5'-P Point mutations, 641, 644 Polarography, 3 Polymerase chain reaction (PCR), 236, 646 Polypeptides, 215—218, 238—239, see also Peptides; specific types Potassium bromide, 6 Potentiometry, 31 Prenatal diagnosis, 640 Presteady-state kinetics of glutathione reductase, 156—157 of lipoamide dehydrogenase, 139—141 of mercuric ion reductase, 179 of thioredoxin reductase, 172 Primary structure, 618—621 L-Proline, 74 Prolyl 4-hydroxylase, 166 a-Propio-a-hydroxybutyrate, 533 Propionyl CoA, 602, 628—630 Proteases, 494, 500—502, see also specific types Protein-bound phosphates, 560 Protein-protein complex, 415—417 Proteins, 532, see also specific types acyl carrier, 599 apo-riboflavin-binding, 585 conversion of to semiquinone, 398 cyclic AMP, 475 electron transfer, see Electron transfer protein flavin-binding, 9 flavin interactions with, 400—406

flavin mononucleotide interaction with, 437_440, 446 fluorescence of, 10 lumazine, 512—513 riboflavin-binding, 573, 580, 581 solvent interaction with, 436—437 yellow fluorescent, 511—512 Proteolysis, 322, 488, 494, 499—500 Pulse fluorometry, 138 Pulse radiolysis, 36, 398 Purification of D-amino acid oxidases, 71, 75 of L-amino acid oxidases, 84 of coenzyme A, 601 of flavodoxins, 395—395 of fumarate reductase, 231—238 of methanol oxidase, 98—99 of pyridoxine-5'-P oxidase, 3—6 of succinate dehydrogenase, 231—238 of xanthine dehydrogenase, 52 Putidamonoxin, 546 Putidamonoxin oxido-reductase, 553 Pyocyanine, 603, 631 Pyrazon dioxygenase, 545, 546 Pyridine, 168, 558 Pyridine nucleotide, 158, 364 Pyridine nucleotide-disulfide oxidoreductases, 189—195, see also Glutathione reductase; Lipoamide dehydrogenase; Mercuric ion reductase; Thioredoxin reductase; specific types active-site studies of, 169—171 amino acid sequence homology in, 127—134 comparisons of, 123—127, 168—174 defined, 123 kinetics of, 123 mechanistic similarities among, 123—127 oxidation-reduction potentials of, 170 redox properties of, 169—171 residues in, 126—127 structural properties of, 127—135 Pyridine nucleotides, 548 6-Pyridone, 58 Pyridoxal (PL), 2, 16 Pyridoxal kinase, 17 Pyridoxal-P (PLP), 17, 136 Pyridoxal 5'-P, 2 Pyridoxal 5'-P oxidase, 17—18 Pyridoxamine, see Pyridoxine Pyridoxine, 2, 17 Pyridoxine 5'-P, 14, 16 Pyridoxine-P oxidase, 17 Pyridoxine-5'-P oxidase, 1—18 aggregation of, 5—6 anisotropy and, 12 assay of, 3 chemical modification studies and, 12—14 coenzyme for, 8, 9 colorimetry of, 3 discovery of, 2 electrophoresis of, 4

Volume III emission anisotropy and, 12 FMN-binding site and, 7—12 FMN-dependent, 2, 6 kinetics of, 14—-16 optimum pH for, 3 polarography of, 3 purification of, 3—6 pyridoxal kinase interaction with, 17 regulation of, 17—18 resolution of, 6 sheep brain, 4 spectrophotometry of, 3 spectroscopy of, 10—12 stereospecificity of, 16—17 substrate-binding site for, 12—17 substrate specificities and, 6—7 Pyrimidine, 578 Pyrophosphatase, 336 Pyrophosphate, 33, 221, 334 Pyrrole, 558 Pyruvate, 135, 532, 538 Pyruvate dehydrogenase, 391, 585 Pyruvate-formate lyase, 394 Pyruvate oxidase, 532, 534

Q Quaternary structure, 321—322, 616—618 Quinone reductase, 241 Quinones, 31, see also specific types binding of, 540 fumarate reductase interaction with, 273—278 succinate dehydrogenase interaction with, 273—278

R Racemization reactions, 2 Radiation inactivation analysis, 322 Radiolysis, 36, 398 Raman spectrum, 254 of acyl-CoA dehydrogenases, 611 of dimethylsulfoxide reductase, 51 of xanthine oxidase, 26, 51 Randomization of label, 17 Rapid equilibrium model, 31, 34 Reactive thiols, 475, 482, 495—499, 505 Reactivity of acyl-CoA dehydrogenases, 630—636 of lipoamide dehydrogenase, 144 of mercuric ion reductase, 186—189 oxygen, 603—604 of xanthine dehydrogenase, 56 Reagents, 622—626, see also specific types Recombinant DNA technology, 468 Redox-active centers, 24, 36 Redox properties, 260 of D-amino acid oxidases, 72 of L-amino acid oxidases, 86 of APS reductase, 347 of Clostridial flavodoxins, 430, 453

669

of l-deaza-FMN flavodoxin, 453 electron transfer rate constants and, 410—411 of flavin adenine dinucleotide, 130, 135 of flavodoxins, 392—393, 401, 405, 408—411, 430 of fumarate reductase, 253 of glutathione reductase, 154 kinetics and, 408—410 of lipoamide dehydrogenase, 149 of mercuric ion reductase, 181, 185 of phthalate dioxygenase reductase, 550 of pyridine nucleotide-disulfide oxidoreductases, 169—171 of succinate dehydrogenase, 253 of thioredoxin reductase, 169—171 Red semiquinone, 614—615 Reduction, see also Oxidation-reduction of D-amino acid oxidases, 72, 75 assimilatory sulfate, 334—336 dissimilatory sulfate, 334, 336—337 dithionite, 313 of flavin, 77 of glutathione disulfide, 214 of hydroxylamine, 103 kinetics of, 451 of mercuric ion reductase, 179 of NAD, 124 sulfate, 334—337 of thioredoxin, 172 of thioredoxin reductase, 174 of xanthine oxidase, 42 Reduction potentials, 31, 33 Reductive half-reaction, 41—51 Reductive radiolysis, 36 Reoxidation, 78 Resistance to herbicides, 533 Resolution of pyridoxine-5'-P oxidase, 6 Restriction fragment length polymorphism (RFLP) analysis, 641 RFLP, see Restriction fragment length polymorphism Riboflavin, 8, 245, 403, 404, 431 Riboflavin-binding protein, 573, 580, 581 Riboflavin 3',5'-bisphosphate, 403, 564 Riboflavin deficiency, 2 Riboflavin kinase, 98 Riboflavin 5'-methylphosphate, 8 Riboflavin 4'-phosphate, 403 Riboflavin synthase, 98 Ribonculeotide reductase, 166 Ring bending, 456—457

s Salicylate hydroxylase, 580 Schiff base, 102 SDH, see Succinate dehydrogenase Sedimentation coefficients, 85 Semiquinone, 24, 31, 32, 35, 60, 156, 174 acyl-CoA dehydrogenases and, 604—607 blue, 314, 614—615

670

Chemistry and Biochemistry of Flavoenzymes

charge-transfer complexes and, 604—607 Clostridial flavodoxins and, 442—444 electron transfer and, 415, 417 fumarate reductase and, 273 glutamate synthase and, 314 green, 608—614 ionic strength and, 410 ionization of, 398 methanol oxidase and, 103—104 oxidized, 448—452 protein converted to, 398 red, 614—615 stabilization of, 400 succinate dehydrogenase and, 273 Semiquinone-reduced equilibrium, 452—457 Serine proteases, 226, see also specific types Sheep brain pyridoxine-5'-P oxidase, 4 Short-chain acyl-CoA dehydrogenases, 605, 628, 632, 644 Site-directed mutagenesis, 402 SKIE, see Solvent kinetic isotope effects SOD, see Superoxide dismutase Sodium borohydride, 75 Solvent kinetic isotope effects (SKIE), 163 Solvents, 373, 436—437, see also specific types Sources of acetolactate synthase, 532—533 of L-amino acid oxidases, 84 of glutamate synthase, 310 of glutathione reductase, 153 of methanol oxidase, 98—99 Spectral properties, see also specific types of acyl-CoA dehydrogenases, see under AcylCoA dehydrogenases of aldehyde oxidase, 59 of APS reductase, 342—346 of dimethylsulfoxide reductase, 51 of glutamate synthase, 313—314 of glutathione reductase, 156, 161—162 of lipoamide dehydrogenase, 138—139, 147—153 of methanol oxidase, 100—102, 106, 114 of nitrate reductase, 325—326 of phthalate dioxygenase reductase, 548—551 of xanthine dehydrogenases, 52 of xanthine oxidase, 26, 51 Spectrometry, 639, see also Spectroscopy; specific types Spectrophotometry, 3, 402, 407, see also Spectroscopy; specific types Spectroscopy, see also specific types of APS reductase, 342—346 circular dichroism, 325 electron paramagnetic resonance, see Electron paramagnetic resonance (EPR) magnetic circular dichroism, 49, 250, 254, 257, 258, 260 of nitrate reductase, 325—326 nuclear magnetic resonance, see Nuclear magnetic resonance (NMR) of phthalate dioxygenase reductase, 548—551

of pyridoxine-5'-P oxidase, 10—12 of xanthine oxidase, 24 X-ray absorption, 45, 46, 47, 51 Spinach ferredoxin, 26 Spinach glycolate oxidase, see Glycolate oxidase Spin echo method, 103 SRB, see Sulfate reducing bacteria Steady-state kinetics, 194 of glutamate synthase, 311 of glutathione reductase, 157—159 of lipoamide dehydrogenase, 141 of mercuric ion reductase, 180 of thioredoxin reductase, 175 of xanthine oxidase, 24 Stellacyanin, 411 Stereochemistry, 357—364, 538, 631—632 Stereospecificity, 16—17, 70, 78, 359 Steric effects on electron transfer rates, 411—415 Stoichiometry, 613 of aldehyde oxidase, 60 of flavin adenosine dinucleotide, 107 of luciferase enzymology, 478—479 of molybdenum hydroxylases, 22 of superoxide, 39, 61 Stopped-flow spectrophotometry, 402, 407 S-transferases, 155, see also specific types Structural properties of acetolactate synthase, 532—533 of acyl-CoA dehydrogenases, see under AcylCoA dehydrogenases of D-amino acid oxidases, 78—83 of APS reductase, 342—346 of Clostridial flavodoxins, 430—435 crystal, see Crystals of 1-deaza-FMN-flavodoxin, 453 of flavodoxins, 396—398, 447—448 of fumarate reductase, 279—281 of glutathione reductase, see under Glutathione reductase of glycolate oxidase, see under Glycolate oxidase of lipoamide dehydrogenase, 143 of luciferase, see under Luciferase of lux regulon, 515—516 of medium-chain acyl-CoA dehydrogenase, 301—304 of methanol oxidase, 99—100 of nitrate reductase, 321—322 of phthalate dioxygenase reductase, 544, 548 primary, 618—621 of protein-protein complex, 415—417 of pyridine nucleotide-disulfide oxidoreductases, 127—135 quaternary, 321—322, 616—618 secondary, 370—373, 435 of succinate dehyddrogenase, 279—281 tertiary, 621—622 of water, 373—374 X-ray, see X-ray analysis Structure-function studies of flavodoxins, 430—433 Substrate-binding site, 12—17, 246—249 Substrate pocket, 376—377

Volume HI Substrate specificities acyl-CoA dehydrogenases and, 599—602 L-amino acid oxidases an, 86 D-amino acid oxidases and, 73, 76 glutamate synthase and, 311 glycolate oxidase and, 368—369 pyridoxine-5'-P oxidase and, 6—7 Succinate, 246 Succinate dehydrogenase, 229—284 active-site studies of, 248, 249 assays of, 239—241 assembly of complex of, 270—273 carboxanilides and, 278—279 catalytic properties of, 239—242 flavin moiety of, 243—246 gene expression in, 281—284 gene organization of, 279—281 genetics of, 279—284 iron-sulfur clusters and, 249—266 isolation of, 231—232 membrane-bound types of, 230—232 molecular biology of, 279—284 paniculate forms of, 241—242 purification of, 231—238 quinone interaction with, 273—278 redox properties of, 253 soluble forms of, 241—242 structural properties of, 279—281 substrate-binding site and, 246—249 subunit processing and, 270 thenoyltrifluoroacetone and, 241, 278—279 Succinate-ubiquinone oxidoreductase, 230, 234—235 Succinyl-CoA, 135 Sudden infant death syndrome, 640 Suicide inactivation, 627 Suicide inhibition, 628 Sulfate-reducing bacteria (SRB), 334, 339—341, 351 Sulfate reduction, 334—337 Sulfhydryl oxidase, 192 Sulfite, 399, 615 Sulfite-L-amino acid oxidase complex, 85 Sulfite oxidase, 323 4-Sulfobenzoate dioxygenase, 546 Sulfometuron methyl, 540 Sulfonanilide, 540 Sulfonylurea, 540 Sulfotransferases, 334—336 Sulfur cycle, 334 Sulfur-iron, see Iron-sulfur Sulfur-sulfur stretching frequency, 611 Sulindac, 58 Superoxide, 38, 39, 41, 61 Superoxide dismutase, 61

T Tartronate-semialdehyde synthase, 538 Taurine, 609 TCA, see Trichloracetic acid

671

Temperature-jump methods, 402 Tertiary structural properties, 621—622 Tetrahydrofolate, 136 Tetrahydro-thiamin pyrophosphate, 540 Tetranitromethane, 626 Tetrazolium dyes, 603 Thenoyltrifluoroacetone (TTFA), 241, 273, 274, 278—279 Thermodynamics, 417 Clostridial flavodoxins and, 428 of nitrate reductase, 327—328 oxidation-reduction, 57 of xanthine oxidase, 28—37 Thiamin pyrophosphate, 136, 532, 533, 540, 541 Thiazoles, 533 Thioesterase, 603 Thioesters, 300, 600—602 Thioether, 43, 601, 602 4-Thio-FMN, 406 2-Thio-FMN-flavodoxin, 405 4-Thio-FMN-flavodoxin, 405 Thiolate, 43, 124 Thiols, 43, see also specific types active-site, 180—185, 249 acyl-CoA dehydrogenases and, 600—602 auxiliary, 181 — 185 chemical modification of, 503 disulfide interchange with, 123, 163, 168 electron transfer and, 186—189 interchange (distal), 124 nascent, 128, 142—145, 155—156, 174—175 reactive, 475, 482, 495—499, 505 sulfotransferases and, 334—336 Thioltransferase, 166 Thionitrobenzoate, 403 Thiophenol, 43 Thiopropyl-Sepharose, 610 Thioredoxin, 155, 165—167, 172 Thioredoxin reductase, 165—178, 186 active-site studies of, 169—171 catalytic properties of, 175—178 comparison with other family members, 168—174 crystals of, 178 distribution of, 165—168 glutavhione reductase compared with, 168 kinetics of, 172 metabolic function of, 165—168 oxidation-reduction potentials of, 170 photoreduction of, 174 presteady-state kinetics of, 172 redox properties of, 169—171 reduction of, 174 steady-state kinetics of, 175 Threonines, 382 Time-resolved fluorescence, 406 Toluene dioxygenase, 544—546, 548, 551, 553, 554 Toluene 2,3-dioxygenase, 546 S-Transferases, 155, see also specific types

672

Chemistry and Biochemistry of Flavoenzymes

Transhydrogenases, 143, 155, 165, see also specific types Transition state analogues, 312 Triazolo pyrimidine, 540 Trichloracetic acid (TCA), 610, 616 Trimethylamine dehydrogenase, 363, 380 2,4,6-Trinitro-benzene-sulfonate, 165 Trithionate reductase, 391 Tritium, 641 Trypanothione, 155, 189 Trypanothione reductase, 127—134, 155, 189—191 Tryptophan, 2, 9, 10 TTFA, see Thenoyltrifluoroacetone Tumors, 17—18, see also Cancer; specific types Tyrosine, 9, 52, 626

u Ubiquinone, 236 Ubiquinone-40, 538 Ubiquinone-O, 540 Ultraviolet/visible spectrum, 52, 59, 342

Valine, 532 Vanillate demethylase, 545 Vanillate demethylate oxido-reductase, 546—548 Vinylglycine, 88 Vitamin B6, 2

w Water structure, 373—374 X

Xanthine dehydrogenase, 22, 51—58 dithionite-reduced, 54 electron transfer within, 33 genetics of, 57—58 kinetics of, 53—55

molecular genetics of, 57-—58 oxidation-reduction properties of, 57 physicochemical properties of, 52—53 purification of, 52 reactivity of, 56 xanthine oxidase interconversion, 55—57 Xanthine oxidase, 22—51 electron transfer and, 28—37 flavin site and, 37—41 kinetics of, 24, 28—37 milk, 567 oxidative half-reaction of, 37—41 physicochemical properties of, 23—28 Raman spectrum of, 26, 51 reduction of, 42 reductive half-reaction of, 41—51 spectroscopy of, 24 steady-state kinetics of, 24 thermodynamics of, 28—37 xanthine dehydrogenase interconversion, 55—57 X-ray absorption fine structure (EXAFS), 45, 51, 545 X-ray absorption spectroscopy, 45—47, 51 X-ray analysis, 363, see also specific types of acyl-CoA dehydrogenases, 621 of D-amino acid oxidases, 79 of Clostridial flavodoxins, 430, 433—448 polypeptide fold and, 435—437 structure determination in, 433—435 dispersive, 42 of lipoamide dehydrogenase, 143 of medium-chain acyl-CoA dehydrogenase, 300 of methanol oxidase, 100 of phthalate dioxygenase reductase, 544, 551 of pyridine nucleotide-disulfide oxidoreductases, 134—135 of thioredoxin reductase, 178 X-ray crystallography, 300, 363, 433—435, 586, 588, 621, see also X-ray analysis

Y Yellow fluorescent protein (YFP), 511—512